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Mouse Cell Culture: Methods and Protocols

Methods in
Molecular Biology 1940
Ivan Bertoncello Editor
Mouse
Cell Culture
Methods and Protocols
METHODS IN MOLECULAR BIOLOGY
Series Editor
John M. Walker
School of Life and Medical Sciences
University of Hertfordshire
Hatfield, Hertfordshire, AL10 9AB, UK
For further volumes:
http://www.springer.com/series/7651
Mouse Cell Culture
Methods and Protocols
Edited by
Ivan Bertoncello
Lung Health Research Centre, Department of Pharmacology and Therapeutics, University of
Melbourne, Melbourne, Victoria, Australia
Editor
Ivan Bertoncello
Lung Health Research Centre
Department of Pharmacology
and Therapeutics
University of Melbourne
Melbourne, Victoria, Australia
ISSN 1064-3745
ISSN 1940-6029 (electronic)
Methods in Molecular Biology
ISBN 978-1-4939-9085-6
ISBN 978-1-4939-9086-3 (eBook)
https://doi.org/10.1007/978-1-4939-9086-3
Library of Congress Control Number: 2019930653
© Springer Science+Business Media, LLC, part of Springer Nature 2019
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Preface
Mouse models have long underpinned discovery in the biomedical sciences and the development of enabling technologies for the systematic analysis of cellular and molecular
mechanisms of organ development, regeneration, and repair. In particular, mouse cell
culture systems continue to play an important role in identifying key factors, genes, and
pathways regulating cell lineage commitment and differentiation and validating potential
cellular and molecular targets which could be exploited to develop novel therapies for
intractable diseases.
This compendium describes recently devised and refined best-practice cell culture protocols for the maintenance, propagation, manipulation, and analysis of primary explanted
cells from various mouse organ systems commonly used in current research applications.
Each chapter provides a step-by-step description of cell culture methodologies for specific
mouse cell types and lineages, highlighting caveats and commonly encountered pitfalls.
These protocols are preceded by two introductory chapters that review the applicability of
mouse models as a discovery tool and describe factors and variables that influence cell
culture endpoints and need to be considered and controlled in order to achieve optimal
results.
In conclusion, I would like to acknowledge and thank the many authors who have
enthusiastically contributed their protocols to this volume. I also thank John Walker (series
editor) for his invitation to edit the volume and for his advice and assistance in developing
and preparing the volume for publication.
Melbourne, Victoria, Australia
Ivan Bertoncello
v
Contents
Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Contributors. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
PART I
PRACTICAL CONSIDERATIONS
1 The Applicability of Mouse Models to the Study of Human Disease. . . . . . . . . . .
Kristina Rydell-Törm€
a nen and Jill R. Johnson
2 Optimizing the Cell Culture Microenvironment . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Ivan Bertoncello
PART II
v
xi
3
23
METHODS AND PROTOCOLS
3 Propagation and Maintenance of Mouse Embryonic
Stem Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Jacob M. Paynter, Joseph Chen, Xiaodong Liu, and Christian M. Nefzger
4 Production of High-Titer Lentiviral Particles for Stable Genetic
Modification of Mammalian Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Michael R. Larcombe, Jan Manent, Joseph Chen, Ketan Mishra,
Xiaodong Liu, and Christian M. Nefzger
5 Generation of Mouse-Induced Pluripotent Stem Cells by Lentiviral
Transduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Xiaodong Liu, Joseph Chen, Jaber Firas, Jacob M. Paynter,
Christian M. Nefzger, and Jose M. Polo
6 Gene Editing of Mouse Embryonic and Epiblast Stem Cells. . . . . . . . . . . . . . . . . .
Tennille Sibbritt, Pierre Osteil, Xiaochen Fan, Jane Sun,
Nazmus Salehin, Hilary Knowles, Joanne Shen, and Patrick P. L. Tam
7 Identification of Circulating Endothelial Colony-Forming Cells
from Murine Embryonic Peripheral Blood . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Yang Lin, Chang-Hyun Gil, and Mervin C. Yoder
8 Imaging and Analysis of Mouse Embryonic Whole Lung, Isolated Tissue,
and Lineage-Labelled Cell Culture . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Matthew Jones and Saverio Bellusci
9 Mouse Hematopoietic Stem Cell Modification and Labelling
by Transduction and Tracking Posttransplantation . . . . . . . . . . . . . . . . . . . . . . . . . .
Benjamin Cao, Songhui Li, Claire Pritchard, Brenda Williams,
and Susan K. Nilsson
10 Genetic Manipulation and Selection of Mouse Mesenchymal Stem
Cells for Delivery of Therapeutic Factors In Vivo . . . . . . . . . . . . . . . . . . . . . . . . . . .
Donald S. Sakaguchi
11 Isolation and Culture of Primary Mouse Middle Ear Epithelial Cells . . . . . . . . . .
Apoorva Mulay, Khondoker Akram, Lynne Bingle, and Colin D. Bingle
vii
33
47
63
77
97
109
129
143
157
viii
12
13
14
15
16
17
18
19
20
21
Contents
Isolation and Propagation of Lacrimal Gland Putative Epithelial
Progenitor Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Helen P. Makarenkova and Robyn Meech
Organotypic Culture of Adult Mouse Retina . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Brigitte Müller
Langendorff-Free Isolation and Propagation of Adult Mouse
Cardiomyocytes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Matthew Ackers-Johnson and Roger S. Foo
Isolation, Culture, and Characterization of Primary Mouse
Epidermal Keratinocytes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Ling-Juan Zhang
Isolation and Propagation of Mammary Epithelial Stem
and Progenitor Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Julie M. Sheridan and Jane E. Visvader
An Organoid Assay for Long-Term Maintenance and Propagation
of Mouse Prostate Luminal Epithelial Progenitors and Cancer Cells. . . . . . . . . . .
Yu Shu and Chee Wai Chua
Isolation, Purification, and Culture of Mouse Pancreatic Islets
of Langerhans . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Youakim Saliba and Nassim Farès
Identification and In Vitro Expansion of Adult Hepatocyte
Progenitors from Chronically Injured Livers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Naoki Tanimizu
The Preparation of Decellularized Mouse Lung Matrix Scaffolds
for Analysis of Lung Regenerative Cell Potential . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Deniz A. Bölükbas, Martina M. De Santis, Hani N. Alsafadi,
Ali Doryab, and Darcy E. Wagner
Mouse Lung Tissue Slice Culture . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Xinhui Wu, Eline M. van Dijk, I. Sophie T. Bos,
Loes E. M. Kistemaker, and Reinoud Gosens
Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
169
181
193
205
217
231
255
267
275
297
313
Contributors
MATTHEW ACKERS-JOHNSON Cardiovascular Research Institute, Centre for Translational
Medicine MD6, National University Health System, Singapore, Singapore; Genome
Institute of Singapore, Singapore, Singapore
KHONDOKER AKRAM Academic Unit of Respiratory Medicine, Department of Infection,
Immunity and Cardiovascular Disease, University of Sheffield, Sheffield, UK
HANI N. ALSAFADI Department of Experimental Medical Sciences, Faculty of Medicine,
Lund University, Lund, Sweden; Wallenberg Centre for Molecular Medicine, Faculty of
Medicine, Lund University, Lund, Sweden; Stem Cell Centre, Lund University, Lund,
Sweden
SAVERIO BELLUSCI Faculty of Medicine, Excellence Cluster Cardio-Pulmonary System
(ECCPS), University of Giessen, Giessen, Germany
IVAN BERTONCELLO Lung Health Research Centre, Department of Pharmacology and
Therapeutics, University of Melbourne, Melbourne, Victoria, Australia
COLIN D. BINGLE Academic Unit of Respiratory Medicine, Department of Infection,
Immunity and Cardiovascular Disease, University of Sheffield, Sheffield, UK
LYNNE BINGLE Oral and Maxillofacial Pathology, Department of Clinical Dentistry,
University of Sheffield, Sheffield, UK
DENIZ A. BÖLÜKBAS Department of Experimental Medical Sciences, Faculty of Medicine,
Lund University, Lund, Sweden; Wallenberg Centre for Molecular Medicine, Faculty of
Medicine, Lund University, Lund, Sweden; Stem Cell Centre, Lund University, Lund,
Sweden
I. SOPHIE T. BOS Faculty of Science and Engineering, Department of Molecular
Pharmacology, University of Groningen, Groningen, The Netherlands; Groningen Research
Institute for Asthma and COPD, University Medical Center Groningen, University of
Groningen, Groningen, The Netherlands
BENJAMIN CAO Biomedical Manufacturing, CSIRO Clayton, Clayton, VIC, Australia;
Australian Regenerative Medicine Institute, Monash University, Clayton, VIC, Australia
JOSEPH CHEN Department of Anatomy and Developmental Biology, Monash University,
Clayton, VIC, Australia; Development and Stem Cells Program, Monash Biomedicine
Discovery Institute, Clayton, VIC, Australia; Australian Regenerative Medicine Institute,
Monash University, Clayton, VIC, Australia
CHEE WAI CHUA State Key Laboratory of Oncogenes and Related Genes, Renji-Med X
Clinical Stem Cell Research Center, Department of Urology, Ren Ji Hospital, School of
Medicine, Shanghai Jiao Tong University, Shanghai, China
MARTINA M. DE SANTIS Department of Experimental Medical Sciences, Faculty of
Medicine, Lund University, Lund, Sweden; Wallenberg Centre for Molecular Medicine,
Faculty of Medicine, Lund University, Lund, Sweden; Stem Cell Centre, Lund University,
Lund, Sweden
ALI DORYAB Helmholtz Zentrum München, Member of the German Center for Lung
Research (DZL), Institute of Lung Biology and Disease, Neuherberg, Germany
XIAOCHEN FAN Children’s Medical Research Institute, The University of Sydney, Westmead,
NSW, Australia
ix
x
Contributors
NASSIM FARÈS Laboratoire de Recherche en Physiologie et Physiopathologie LRPP, Pôle
Technologie Santé, Faculté de Médecine, Université Saint Joseph, Beirut, Lebanon
JABER FIRAS Department of Anatomy and Developmental Biology, Monash University,
Clayton, VIC, Australia; Development and Stem Cells Program, Monash Biomedicine
Discovery Institute, Clayton, VIC, Australia; Australian Regenerative Medicine Institute,
Monash University, Clayton, VIC, Australia
ROGER S. FOO Cardiovascular Research Institute, Centre for Translational Medicine
MD6, National University Health System, Singapore, Singapore; Genome Institute of
Singapore, Singapore, Singapore
CHANG-HYUN GIL Department of Pediatrics, Herman B. Wells Center for Pediatric
Research, Indiana University School of Medicine, Indianapolis, IN, USA
REINOUD GOSENS Faculty of Science and Engineering, Department of Molecular
Pharmacology, University of Groningen, Groningen, The Netherlands; Groningen Research
Institute for Asthma and COPD, University Medical Center Groningen, University of
Groningen, Groningen, The Netherlands
JILL R. JOHNSON School of Life and Health Sciences, Aston University, Birmingham, UK
MATTHEW JONES Faculty of Medicine, Excellence Cluster Cardio-Pulmonary System
(ECCPS), University of Giessen, Giessen, Germany
LOES E. M. KISTEMAKER Faculty of Science and Engineering, Department of Molecular
Pharmacology, University of Groningen, Groningen, The Netherlands; Groningen Research
Institute for Asthma and COPD, University Medical Center Groningen, University of
Groningen, Groningen, The Netherlands
HILARY KNOWLES Children’s Medical Research Institute, The University of Sydney,
Westmead, NSW, Australia
MICHAEL R. LARCOMBE Department of Anatomy and Developmental Biology, Monash
University, Clayton, VIC, Australia; Development and Stem Cells Program, Monash
Biomedicine Discovery Institute, Clayton, VIC, Australia; Australian Regenerative
Medicine Institute, Monash University, Clayton, VIC, Australia
SONGHUI LI Biomedical Manufacturing, CSIRO Clayton, Clayton, VIC, Australia;
Australian Regenerative Medicine Institute, Monash University, Clayton, VIC, Australia
YANG LIN Department of Pediatrics, Herman B. Wells Center for Pediatric Research,
Indiana University School of Medicine, Indianapolis, IN, USA; Department of
Biochemistry and Molecular Biology, Indiana University School of Medicine, Indianapolis,
IN, USA
XIAODONG LIU Department of Anatomy and Developmental Biology, Monash University,
Clayton, VIC, Australia; Development and Stem Cells Program, Monash Biomedicine
Discovery Institute, Clayton, VIC, Australia; Australian Regenerative Medicine Institute,
Monash University, Clayton, VIC, Australia
HELEN P. MAKARENKOVA Department of Molecular Medicine, The Scripps Research
Institute, La Jolla, CA, USA
JAN MANENT Department of Anatomy and Developmental Biology, Monash University,
Clayton, VIC, Australia; Development and Stem Cells Program, Monash Biomedicine
Discovery Institute, Clayton, VIC, Australia; Australian Regenerative Medicine Institute,
Monash University, Clayton, VIC, Australia
ROBYN MEECH Discipline of Clinical Pharmacology, College of Medicine and Public Health,
Flinders University, Bedford Park, SA, Australia
KETAN MISHRA Department of Anatomy and Developmental Biology, Monash University,
Clayton, VIC, Australia; Development and Stem Cells Program, Monash Biomedicine
Contributors
xi
Discovery Institute, Clayton, VIC, Australia; Australian Regenerative Medicine Institute,
Monash University, Clayton, VIC, Australia
APOORVA MULAY Academic Unit of Respiratory Medicine, Department of Infection,
Immunity and Cardiovascular Disease, University of Sheffield, Sheffield, UK
BRIGITTE MÜLLER Department of Ophthalmology, Justus-Liebig-University Gießen, Gießen,
Germany
CHRISTIAN M. NEFZGER Department of Anatomy and Developmental Biology, Monash
University, Clayton, VIC, Australia; Development and Stem Cells Program, Monash
Biomedicine Discovery Institute, Clayton, VIC, Australia; Australian Regenerative
Medicine Institute, Monash University, Clayton, VIC, Australia; Institute for Molecular
Bioscience, The University of Queensland, St Lucia, QLD, Australia
SUSAN K. NILSSON Biomedical Manufacturing, CSIRO Clayton, Clayton, VIC, Australia;
Australian Regenerative Medicine Institute, Monash University, Clayton, VIC, Australia
PIERRE OSTEIL Children’s Medical Research Institute, The University of Sydney, Westmead,
NSW, Australia; School of Medical Sciences, The University of Sydney, Camperdown, NSW,
Australia
JACOB M. PAYNTER Department of Anatomy and Developmental Biology, Monash
University, Clayton, VIC, Australia; Development and Stem Cells Program, Monash
Biomedicine Discovery Institute, Clayton, VIC, Australia; Australian Regenerative
Medicine Institute, Monash University, Clayton, VIC, Australia
JOSE M. POLO Department of Anatomy and Developmental Biology, Monash University,
Clayton, VIC, Australia; Development and Stem Cells Program, Monash Biomedicine
Discovery Institute, Clayton, VIC, Australia; Australian Regenerative Medicine Institute,
Monash University, Clayton, VIC, Australia
CLAIRE PRITCHARD Biomedical Manufacturing, CSIRO Clayton, Clayton, VIC, Australia;
Australian Regenerative Medicine Institute, Monash University, Clayton, VIC, Australia
€
KRISTINA RYDELL-TÖRMANEN
Lung Biology Group, Department of Experimental Medical
Science, Lund University, Lund, Sweden
DONALD S. SAKAGUCHI Department of Genetics, Development and Cell Biology, Iowa State
University, Ames, IA, USA; Neuroscience Program, Iowa State University, Ames, IA, USA
NAZMUS SALEHIN Children’s Medical Research Institute, The University of Sydney,
Westmead, NSW, Australia
YOUAKIM SALIBA Laboratoire de Recherche en Physiologie et Physiopathologie LRPP, Pôle
Technologie Santé, Faculté de Médecine, Université Saint Joseph, Beirut, Lebanon
JOANNE SHEN Children’s Medical Research Institute, The University of Sydney, Westmead,
NSW, Australia
JULIE M. SHERIDAN Molecular Genetics of Cancer Division, Walter and Eliza Hall Institute
of Medical Research, Parkville, VIC, Australia; Department of Medical Biology, University
of Melbourne, Melbourne, VIC, Australia
YU SHU State Key Laboratory of Oncogenes and Related Genes, Renji-Med X Clinical Stem
Cell Research Center, Department of Urology, Ren Ji Hospital, School of Medicine,
Shanghai Jiao Tong University, Shanghai, China
TENNILLE SIBBRITT Children’s Medical Research Institute, The University of Sydney,
Westmead, NSW, Australia; School of Medical Sciences, The University of Sydney,
Camperdown, NSW, Australia
JANE SUN Children’s Medical Research Institute, The University of Sydney, Westmead,
NSW, Australia
xii
Contributors
PATRICK P. L. TAM Children’s Medical Research Institute, The University of Sydney,
Westmead, NSW, Australia; School of Medical Sciences, The University of Sydney,
Camperdown, NSW, Australia
NAOKI TANIMIZU Department of Tissue Development and Regeneration, Research Institute
for Frontier Medicine, Sapporo Medical University School of Medicine, Sapporo, Japan
ELINE M. VAN DIJK Faculty of Science and Engineering, Department of Molecular
Pharmacology, University of Groningen, Groningen, The Netherlands; Groningen Research
Institute for Asthma and COPD, University Medical Center Groningen, University of
Groningen, Groningen, The Netherlands
JANE E. VISVADER Department of Medical Biology, University of Melbourne, Melbourne,
VIC, Australia; Stem Cells and Cancer Division, Walter and Eliza Hall Institute of
Medical Research, Parkville, VIC, Australia
DARCY E. WAGNER Department of Experimental Medical Sciences, Faculty of Medicine,
Lund University, Lund, Sweden; Wallenberg Centre for Molecular Medicine, Faculty of
Medicine, Lund University, Lund, Sweden; Stem Cell Centre, Lund University, Lund,
Sweden
BRENDA WILLIAMS Biomedical Manufacturing, CSIRO Clayton, Clayton, VIC, Australia;
Australian Regenerative Medicine Institute, Monash University, Clayton, VIC, Australia
XINHUI WU Faculty of Science and Engineering, Department of Molecular Pharmacology,
University of Groningen, Groningen, The Netherlands; Groningen Research Institute for
Asthma and COPD, University Medical Center Groningen, University of Groningen,
Groningen, The Netherlands
MERVIN C. YODER Department of Pediatrics, Herman B. Wells Center for Pediatric
Research, Indiana University School of Medicine, Indianapolis, IN, USA; Department of
Biochemistry and Molecular Biology, Indiana University School of Medicine, Indianapolis,
IN, USA
LING-JUAN ZHANG School of Pharmaceutical Sciences, Xiamen University, Xiamen, China;
Department of Dermatology, School of Medicine, University of California San Diego, La
Jolla, CA, USA
Part I
Practical Considerations
Chapter 1
The Applicability of Mouse Models to the Study of Human
Disease
Kristina Rydell-Törm€anen and Jill R. Johnson
Abstract
The laboratory mouse Mus musculus has long been used as a model organism to test hypotheses and
treatments related to understanding the mechanisms of disease in humans; however, for these experiments
to be relevant, it is important to know the complex ways in which mice are similar to humans and, crucially,
the ways in which they differ. In this chapter, an in-depth analysis of these similarities and differences is
provided to allow researchers to use mouse models of human disease and primary cells derived from these
animal models under the most appropriate and meaningful conditions.
Although there are considerable differences between mice and humans, particularly regarding genetics,
physiology, and immunology, a more thorough understanding of these differences and their effects on the
function of the whole organism will provide deeper insights into relevant disease mechanisms and potential
drug targets for further clinical investigation. Using specific examples of mouse models of human lung
disease, i.e., asthma, chronic obstructive pulmonary disease, and pulmonary fibrosis, this chapter explores
the most salient features of mouse models of human disease and provides a full assessment of the advantages
and limitations of these models, focusing on the relevance of disease induction and their ability to replicate
critical features of human disease pathophysiology and response to treatment. The chapter concludes with a
discussion on the future of using mice in medical research with regard to ethical and technological
considerations.
Key words Mouse, Model, Disease, Genetics, Physiology, Immunology, Ethics
1
The Mouse: From Pest, to Pet, to Predominant Tool in Medical Research
Although the genetic lineages of mice and humans diverged around
75 million years ago, these two species have evolved to live
together, particularly since the development of agriculture. For
millennia, mice (Mus musculus) were considered to be pests due
to their propensity to ravenously consume stored foodstuff (mush
in ancient Sanskrit means “to steal” [1]) and their ability to adapt to
a wide range of environmental conditions. Since the 1700s, domesticated mice have been bred and kept as companion animals, and in
Victorian England, “fancy” mice were prized for their variations in
coat color and comportment; these mouse strains were the
Ivan Bertoncello (ed.), Mouse Cell Culture: Methods and Protocols, Methods in Molecular Biology, vol. 1940,
https://doi.org/10.1007/978-1-4939-9086-3_1, © Springer Science+Business Media, LLC, part of Springer Nature 2019
3
4
€nen and Jill R. Johnson
Kristina Rydell-Törma
forerunners to the strains used in the laboratory today. Robert
Hooke performed the first recorded inquiry-driven experiments
on mice in 1664, when he investigated the effects of changes in
air pressure on respiratory function [2]. More recently, with data
from the Human Genome Project and sequencing of the Mus
musculus genome showing remarkable genetic homology between
these species, as well as the advent of biotechnology and the development of myriad knockout and transgenic mouse strains, it is clear
why the mouse has become the most ubiquitous model organism
used to study human disease. In addition, their small size, rapid
breeding, and ease of handling are all important advantages to
scientists for practical and financial reasons. However, keeping in
mind that mice are fellow vertebrates and mammals, there are
ethical issues inherent to using these animals in medical research.
This chapter will provide an overview of the important similarities
and differences between Mus musculus and Homo sapiens and their
relevance to the use of the mouse as a model organism and provide
specific examples of the quality of mouse models used to investigate
the mechanisms, pathology, and treatment of human lung diseases.
We will then conclude with an assessment of the future of mice in
medical research considering ethical and technological advances.
As a model organism used to test hypotheses and treatments
related to human disease, it is important to understand the complex
ways in which mice are similar to humans, and crucially, the ways in
which they differ. A clear understanding of these aspects will allow
researchers to use mouse models of human disease and primary cells
derived from mice under the most appropriate and meaningful
conditions.
2
Applicability of Mouse Models to Human Disease
2.1
Genetics
In 2014, the Encyclopedia of DNA Elements (ENCODE) program
published a comparative analysis of the genomes of Homo sapiens
and Mus musculus [3], as well as an in-depth analysis of the differences in the regulatory landscape of the genomes of these species
[4]. ENCODE, a follow-up to the Human Genome Project, was
implemented by the National Human Genome Research Institute
(NHGRI) at the National Institutes of Health in order to develop a
comprehensive catalog of protein-encoding and nonproteincoding genes and the regulatory elements that control gene expression in a number of species. This was achieved using a number of
genomic approaches (e.g., RNA-seq, DNase-seq, and ChIP-seq) to
assess gene expression in over 100 mouse cell types and tissues; the
data were then compared with the human genome.
Overall, these studies showed that although gene expression is
fairly similar between mice and humans, considerable differences
were observed in the regulatory networks controlling the activity of
Mouse Models of Human Disease
5
the immune system, metabolic functions, and responses to stress,
all of which have important implications when using mice to model
human disease. In essence, mice and humans demonstrate genetic
similarity with regulatory divergence. Specifically, there is a high
degree of similarity in transcription factor networks but a great deal
of divergence in the cis-regulatory elements that control gene
transcription in the mouse and human genomes. Moreover, the
chromatin landscape in cell types of similar lineages in mouse and
human is both developmentally stable and evolutionarily conserved
[3]. Of particular relevance regarding modeling human diseases
involving the immune system, in its assessment of transcription
factor networks, the Mouse ENCODE Consortium revealed
potentially important differences in the activity of ETS1 in the
mouse and human genome. Although conserved between the two
species, divergence in ETS1 regulation may be responsible for
discrepancies in the function of the immune system in mouse and
human [4]. Certainly, the biological consequences of these differences in gene expression and regulation between human and mouse
invite further investigation.
2.2 Anatomy and
Physiology
The anatomical and physiological differences between model
organisms and humans can have profound impacts on interpreting
experimental results. Virtually every biological process under investigation in experimental studies involves at least one anatomical
structure. To aid in interpretation, many anatomy compendia
have been developed for model organisms; the most useful organize anatomical entities into hierarchies representing the structure
of the human body, e.g., the Foundational Model of Anatomy
developed by the Structural Informatics Group at the University
of Washington [5]. Although an analysis of the myriad differences
between mouse and human anatomy is beyond the scope of this
chapter, a few of the most critical issues that have an impact on the
interpretation of data from mouse experiments should be
mentioned.
The most obvious difference between mice and humans is size;
the human body is about 2500 times larger than that of the mouse.
Size influences many aspects of biology, particularly the metabolic
rate, which is correlated to body size in placental mammals through
the relationship BMR ¼ 70 mass (0.75), where BMR is the basal
metabolic rate (in kcal/day). Thus, the mouse BMR is roughly
seven times faster than that of an average-sized human [6]. This
higher BMR has effects on thermoregulation, nutrient demand,
and nutrient supply. As such, mice have greater amounts of metabolically active tissues (e.g., liver and kidney) and more extensive
deposits of brown fat [6]. Furthermore, mice more readily produce
reactive oxygen species than do humans, which is an important
consideration when modeling human diseases involving the
6
€nen and Jill R. Johnson
Kristina Rydell-Törma
induction of oxidative stress (i.e., aging, inflammation, and
neurodegeneration) [6].
The lung provides an excellent example of the similarities and
differences between human and mouse anatomy. Similar to the
human organ, the mouse lung is subdivided into lobes of lung
parenchyma containing a branching bronchial tree and is vascularized by the pulmonary circulation originating from the right ventricle. There are a number of subtle variations in this general
structure between species, i.e., the number of lobes on the right
and left, the branching pattern, and the distribution of cartilage
rings around the large airways, but the most important differences
between the mouse and human lung are related to the organism’s
size (airway diameter and alveolar size are naturally much smaller in
the mouse) and respiratory rate. Moreover, there are important
differences in the blood supply of the large airways in humans
versus mice [7]. Specifically, the bronchial circulation (a branch of
the high-pressure systemic circulation that arises from the aorta and
intercostal arteries) supplies a miniscule proportion of the pulmonary tissue in mice (the trachea and bronchi) compared to humans;
the majority of the lung parenchyma is supplied by the
low-pressure, high-flow pulmonary circulation. In the mouse,
these systemic blood vessels do not penetrate into the intraparenchymal airways, as they do in larger species [8]. This difference,
although subtle, has important ramifications regarding the vascular
supply of lung tumors which, in humans, is primarily derived from
the systemic circulation [9]. These differences may also have profound consequences when modeling human diseases involving the
lung vasculature.
2.3
Immunology
The adaptive immune system evolved in jawed fish about 500 million years ago, well before the evolution of mammals and the
divergence of mouse and human ancestral species [10]. Many features of the adaptive immune system, including antigen recognition, clonal selection, antibody production, and immunological
tolerance, have been maintained since they first arose in early vertebrates. However, the finer details of the mouse and human immune
systems differ considerably, which is not surprising since these
species diverged 75 million years ago [6]. While some have claimed
that these differences mean that research into immunological phenomena in mice is not transferable to humans, as long as these
differences are understood and acknowledged, the study of mouse
immune responses can continue to be relevant.
Research on mice has been vital to the discovery of key features
of both innate and adaptive immune responses; for example, the
first descriptions of the major histocompatibility complex, the T cell
receptor, and antibody synthesis were derived from experiments
performed on mice [6]. The general structure of the immune
system is similar in mice and humans, with similar mediators and
Mouse Models of Human Disease
7
Table 1
A brief overview of the immunological differences between mice and humans
Attribute
Mouse
Human
References
Proportion of
leukocytes in
the blood
75–90% lymphocytes
10–25% neutrophils
50–70% neutrophils
30–50% lymphocytes
[13]
Antigen
presentation
Endothelial cells do not express Endothelial cells express MHC
Class II and present antigen to
MHC Class II, cannot activate
CD4+ T cells
CD4+ T cells
Costimulatory
signaling
80% of CD4+ and 50% of CD8+
T cells express CD28
ICOS is not required for B cell
maturation
B7-H3 inhibits T cell activation
[14]
100% of CD4+ and CD8+ T cells [12]
express CD28
ICOS is required for B cell
[15, 16]
maturation and IgM production
B7-H3 promotes T cell activation [17]
Immunoglobulin IgD, IgM, IgA, IgE, IgG1,
isotypes
IgG2a/c, IgG2b, IgG3
IgD, IgM, IgA1, IgA2, IgE, IgG1, [12]
IgG2, IgG3, IgG4
Immunoglobulin IL-4 induces IgG1 and IgE
class switching
IL-4 induces IgG4 and IgE
[18]
Helper T cell
differentiation
IFN-α does not activate STAT4 IFN-α induces Th1 polarization via [19]
STAT4
and does not induce Th1
polarization
Clear Th1/Th2 differentiation in Multiple T helper cell subsets occur [20]
mice
simultaneously
Responses to
infection
Eradication of schistosomiasis
requires a Th1 response and
IFN-γ
Low susceptibility to
Mycobacterium tuberculosis;
noncaseating granulomas; no
latent infection
[21]
Eradication of schistosomiasis
requires a Th2 response and IgE
Highly susceptible to
Mycobacterium tuberculosis;
caseating granulomas; latent
infection is common
[22]
cell types involved in rapid, innate immune responses (complement,
macrophages, neutrophils, and natural killer cells) as well as adaptive immune responses informed by antigen-presenting dendritic
cells and executed by B and T cells. However, due to the anatomical
and physiological differences between these species as described
above, divergence in key features of the immune system, such as
the maintenance of memory T cells (related to the life span of the
organism) and the commensal microbiota (related to the lifestyle of
the organism), has arisen [11].
Similar to what has been discovered regarding the genetics of
mice and humans, i.e., broad similarities in structure but considerable differences in regulation, there are a number of known discrepancies in the regulation of innate and adaptive immunity in
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Kristina Rydell-Törma
mice versus humans, including the balance of leukocyte subsets, T
cell activation and costimulation, antibody subtypes and cellular
responses to antibody, Th1/Th2 differentiation, and responses to
pathogens (described in detail in Table 1). In addition to these
differences in immune cell functions, the expression of specific
genes involved in immune responses also differs, particularly those
for Toll-like receptors, defensins, NK inhibitory receptors, Thy-1,
and many components of chemokine and cytokine signaling; additionally, differences between mouse strains are known to exist for
many of these mediators [12].
Another important consideration when using mice to perform
immunological research (with a view to translating these findings to
human medicine) is the availability of hundreds of strains of genetically modified mice that have enabled exquisitely detailed studies
on immune cell function, regulation, and trafficking. Many of these
strains involve the expression of inducible Cre or Cas9 that allow
for targeted knockdown or overexpression of key immune
function-related genes in specific cell types at specific moments in
time. However, it is important to note that drift between mouse
colonies has long been known to occur. In fact, a recent report
described the fortuitous discovery of a point mutation in the natural cytotoxicity receptor 1 (NCR1) gene in the C57/Bl6 CD45.1
mouse strain, resulting in absent NCR1 expression. This mutation
was found to have profound effects on the response of mice to viral
infection, i.e., the mice were resistant to cytomegalovirus infection
but more susceptible to influenza virus [23]. This cautionary tale
highlights the importance of understanding the genetic evolution
of laboratory strains of mice, the effect of these genetic and immunological changes on mouse biology, and the impact on the translation of these results to human medicine.
In addition to the differences between mouse and human
genetics, physiology, and immunology highlighted above, several
factors must also be taken into account when performing in vitro
assays using isolated mouse cells and applying these findings to our
understanding of human disease. Particularly with regard to stem
cell research, it should be noted that the telomeres of mouse cells
are five- to tenfold longer than human telomeres, resulting in
greater replicative capacity [24]. There are also important differences in the regulation of pluripotency and stem cell differentiation
pathways in humans and mice [25]. Moreover, there are considerable species differences in the longevity of cultured cells; for example, mouse fibroblasts are capable of spontaneous immortalization
in vitro, whereas human fibroblasts become senescent and ultimately fail to thrive in culture [26].
In summary, although there are considerable differences
between mice and humans, constant improvement in the analytical
techniques used to delineate these differences and their effects on
whole organism and cell function have provided vital information
Mouse Models of Human Disease
9
and contributed to our understanding of both murine and human
biology. Experimentation employing mouse models of human disease will continue to provide key insights into relevant disease
mechanisms and potential drug targets for further clinical investigation. However, several important considerations must be taken
into account when selecting a mouse model of human disease, as
described in the following section, using mouse models of human
lung disease to illustrate this point.
3
Mouse Models of Human Disease
The two most salient features of a mouse model of human disease
are the accuracy of its etiology (it employs a physiologically relevant
method of disease induction) and its presentation (its ability to
recapitulate the features of human disease). The relevance of any
given mouse model can be judged on the basis of these two criteria,
and there is considerable variation within mouse models of human
disease in this regard. As a full assessment of the advantages and
limitations of all currently available mouse models of human disease
would be prohibitively long and complex, here we have elected to
assess the accuracy of currently available models of human lung
diseases, i.e., asthma, chronic obstructive pulmonary disease, and
pulmonary fibrosis, focusing on the relevance of disease induction
in these models and their ability to replicate critical features of
human disease pathophysiology and response to treatment.
The first and foremost notion when modeling human disease in
mice is to acknowledge the species differences, which are significant
[27]. As described above, genetics, anatomy, physiology, and
immunology differ between mice and humans, but despite these
differences, mouse models of human disease are useful and necessary, as long as data interpretation is performed appropriately.
4
Asthma
An elegant example of differences between mice and humans that
must be considered when designing a mouse model of human
inflammatory lung disease is the key effector cell type in human
asthma, i.e., mast cells. These leukocytes differ in granule composition as well as localization in the mouse and human airways
[28]. Mice mostly lack mast cells in the peripheral lung [29],
whereas humans have numerous mast cells of multiple subpopulations in the alveolar parenchyma [30]. Another example is anatomy:
in contrast to humans, mice lack an extensive pulmonary circulation, which may have significant effects on leukocyte adhesion and
migration, and subsequently inflammation [31]. Still, as long as
these differences are taken into consideration, mouse models can be
10
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Kristina Rydell-Törma
powerful tools in the discovery and exploitation of new targets for
the treatment of human disease.
The World Health Organization (WHO) defines asthma as a
chronic disease characterized by recurrent attacks of breathlessness
and wheezing, which may vary in severity and frequency from
person to person. The disease is characterized by airway hyperresponsiveness, airway smooth muscle thickening, increased mucus
secretion and collagen deposition, as well as prominent inflammation affecting both large and small airways [32]. Nowadays, it is
recognized that asthma is not a single homogenous disease but
rather several different phenotypes united by similar clinical symptoms [32, 33].
Only a few animal species develop asthma naturally, including
cats and horses [34, 35], whereas mice do not [31]. However, mice
can be manipulated to develop a type of allergic airway inflammation, which is similar in many ways to the human disease, in
response to different aeroallergens [36]. Importantly, these models
are capable of recapitulating only the allergic type of human asthma
and have less relevance for other types of asthma (i.e., endotypes
induced by medication, obesity, and air pollution).
As with many human diseases, asthma has a complex and
multifaceted etiology, where environmental factors, genetic susceptibility, and microbial colonization all contribute; thus, it is important to take strain differences into consideration. Generations of
inbreeding have created mouse strains that differ not only in coat
color and disposition but also from a physiological, immunological,
and genetic perspective. Different strains may be more susceptible
to allergic airway inflammation or pulmonary fibrosis, whereas
others are more or less resistant. Choosing the right strain to
model a specific disease or pathologic event is thus essential. The
most widely used strains for models of allergic airway inflammation
are BALB/c and C57BL/6. These strains differ regarding the type
of immune response mounted to an inhaled allergen: C57BL/6 is
generally considered a TH1-skewed strain, whereas BALB/c is
regarded as a TH2-skewed strain [36]. Due to their strong TH2response, and subsequent development of robust asthmatic
responses, BALB/c has been commonly used to model asthma
[37]. However, most humans do not express such a strongly
TH2-skewed immune system, suggesting this strain may not be
the best model of human disease; instead, C57BL/6 may be
more suitable as immune responses in this strain are more similar
to those of atopic human subjects [37]. Furthermore, as C57BL/6
is the most commonly used strain for the development of genetically manipulated mice, using these mice allows for very specific
investigations into disease pathology; thus, this strain is increasingly
used in models of human lung disease.
Mouse Models of Human Disease
11
4.1
Ovalbumin
Besides the genetic differences in the mouse strains used in these
models, the etiology (the method of disease induction) of commonly used models of asthma is highly variable. In humans with
allergic asthma, environmental allergen exposure occurs at the
airway mucosa; the immune response is coordinated in the bronchopulmonary lymph nodes, and the T cells, macrophages, and eosinophils recruited as part of this response travel to the lung where
they mediate the cardinal features of asthma: airway inflammation,
structural remodeling of the airway wall, and airway hyperreactivity
[38]. Ideally, these features should be found in a physiologically
relevant mouse model of asthma. However, for the sake of cost and
convenience, early mouse models of asthma used the surrogate
protein ovalbumin (OVA) [31] rather than an environmental allergen to induce an immune response, which also requires the use of a
powerful TH2-polarizing adjuvant such as alum delivered via the
intraperitoneal route, followed by OVA nebulization—a clear
divergence from the etiology of human asthma [36]. In terms of
disease presentation, mice develop some hallmarks of asthma,
including airway eosinophilic inflammation, goblet cell metaplasia,
and increased airway smooth muscle density [31]. After the cessation of OVA exposure, most of the remodeling resolves, although
some structural alterations remain up to 1 month after the last
challenge [39]. Based on these attributes, the OVA model is primarily a model to investigate the initiation of inflammation, rather
than the
chronic progression
and
maintenance
of
inflammation [31].
A clear advantage with the OVA model is the number of studies
where it is used; both the pros and cons are familiar. It is easy to find
a suitable protocol, and the model is readily accessible and flexible
regarding the number of sensitizations and allergen doses. The
model is relatively easy to reproduce, as OVA and different adjuvants are easily obtained. However, the resolution of remodeling
following the cessation of allergen provocations is a disadvantage,
as is the practical problem with the nebulization of an allergen—it
ends up in the mouse’s coat and is ingested during grooming,
potentially resulting in systemic exposure (this is particularly relevant in models employing systemic, intraperitoneal sensitization).
In addition, concerns have been raised against the use of adjuvants
to induce the immunological response, as well as the clinical relevance of OVA as an allergen, which have driven the development of
more clinically relevant allergens and models [31].
4.2
House Dust Mite
The common environmental aeroallergen house dust mite (HDM)
extract is increasingly used to initiate disease in mouse models of
allergic airway inflammation, as it is a common human allergen
(around 50% of asthmatics are sensitized to HDM [40]) that evokes
asthma attacks and other allergic responses in susceptible individuals. In addition, HDM has inherent allergenic properties, likely
12
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Kristina Rydell-Törma
due to components with protease activity [40], so there is no need
to use an adjuvant, thus improving the etiological similarity of these
models with the clinical situation [41]. In contrast to OVA, prolonged exposure of HDM (up to 7 weeks) induces asthma-like
severe airway inflammation with prominent eosinophilia, severe
hyperreactivity to methacholine, and robust remodeling of the
airway wall [41], i.e., the presentation of chronic respiratory
HDM exposure in mice effectively recapitulates the key features
of human allergic asthma. Importantly, the airway structural
changes induced by chronic HDM exposure, such as increased
collagen deposition, airway smooth muscle thickening, and microvascular alterations, persist for at least 4 weeks after the cessation of
HDM exposure [42], another commonality with human asthma in
which airway remodeling is currently considered to be irreversible.
Thus, the advantages of using HDM as the allergen in mouse
models of asthma are the clinical relevance of the allergen [43] and
the route of delivery via the respiratory tract. Moreover, studies
have shown that the type of inflammation and characteristics of
tissue remodeling are relatively similar to those seen in human
asthmatics [35, 41, 43]. One disadvantage is the complexity of
HDM extract; as a consequence of this complexity, variations exist
in some components between batches, particularly regarding the
content of lipopolysaccharide, so reproducibility in these studies
may be problematic.
4.3 Cockroach,
Aspergillus, and Other
Model Allergens
With similarity to HDM, these models were developed to be as
clinically relevant as possible, as many patients suffer from allergy
toward cockroach allergen, molds, and other environmental irritants. A common feature of these allergens is their complex nature,
as they commonly consist of a mix of different allergic epitopes and
fragments. This complexity is most likely why the immunological
reaction in mice is relatively similar to that seen in asthmatics [44].
Cockroach allergen (CRA) is a common allergen, known to
induce asthma in susceptible individuals; thus, it shares with HDM
the advantage of being highly clinically relevant [45]. CRA induces
peribronchial inflammation with significant eosinophilic inflammation and transient airway hyperresponsiveness, both of which can
be increased by repeated administrations of the allergen [45].
Colonization of the airways with Aspergillus fumigatus is the
cause of allergic bronchopulmonary aspergillosis (ABPA), a disease
where the lungs are colonized by the fungus, but allergens from
Aspergillus fumigatus can also induce asthma similar to other allergens [46]. The reaction to Aspergillus allergens is robust, and often
no adjuvants are needed to elicit inflammation [46]. In addition to
Aspergillus, other fungi such as Penicillium and Alternaria can also
induce asthma in humans and have been used to model disease in
mice [47]. A common difficulty with these allergens is the method
of administration, as the physiological route is believed to be the
Mouse Models of Human Disease
13
inhalation of dry allergens; mimicking this route with a nebulizer
introduces the risk of the animals ingesting the allergen and thus
causing systemic responses [47].
4.4 Modeling Asthma
Exacerbations
Exacerbations of asthma are defined as the worsening of symptoms,
prompting an adjustment in treatment, and are believed to be
associated with increased inflammation in the distal airways. Clinically, exacerbations are believed to be induced by infections (most
common), allergen exposure, or pollutants, which can be modeled
in different ways [48, 49]:
1. Infections with viruses and bacteria or exposure to proteins/
DNA/RNA derived from these microbes.
2. Administration of a high dose of allergen in a previously sensitized animal.
3. Exposure to environmental pollutants, such as diesel exhaust or
ozone.
Modeling exacerbations adds a layer of complexity, as robust
ongoing allergic airway inflammation needs to be established first,
before challenge with the exacerbating agent. Both the OVA and
HDM models are used in this respect, and in both cases chronic
protocols extending for several weeks before triggering an exacerbation have been used [48].
5
Chronic Obstructive Pulmonary Disease
Chronic obstructive pulmonary disease (COPD) is characterized by
chronic airway obstruction, in contrast to asthma where the
obstruction is reversible (particularly in response to bronchodilator
treatment). Clinically, in COPD, chronic bronchitis and emphysema can occur either separately or in combination. COPD is
almost always associated with either first- or secondhand tobacco
smoking or in rare cases with a deficiency in the production of α1antitrypsin (a serpin that prevents elastin breakdown as a result of
neutrophil degranulation) [50]. The etiology of COPD is highly
complex and is believed to develop after many years of smoking in
combination with other known factors such as genetic susceptibility
or environmental factors [51]. In similarity to asthma, inflammation is a major component in COPD, but the leukocyte profile is
very different: the most prominent players in COPD-related
inflammation are neutrophils and, to some degree, macrophages
[51]. Due to the complex etiology of COPD, it is difficult to
recapitulate all aspects of this disease in a single model, so in most
cases, the aim is to induce COPD-like lesions by exposing mice to
tissue-damaging substances (usually cigarette smoke) or to mimic
emphysema by the administration of tissue-degrading enzymes
[27, 51].
14
5.1
€nen and Jill R. Johnson
Kristina Rydell-Törma
Cigarette Smoke
5.2 Protease
Instillation
6
Clearly, mice do not smoke cigarettes on their own, so to model
COPD by cigarette smoke (CS) inhalation, the mice need to be
exposed to unfiltered CS in an induction chamber; moreover, in an
attempt to better model the chronic aspects of COPD, this needs to
be performed for a prolonged period of time. Mice are very tolerant
to CS, but eventually (over a period of several weeks), CS induces
pulmonary neutrophilic inflammation that is associated with some
degree of tissue degradation and destruction [51]. An important
advantage of this model is the fact that CS is the actual irritant
responsible for disease in humans, and mice develop several features
similar to the clinical disease, making this model highly clinically
relevant [27]. A significant drawback is the self-limitation of the
model—the pathological changes do not progress after the cessation of CS exposure [51]. Furthermore, the exposure time needed
for mice to develop COPD-like pathology is extensive, i.e., studies
have shown that an exposure protocol of 5 days per week for a
minimum of 3 months is needed to generate robust structural
changes to the lung [52]. The pathological image in COPD is
complex and varies greatly between patients, commonly encompassing chronic bronchitis and bronchiolitis, emphysema, fibrosis,
and airway obstruction. Although mice develop some of these
symptoms when exposed to CS, they do not develop all the symptoms of human disease; thus, CS has advantages as a model but fails
to mimic the complexity of the clinical situation and disease
presentation [27].
Other models of COPD rely on the administration of proteases
(protein-degrading enzymes) that are believed to be involved in the
pathology of this disease in a subset of patients, such as elastindegrading elastase. This approach mimics the emphysematous
changes seen in COPD, but the pathological process underlying
tissue destruction is likely very different compared to the clinical
situation [51], as very few patients show evidence of elastase dysregulation [27]. However, if the aim of the study is to investigate the
general effect of protease-induced tissue destruction and regeneration, then this is a highly relevant method [51]. Some studies on
COPD have also used genetically modified animals, such as mice
overexpressing collagenase, which results in tissue destruction
without inflammation or fibrosis with an end result fairly similar
to the type of emphysema observed in COPD [53].
Pulmonary Fibrosis
Pulmonary fibrosis, the accumulation of fibrotic tissue within the
alveolar parenchyma, is merely a symptom of disease, and the
etiology of this pathology in humans varies greatly [54]. The
Mouse Models of Human Disease
15
most enigmatic class is perhaps the idiopathic interstitial pneumonias, especially idiopathic pulmonary fibrosis (IPF). IPF is a debilitating and progressive disease with a grave prognosis, characterized
by progressive fibrosis believed to reflect aberrant tissue regeneration [55]. As the reason behind this defective repair is unknown,
although a combination of immunological, genetic, and environmental factors are suspected, it is very difficult to model disease in a
clinically relevant fashion [56]. The most common method used to
model pulmonary fibrosis in mice is administration of the chemotherapeutic agent bleomycin; this agent is known to cause pulmonary fibrosis in humans as well, but this may not accurately reflect
the true etiology of most cases of human disease. The strain of
choice is C57BL/6, as it is prone to developing pulmonary fibrosis,
whereas BALB/c is relatively resistant, a feature believed to reflect
the cytokine response following cellular stress and damage
[57]. Bleomycin administration can be performed locally or systemically, producing very different results.
6.1 Local Bleomycin
Administration
The most common model of pulmonary fibrosis is a single intranasal or intratracheal administration of bleomycin, with analysis 3 to
4 weeks later. During this time, the drug causes acute tissue damage
in a restricted area of the lung (where the solution ends up during
administration), followed by intense inflammation in this area and
subsequent fibrosis, which gradually resolves within weeks. However, if older mice are used, the fibrosis will persist longer than in
younger mice, which is in accordance with clinical IPF, where the
majority of the patients are 65 years of age or older [56, 58].
A great advantage of this model is how well-characterized it
is. In addition, local administration is labor-effective, as only one
administration is required and the result is highly reproducible. The
fibrosis is robust, only affects the lungs, and the accumulation of
extracellular matrix can be easily measured using standard techniques [58]. Furthermore, as it is used throughout the world, studies
performed in different labs and by different groups can be compared relatively easily. Unfortunately, the intense pulmonary
inflammation may be lethal, and fatalities are to be expected with
this model [59], representing an important ethical limitation. Furthermore, fibrosis is heterogeneous—it develops where the bleomycin solution is deposited. The solution usually deposits within
the central lung, a localization that is not in agreement with the
clinical situation where fibrosis is located in the more distal regions
of the lung parenchyma. In addition, the fibrosis that develops as a
result of severe tissue damage is self-limiting and reversible, unlike
what is observed clinically [58]. The severe degree of tissue damage
induced by bleomycin may in fact be more relevant for modeling
acute lung injury (ALI) or acute respiratory distress syndrome
(ARDS).
16
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Kristina Rydell-Törma
6.2 Systemic
Bleomycin
Administration
Bleomycin can also be administered systemically, through intravenous or subcutaneous injection. In contrast to local administration,
this route requires multiple administrations and is thus more laborintensive [58]. Some studies have described the usage of osmotic
mini-pumps, where bleomycin is slowly administered over a short
period of time, and then fibrosis continues to develop over
subsequent weeks [60]. Irrespective of the route of delivery, systemic administration results in more homogenous fibrosis, affecting the entire lung through the pulmonary endothelium and
persisting much longer than following local administration
[61]. The main advantages of systemic administration are that
inflammation is limited, while the fibrosis is more apparent and
displays a more distal pattern, all of which mimics the clinical
situation relatively well. The multiple administrations allow for
lower doses with each injection; this is less stressful to the animals
and results in little to no mortality [61] and is thus more ethically
acceptable. A major disadvantage with this model is that it takes
time for fibrosis to develop [58], which may be the reason it is used
relatively scarcely, and thus the pathological development is less
well-understood. In addition, as IPF is a local disease, local administration of the etiologic agent may better mimic the clinical
reality [56].
6.3 Fluorescein
Isothiocyanate
Administration
The administration of fluorescein isothiocyanate (FITC) induces
focal inflammation, primarily involving mononuclear cells and neutrophils, and localizes in areas where the FITC solution is deposited
[58]. Antibodies against FITC can be detected after 1 week, and
the fibrosis persists for up to 5 months after instillation [58]. The
benefits of this model are mainly related to the persistent fibrosis
that does not appear to be self-limiting, thus reflecting the clinical
situation, and it is also very easy to determine which part of the lung
has been exposed to FITC, as the molecule is fluorescent [58]. It is
also an advantage that both C57BL/6 and BALB/c mice are
susceptible and develop fibrosis following FITC administration
[56]. The disadvantages of this model include profound variability
due to differences between batches of FITC, as well as in the
method used to prepare the solution before instillation. Importantly, given the characteristics of the etiologic agent used to induce
this model of IPF, this model is considered a very artificial system
with limited clinical relevance [56].
6.4 TGF-β
Overexpression
Adenovirus vectors have been used to overexpress the pro-fibrotic
cytokine transforming growth factor (TGF)-β, which results in
pulmonary fibrosis. As TGF-β overexpression in the lungs is
known to be crucial in the development of fibrosis in humans
[62], this model mimics an important feature of disease etiology.
However, the delivery system has some drawbacks, as the virus itself
initiates an immune response. Moreover, adenoviruses display
Mouse Models of Human Disease
17
significant tropism for epithelial cells and rarely infect other cell
types such as fibroblasts [58], which are the cells meant to be
targeted in this model. As TGF-β has major effects on fibroblast
biology, the main feature of this model is the effect of epitheliumderived TGF-β on fibroblasts and myofibroblasts, resulting in the
deposition of ECM proteins and areas of dense fibrosis [63]. An
advantage of this model is the relatively low degree of inflammation, as well as what appears to be a direct effect on fibroblasts/
myofibroblasts [63], which is in accordance with the clinical situation (as we understand it today).
Silica administration induces a similar pathology in mouse lungs as
in humans exposed to silica, and as is also observed in human silicainduced fibrosis, structural remodeling persists when administration is halted [56]. Following the administration of silica particles,
fibrotic nodules develop in mouse lungs, with considerable resemblance to the human lesions that develop after exposure to mineral
fibers [56]. The fibrotic response is accompanied by a limited
inflammatory response, and different pro-fibrotic cytokines such
as TGF-β, platelet-derived growth factor, and IL-10 are involved in
disease development, which is in accordance with the clinical situation [56]. Another advantage is that nodules develop around silica
fibers, and these fibers are easy to identify by light microscopy. The
response in this model is strain-dependent, with C57BL/6 mice
being the most susceptible. The main drawbacks are the time
required to establish disease, i.e., 30–60 days, and the need for
special equipment to aerosolize the silica particles. However, since
the route of administration, the driving etiologic agent, and the
resulting pathobiology are all similar to the characteristics of this
subtype of pulmonary fibrosis [56, 58], the silica exposure model
can be considered to have very good clinical relevance.
6.5
Silica
7
What Does the Future Hold for Mouse Models of Human Disease?
Medical research using experimental animals (not only mice but
other animals including rats, guinea pigs, zebrafish, and fruit flies)
has greatly contributed to many important scientific and medical
advances in the past century and will continue to do so into the near
future. These advances have contributed to the development of
new medicines and treatments for human disease and have therefore played a vital role in increasing the human life span and
improving quality of life.
Despite the acknowledged benefits of performing research
using experimental animals, a number of considerations must be
made before embarking on this type of research. Of course, the
financial aspects of conducting this type of work are an important
limitation, as the costs of purchasing and housing mice can be
prohibitive, especially when genetically modified mice and colony
18
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Kristina Rydell-Törma
maintenance are required for the study. The practicalities of working with animals such as mice may also be an issue, as this type of
work requires specialized facilities, equipment, and staff to ensure
studies are carried out in a manner that is safe for both the researchers and the animals. Moreover, as discussed in detail in this chapter,
the relevance of the selected animal model to human disease must
be carefully evaluated to ensure that these experiments provide
robust results that are translatable to human health and disease.
Another important and demanding aspect of biomedical research
using animals is the ethics of imposing pain and suffering on live
animals.
Although there has been a considerable reduction in the numbers of animals used in research in the last 30 years, animal research
remains a vital part of biomedical research. However, no responsible scientist wants to cause unnecessary suffering in experimental
animals if it can be avoided, so scientists have accepted controls on
the use of animals for medical research. In the UK, this ethical
framework has been enshrined in law, i.e., the Animals (Scientific
Procedures) Act 1986. This legislation requires that applications for
a project license to perform research involving the use of “protected” animals (including all vertebrates and cephalopods) must
be fully assessed with regard to any harm imposed on the animals.
This involves a detailed examination of the proposed procedures
and experiments, and the numbers and types of animal used, with
robust statistical calculations to support these numbers. The
planned studies are then considered in light of the potential benefits
of the project. Both within and outside the UK, approval for a study
involving protected animals also requires an internal ethical review
process, usually conducted by the research institution where the
work is taking place, with the aim of promoting animal welfare by
ensuring the work will be carried out in an ethical manner and that
the use of animals is justified. Additionally, the UK has a national
animal use reduction strategy supported by the National Centre for
the Replacement, Refinement and Reduction of Animals in
Research (NC3Rs; London, UK). This consortium was established
in 2004 to promote and develop high-quality research that takes
the principles of replacement, refinement, and reduction (the 3Rs)
into account.
7.1
Replacement
Replacement strategies often involve the use of alternative,
non-protected species (e.g., zebrafish, fruit flies, flatworms) and
in vitro correlates (two-dimensional cell culture or threedimensional organoids containing multiple cell types) to test
hypotheses and assess the effects of therapeutic interventions. The
main obstacle with studies on non-protected animals is the difficulty of accurately mimicking the complex physiological systems
involved in human health and disease, as described in detail above.
For example, the fruit fly Drosophila melanogaster is an excellent
Mouse Models of Human Disease
19
model organism for studies on genetic diseases, aging, and
pathogen-borne illnesses but may be less relevant for studies on
complex lung diseases. Importantly, model organisms such as fruit
flies, zebrafish, and flatworms do not possess lungs, which somewhat limits the translatability of research on these animals in the
field of respiratory disease. As such, it is likely that rodents will
remain the model organism of choice for studies into lung disease
for some time to come.
There has been considerable progress recently in imitating
single organs such as the liver, lung, and brain in vitro using
multiple cell types and a physical scaffold. As an important advantage, these in vitro tests have replaced a large number of rodents in
initial drug discovery experiments, while also speeding up the
process [64]. These studies still require further refinement and
validation to establish them as suitable models for an entire organ;
importantly, these in vitro organoids cannot take into account
interactions between organ systems in complex, multisystem diseases such as COPD.
7.2
Refinement
Refinement involves selecting the most clinically relevant model for
the disease available, informed by the discussion above on closely
recapitulating the etiologic agent and disease pathobiology associated with clinical cases. Another important factor is refining the
management of pain. An assessment of the procedures used and the
effects of the substance on the animal, as well as the degree of
handling, restraint, and analgesia, are other important aspects of
refinement. This standard of animal care is achieved through strict
regulations and controls on how personnel are trained to carry out
experiments on live animals. Adequate training is an important
aspect of refinement and should be reviewed and improved on an
ongoing basis. Moreover, refinement can be achieved by improving
animal housing by environmental enrichment, e.g., providing a
place for mice to hide in the cage and housing social animals such
as mice in appropriate-sized groups. These simple changes can
improve the physiological and behavioral status of research animals;
this not only increases animal well-being but also contributes to the
quality of the experimental results by reducing stress levels.
7.3
Reduction
The 3Rs aspect of reduction focuses on the statistical power of
experiments and by following the Animal Research: Reporting of
In Vivo Experiments (ARRIVE) guidelines, originally published in
PLOS Biology in 2010. These guidelines provide a framework to
improve the reporting of research performed on live animals by
maximizing the quality of the scientific data and by minimizing
unnecessary studies. The ARRIVE guidelines provide a checklist of
aspects that must be considered in good quality research using live
animals. The guidelines are most appropriate for comparative studies involving two or more groups of experimental animals with at
least one control group, but they also apply to studies involving
20
€nen and Jill R. Johnson
Kristina Rydell-Törma
drug dosing in which a single animal is used as its own control
(within-subject experiments). The guidelines provide recommendations on what should be considered when preparing to report on
the results of experiments involving live animals, i.e., by providing a
concise but thorough background on the scientific theory and why
and how animals were used to test a hypothesis, a statement on
ethical approvals and study design including power and sample size
calculations, a clear description of the methods used to ensure
repeatability, objective measurements of outcomes and adverse
effects, and interpretation of the results in light of the available
literature and the limitations of the study. In addition to the positive impact of the ARRIVE guidelines on reducing the number of
animals used in experiments, this checklist provides an easy-tofollow roadmap on what is required for good quality reporting of
experimental results.
8
Conclusion
In conclusion, the use of animals in research will continue to be an
important aspect of medical research, and these procedures can be
ethically justified provided the proper controls are in place. The
benefits of animal research have been vital to the progress of medical science; abandoning these studies would have severe negative
consequences on human health. By considering aspects such as the
3Rs and the ARRIVE guidelines in planning experiments involving
live animals, the number of animals used and suffering of these
animals for the benefit of human health can be minimized. This
requires a strong regulatory framework such as that found in the
UK and many other countries, as well an ongoing public debate on
the advantages and limitations of animal experimentation.
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Chapter 2
Optimizing the Cell Culture Microenvironment
Ivan Bertoncello
Abstract
The survival, proliferation, and differentiation of cells in culture are determined not only by their intrinsic
potential but also by cues provided by the permissive or restrictive microenvironment in which they reside.
The robustness and reproducibility of cell culture assays and endpoints relies on the stability of that
microenvironment and vigilant attention to the control of variables that affect cell behavior during culture.
These often underappreciated variables include, but are not limited to, medium pH and buffering,
osmolarity, composition of the gas phase, the timing and periodicity of refeeding and subculture, and the
impact of fluctuations in temperature and gas phase composition on frequent opening and closing of
incubator doors. This chapter briefly describes the impact of these and other variables on the behavior of
cultured cells.
Key words Cell culture, Culture medium, Medium pH, Medium buffering, Oxygen tension, Incubation conditions
1
Introduction
The niche hypothesis first articulated by Schofield [1] to explain
hematopoietic regulation posits that the regenerative potential of a
cell population is defined in context: by its intrinsic potential and by
its interaction with the microenvironment in which it resides
[2, 3]. The dynamic interaction of regenerative cells with soluble
and insoluble factors, signaling pathways, accessory cells, and
matrix proteins comprising their microenvironment regulates
their developmental potential, proliferation, and differentiation
(Fig. 1). Conversely, reciprocal signaling mediated by proliferating
and differentiating cells is able to modify their niche microenvironment, potentially leading to dysregulated cell growth [4–6].
The degree of difficulty encountered in deconstructing and
elucidating these regulatory processes to develop informative and
instructive in vitro cell culture models cannot be underestimated.
In 1993, Quesenberry [7] estimated that there were at least 2.248
possible combinations of 40 known hemolymphopoietic cytokines
with order being important and without allowing for dose-
Ivan Bertoncello (ed.), Mouse Cell Culture: Methods and Protocols, Methods in Molecular Biology, vol. 1940,
https://doi.org/10.1007/978-1-4939-9086-3_2, © Springer Science+Business Media, LLC, part of Springer Nature 2019
23
24
Ivan Bertoncello
Fig. 1 The developmental potential, proliferation, and differentiation of cultured cells are determined by their
spatial orientation, their dynamic interaction, and their elaboration of soluble and insoluble stimulatory and
inhibitory factors and matrix proteins that comprise their microenvironment
dependent differences in activity or target cell heterogeneity. Variables including cell adhesion and cell geometry [8], cell polarity
[9], and extracellular matrix stiffness [3, 10] have also been identified as significant factors that influence the developmental fate,
proliferation, and differentiation of different cell types. Over the
years, reductionist experimentation exploiting powerful biochemical and molecular genetic technologies and cell separative strategies
has enabled the progressive refinement of cell culture technologies.
However, the optimization of cell culture assays and the identification and elimination of sources of experimental variability remain a
work in progress [11].
Historically, the development of protocols for the maintenance
and propagation of specific cell types has focused on the formulation and optimization of chemically defined tissue culture media
that meet their requirement for unique combinations of nutrients,
trace elements, growth factors, and hormones in order to thrive
in vitro [11–13]. Often, optimal growth requires the addition of
uncharacterized supplements such as fetal calf serum, conditioned
media, tissue extracts, extracellular matrices, or hydrogels to chemically defined media. In addition to variable concentrations of
known factors, these supplements contain a plethora of ill-defined
peptides, proteins, and organic and inorganic molecules potentially
capable of stimulating or inhibiting target cells. In one study [14],
in-depth proteomic analysis identified a total of 14,060 unique
peptides and 1851 unique proteins in different lots of standard
and growth factor reduced (GFR) Matrigel preparations obtained
from different suppliers, with only 53% batch-to-batch similarity
in GFR Matrigel lots based on protein identification. These constituents will potentially affect the developmental potential,
The Cell Culture Microenvironment
25
proliferation, and differentiation of cultured cells, by directly stimulating target cells synergistically or additively in a dosedependent fashion, or by indirectly activating or suppressing accessory cells inducing cytokine loops and cascades. Batch variation in
these supplements is often a significant source of unexplained qualitative and quantitative differences in cell culture outcomes and
assay readouts within laboratories or among different laboratories.
Consequently, batch testing of these reagents is critical in order to
ensure the stability and reproducibility of cell culture systems over
time. Ideally aliquots of reagents with optimal growth supporting
properties should be stored long term as a reference standard or as
an aid in identifying confounding factors affecting reproducibility
or replication of assay readouts.
The commercial availability of highly defined cell culture media
supplemented with unique combinations of essential nutrients,
trace elements, hormones, and growth factors has enabled the
development of cell culture systems for the maintenance, propagation, and manipulation of virtually all embryonic and adult cell
types and cell lineages. However, cell-dependent variables, colligative properties of cell culture media, and the stability of the cell
culture environment (Table 1) merit much greater attention when
looking to optimize cell culture systems and improve the stability
and reproducibility of cell culture assays.
Table 1
Variables affecting the proliferation and differentiation of cultured cells
and the reproducibility of cell culture endpoints
Heterogeneity of the initial cell inoculum
Cell plating density
Split ratio, growth phase, and cycling status of cultured cells
Timing and frequency of refeeding
Composition of the gas phase: pO2 and pCO2
Culture medium pH and buffering
Volume and depth of cell culture medium
Medium osmolarity
Stability of tissue culture reagents
Stability of incubation conditions—temperature, gas phase
Period of incubation and criteria for culture endpoint (e.g., colony size)
26
2
Ivan Bertoncello
Cell-Dependent Variables
The defined medium in which cells are propagated is only
“defined” at the initiation of culture. From the moment cells are
suspended in the defined medium, the properties of the culture
system are instantly and progressively altered. The initial cell density
and the heterogeneity of the cell inoculum are significant variables
affecting the stability and reproducibility of cell culture outcomes.
Cells interact in culture to secrete autocrine, juxtacrine, holocrine, and/or endocrine factors that modify their microenvironment to affect their survival, developmental potential, and rate of
proliferation and differentiation (Fig. 1). Single cells or cells growing at clonal cell densities have more fastidious requirements than
cells propagated at high cell densities [12]. The large volume of
medium per cell in clonal cell cultures significantly compromises
their ability to quickly generate optimal concentrations of secreted
factors required for their survival and growth, potentially
compromising their survival, proliferation, and differentiation.
The replicative capacity, viability, and dynamic changes in the
pattern of growth and differentiation of cells propagated in longterm culture also vary in response to the periodicity of refeeding,
the phase of cell growth (exponential or stationary) at the time
cultures are split, and the uniformity of split ratio. For example,
hematopoietic stem cells secrete a large repertoire of stimulatory
and inhibitory factors in the course of their proliferation and differentiation. Therefore, large fluctuations are observed in the repertoire and concentration of these factors each time medium is
replenished during periodic refeeding of long-term cultures [15]
markedly affecting the dynamics and heterogeneity of cell growth.
The optimal performance and stability of cell culture assays, and the
reproducibility of cell culture endpoints, relies on precise standardization of culture conditions. This includes number of cells, and
the cell density of the cell inoculum, the ratio of the cell number to
the volume of medium, and the timing and periodicity of subculture and refeeding.
3
Medium pH and Buffering
Early pioneers of cell culture recognized that the behavior of
cultured cells is profoundly sensitive to changes in environmental
pH, affecting parameters including protein synthesis, metabolism,
cell growth rate [12, 16, 17], and cell differentiation and cloning
efficiency [18]. Medium acidification as a result of catabolic and
anabolic metabolism and the generation of inhibitory metabolites
also affects the availability of nutrients due to complex interactions
of medium constituents [17, 19] including sequestration of
The Cell Culture Microenvironment
27
CO2
CO2 + H2O
H2CO3
HCO3- + H+
Fig. 2 The pH in cell culture is stabilized by the bicarbonate buffering system.
Equilibrium is maintained by the relationship between the concentration of CO2
in the gas phase and the concentration of HCO3 in the cell culture medium.
Acidification of culture medium drives the equation to the left raising CO2
concentration, whereas raising the concentration of CO2 in the gas phase, or
in the medium due to the metabolic activity of cultured cells, drives the equation
to the right lowering medium pH. For optimal buffering, the concentration of
NaHCO3 in culture medium should be adjusted in line with the concentration of
CO2 in the gas phase
essential nutrients by binding to albumin [20]. Optimal pH differs
markedly for different cell types [16, 17] with some cell types
exhibiting extreme sensitivity to medium acidification [21, 22]
affecting cell cycling, cell growth, and differentiation and also
causing DNA damage and genomic instability [23].
The regulation of pH in cell culture is primarily achieved by
bicarbonate buffering as described by the Henderson-Hasselbalch
equation (Fig. 2). Medium pH is maintained by the equilibrium
between the CO2 concentration in the gas phase and the sodium
bicarbonate (NaHCO3) concentration in the culture medium
[11, 24]. An elevated concentration of CO2 in the gas phase will
drive the equation to the right resulting in elaboration of an
increased concentration of hydrogen ion (H+) and medium acidification. Conversely, medium acidification will drive the equation to
the left increasing the elaboration of CO2. It is not uncommon for
tissue culture protocols developed in different laboratories to specify different CO2 concentrations for the incubator gas phase: commonly 5% CO2 or 10% CO2. In my experience the fact that
differences in the CO2 concentration of the incubator gas phase
affects the buffering capacity of the medium is often overlooked,
potentially affecting the optimal growth of cells and cell lines
sensitive to medium acidification. Ideally, the CO2 tension in the
incubator gas phase and/or the NaHCO3 concentration in different media should be adjusted accordingly [11, 24].
4
The Gas Phase
Cultured cells are most commonly maintained and propagated
under normoxic conditions in a gas phase of 5% CO2 or 10% CO2
in air (i.e., 20% O2). However, there has been a growing awareness
of the benefits of cell culture at low oxygen tension (5% O2),
mimicking the hypoxic environment in which regenerative cells
28
Ivan Bertoncello
reside in tissues and organs in vivo [25, 26]. The reader is also
referred to a comprehensive bibliography of historical studies in this
field provided in supplementary information in the commentary by
Wion et al. [26].
The beneficial effect of low O2 tension is less evident when
analyzing the behavior of cell lines originally selected and propagated long term under normoxic conditions. Nor when cells or cell
lines are grown at high cell density where O2 and CO2 tension in
the pericellular microenvironment tends to be adjusted and regulated by cell metabolism. However, it is a different matter for many
primary explanted cell types and cells grown at clonal or low cell
densities where growth at low O2 tension often results in improved
replicative capacity and cloning efficiency and qualitative and quantitative differences in differentiation potential or the elaboration of
secreted factors.
The expense of purchasing and maintaining triple gas incubators that regulate the delivery of CO2, O2, and nitrogen (N2) is
often cited as an impediment to routine culturing of cells at low O2
tension. However, investigators should be aware that O2 toxicity
and oxidative stress are detrimental to cells in culture [27–29] and
that O2 toxicity can cause spontaneous genetic mutations [30]. A
recent study has also demonstrated that low O2 tension is not only
important during cell propagation but also during cell processing,
noting that the incidence and recovery of hematopoietic stem cells
(HSC) are significantly impaired in hematopoietic tissues processed
in air [31].
5
Osmolarity
Medium osmolarity is another important variable affecting cell
membrane transport and the metabolism, growth, and differentiation of cultured cells [32]. When evaluating the activity of cytokines, supplements, or reagents on cell proliferation and
differentiation, cell metabolism, or gene expression, investigators
should be aware of the possible contribution of these substances to
changes in osmolarity that could affect culture outcome.
Osmolarity of culture medium in open culture vessels can also
be adversely affected by evaporation due to fluctuations in incubator temperature and humidity during injection of dry gases while
purging to re-equilibrate the gas phase following opening and
closing of the incubator. Frequent opening and closing of incubator doors to retrieve and examine cell cultures further exacerbate
this problem creating a progressively hyperosmotic microenvironment in long-term cultures that will ultimately inhibit their proliferation and clonogenicity.
The Cell Culture Microenvironment
6
29
Stability of Incubation Conditions
The robustness of cell culture protocols, and the reproducibility of
cell culture endpoints, relies on careful attention paid to the stability
of incubation conditions during the period of culture: a factor
underappreciated by many investigators. The previous section
alluded to the impact of frequent opening and closing of incubator
doors on the evaporation of medium and medium osmolarity due to
loss of humidity. However, fluctuations in incubator temperature,
and O2 and CO2 tension during examination of cultures, or following purging and re-equilibration of the incubator gas phase are
equally significant, affecting medium pH and medium buffering.
Investigators need to be aware that the equilibration of the gas
phase in the incubator and gas phase inside a tissue culture vessel
can take 30 min [33, 34]. Because equilibration of gas concentration
in culture medium relies on diffusion, the rate of equilibration is also
affected by the depth of the medium. Allen et al. [33] have shown
that equilibration of O2 content in unstirred culture medium can
take more than 3 h due to the low solubility and limited diffusion of
O2 in aqueous solutions, potentially affecting the precision and
reproducibility of cell culture endpoints over time.
7
Conclusion
Cell culture protocols for specific cell types will continue to evolve in
lockstep with our understanding of the nature and function of the
factors and signaling pathways that specify their fate and regulate their
replicative capacity, proliferation, and differentiation. The impact of
each of the variables discussed in this brief review on individual
primary cell cultures or established cell lines will differ. But together,
they are a source of experimental variation that if not appreciated and
controlled potentially affect the reproducibility of cell culture endpoints within laboratories, as well as the ability of investigators to
replicate assays and predictive models in different laboratories.
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(ed) Handbook of experimental pharmacology,
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13. Ham RG (1984) Formulation of basal nutrient
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113:311–322
Part II
Methods and Protocols
Chapter 3
Propagation and Maintenance of Mouse Embryonic
Stem Cells
Jacob M. Paynter, Joseph Chen, Xiaodong Liu, and Christian M. Nefzger
Abstract
Mouse embryonic stem cells (mESCs) are pluripotent cells derived from preimplantation embryos that have
the capacity to self-renew indefinitely in vitro. mESCs are an indispensable tool for studying cellular
differentiation in vitro, generating disease in a dish models, and have been used extensively for the
generation of transgenic animals. Therefore, maintaining their pluripotent state, even after extended
culture, is crucial for their utility. Herein, we describe in detail a protocol for the culture of mESCs in the
presence of fetal calf serum (FCS), leukemia inhibitory factor (LIF), and a layer of irradiated mouse
embryonic fibroblasts (iMEFs). This culture system reliably sustains mESC pluripotency and self-renewal
capacity, allowing their use in a wide range of experimental settings.
Key words Mouse embryonic stem cells, Cell culture, Pluripotency, Mouse embryonic fibroblasts,
Leukemia inhibitory factor
1
Introduction
Pluripotency is defined as the capacity of a cell to give rise to all
somatic cell lineages and the germline of the embryo [1]. Our
ability to maintain pluripotent stem cells in vitro was crucial to
establish them as a platform for the study of early development
in vitro and for the generation of transgenic animals [2]. Historically, pluripotent stem cell culture emerged from findings by
Stevens and Little in 1954, who described a population of undifferentiated cells in mouse testicular teratocarcinomas termed embryonal carcinoma (EC) cells [3]. Most importantly, Kleinsmith and
Pierce found that individual EC cells could give rise to bona fide
teratocarcinomas when deposited into the peritoneum of secondary
mice [4]. These tumors contained cells from each of the three germ
layers, indicating that EC cells were pluripotent. In 1970, the
groups of Sato and Ephrussi succeeded in maintaining EC cells
Jacob M. Paynter and Joseph Chen contributed equally to this work.
Ivan Bertoncello (ed.), Mouse Cell Culture: Methods and Protocols, Methods in Molecular Biology, vol. 1940,
https://doi.org/10.1007/978-1-4939-9086-3_3, © Springer Science+Business Media, LLC, part of Springer Nature 2019
33
34
Jacob M. Paynter et al.
in vitro as monolayer cultures in the presence of fetal calf serum
(FCS) and a basal feeder layer of mitotically inactivated mouse
fibroblasts [5, 6]. Over the next decade, biochemical and functional
analyses showed that EC cells resemble cells of the early embryo
[7–14]. The most stringent of these assays was the ability of EC
cells to form chimeras upon blastocyst injection [15–17]. However,
in the majority of cases, EC cells failed to contribute to the germline
due to their excessive chromosomal abnormalities [18] which precluded their use for the generation of transgenic animals.
Seminal work in 1981 culminated in two independent publications by Evans and Kaufman [19] and Martin [20] which described
the direct in vitro derivation of pluripotent cells from preimplantation mouse embryos. These cells were named “embryonic stem
cells (ESCs)” and could be maintained on a feeder layer of mouse
embryonic fibroblasts (MEFs) in the presence of FCS. Mouse ESCs
(mESCs) were found to readily give rise to germline chimeras [21]
and, remarkably, whole mice when injected into a tetraploid blastocyst [22]. As mESCs are amenable to genetic modification in vitro,
gene knockouts and knock-ins can be performed, including the
insertion of conditional and reporter alleles, multiplexed gene targeting, and genome-wide mutagenesis. Transgenic animals derived
via these routes have been instrumental for disease modeling and
the study of gene function [23–26].
Subsequent work focused on characterizing the molecular
pathways underpinning the maintenance of pluripotency in ESCs.
In 1988, leukemia inhibitory factor (LIF) was identified as the
principal component produced by feeder cells and can maintain
mESCs in their absence [27, 28], albeit less efficiently, as feeder
cells provide an additional attachment matrix as well as factors that
support the maintenance of pluripotency in ESCs in addition to
LIF [29, 30]. LIF activates STAT3 which feeds into the pluripotency network by upregulating the expression of pluripotency factors such as KLF4, Gbx2, and Tfcp2l1 [31–34]. Serum factors,
particularly bone morphogenetic proteins, stimulate the SMAD
signaling pathway and constrain lineage commitment by inducing
expression of inhibitor of differentiation (ID) proteins
[35, 36]. Building on these insights, in 2008, Ying et al. [37]
established a culture system to maintain ESCs in the absence of
LIF, serum, and feeders, with two small molecule inhibitors they
termed “2i.” The components of 2i are PD03 (PD0325901) and
CHIRON (CHIR99021) which modulate key pathways involved
in lineage commitment and pluripotency. PD03 is a MEK inhibitor
which blocks the auto-inductive effects of the FGF/ERK1/2 signaling cascade on differentiation [37], while CHIRON inhibits
GSK-3 which mimics the effects of canonical Wnt signaling and
thereby alleviates the repressive effects of TCF3 on pluripotency
genes.
Mouse Embryonic Stem Cell Culture
35
Although providing a well-defined milieu, culture in 2i anchors
mESCs in a so-called ground state of pluripotency [1], biasing
lineage commitment under differentiation-inducing conditions
[38]. Hence, most current mESC differentiation protocols still
use cells cultured under serum/LIF conditions in the presence of
feeder cells as a starting point. Albeit not chemically defined, this
system remains the gold standard for mouse pluripotent stem cell
culture. In this chapter, we describe the culture of mESCs under
serum/LIF conditions on a feeder layer. We provide stepwise protocols for (I) the generation of high-quality growth-inactivated
mouse embryonic fibroblast (iMEF) feeders, (II + III) the recovery
of and routine culture of mESCs, and (IV) a protocol for their
cryopreservation. The protocol described herein has broad applicability and is also compatible with the culture of induced pluripotent
stem cells derived from a variety of different cell types [39].
2
Materials
1. MEF culture medium: Dulbecco’s modified eagle medium
(DMEM) containing 10% fetal calf serum (FBS) (v/v); 1%
GlutaMAX™ Supplement (100) (v/v); 1% MEM nonessential amino acid solution (100) (v/v); 100 μM sodium pyruvate; 55 μM β-mercaptoethanol.
2. mESC culture medium: KnockOut™ DMEM containing 15%
FBS (v/v) (see Note 1); 1% GlutaMAX™ Supplement (100)
(v/v); 1% MEM nonessential amino acid solution (100)
(v/v); 55 μM β-mercaptoethanol; 1000 units/mL recombinant murine LIF.
3. Cryopreservation medium: FBS containing 10% DMSO (v/v).
4. Dulbecco’s phosphate-buffered saline (DPBS).
5. 0.25% Trypsin-EDTA (1), phenol red (380 mg/L EDTA,
2500 mg/L trypsin).
6. 175 cm2, angled neck, vented cap cell culture flasks.
7. 25 cm2, angled neck, vented cap cell culture flasks.
8. “Mr. Frosty” freezing containers.
9. 0.1% gelatine solution (w/v): Mix 1 g gelatine from porcine
skin with 1 L ultrapure water (milli-Q). Autoclave to dissolve.
10. 15 mL centrifuge tubes.
11. 50 mL centrifuge tubes.
12. EmbryoMax® Primary Mouse Embryo Fibroblasts, Strain
CF1, passage 1.
13. γ-Radiation source (e.g., Gammacell® 40 Exactor Low DoseRate Research Irradiator).
36
Jacob M. Paynter et al.
Fig. 1 Generation of iMEFs. (a) Schematic overview of the process. (b–d) Morphology of MEFs prior splitting at
the end of passages 1, 2, and 3, respectively. (e) Morphology of iMEFs 24 h after recovery from cryopreservation on a gelatine-coated surface. Scale bar: 25 μm
3
Methods
3.1 Generation
of iMEFs
High-quality iMEFs are crucial to propagate mESC in a highly
undifferentiated state. To generate feeder cells in large quantities,
passage 1 (p1) MEFs are expanded by serial passaging, mitotically
inactivated by irradiation, and cryopreserved for future use
(Fig. 1a). While this protocol makes use of commercially available
p1 MEFs, they can also be isolated de novo via dissection and
homogenization of embryonic day 13.5 mouse embryos as
described previously [40]. The procedure described in Subheading
3.1 is expected to yield between 2.4 and 3.6 108 iMEFs from the
expansion of a single embryo.
3.1.1 Recovery of MEFs
from Cryopreservation
(Passage 1)
1. Dispense 10 mL MEF medium into a 15 mL conical tube, and
place in a water bath heated to 37 C.
2. Retrieve cryovials from liquid nitrogen storage collectively
containing the MEFs from one embryo (~5 106) frozen
down at the first passaging event (as described in Ref. 35),
and place them in a 37 C water bath to thaw (see Note 2).
3. Once the contents of the vials have completely thawed, transfer
the contents to the pre-warmed MEF medium prepared in
step 1 of this section, and mix by gentle inversion.
4. Pellet the cells by centrifugation at 400 g for 5 min.
Mouse Embryonic Stem Cell Culture
37
5. Aspirate the supernatant from the pellet, and resuspend in
5 mL MEF medium by gentle pipetting.
6. Transfer the suspension in equal volumes to 2 175 cm2 culture flasks, and add additional MEF medium to achieve a
working culture volume of 20 mL per flask. Mix by gentle
pipetting.
7. Incubate the flasks at 37 C under low oxygen conditions (5%
O2, 7% CO2) for 24 h (see Note 3).
8. Perform a media change to remove any dead cells and traces of
DMSO: Aspirate culture medium and gently overlay the cells
with 20 mL fresh MEF medium (see Note 3).
9. Incubate at 37 C under low oxygen conditions for a further
24–48 h. The cells will require passaging upon reaching ~90%
confluence (Fig. 1b) (see Note 4).
3.1.2 Passage
2 (Expansion)
1. Aspirate culture media, and gently overlay the cells in each flask
with 15 mL DPBS, and aspirate to remove traces of culture
medium (see Note 5).
2. Dispense 3 mL Trypsin-EDTA solution into each flask, and
rock back and forth to ensure the solution is distributed evenly
over the cell layer.
3. Incubate the flasks at room temperature for 3–5 min.
4. Tap the flasks to dislodge the cells. Examine under a microscope to ensure all cells are fully detached (see Note 6).
5. Dispense 3 mL MEF medium into each flask to neutralize the
trypsin. Gently pipette up and down several times to homogenize the cell suspension.
6. Pool the contents of each flask into a single 50 mL
centrifuge tube.
7. Wash the surface of each flask with an additional 4 mL of MEF
medium to recover any remaining cells, and pool into the
50 mL tube.
8. Pellet the cells by centrifugation at 400 g for 5 min.
9. Aspirate the supernatant from the pellet, and resuspend in
16 mL MEF medium by gentle pipetting.
10. Perform a 1:4 split: Transfer the suspension (20–30 106 cells
total) in equal volumes into 8 175 cm2 culture flasks (see
Note 7), and add additional MEF medium to achieve a working culture volume of 20 mL. Mix by gentle pipetting.
11. Incubate the flasks at 37 C under low oxygen conditions (5%
O2, 7% CO2) for 48–72 h until the cells reach ~90% confluence
(Fig. 1c).
38
Jacob M. Paynter et al.
3.1.3 Passage
3 (Expansion)
1. Perform a 1:3 split: Passage cells as described previously in
Subheading 3.1.2, effectively expanding the 8 p2 flasks
into 24 175 cm2 culture flasks.
2. Incubate the flasks at 37 C under low oxygen conditions (5%
O2, 7% CO2) for 48–96 h until the cells reach ~100% confluence (Fig. 1d) (see Note 8).
3.1.4 Harvesting,
Irradiation,
and Cryopreservation
of MEFs
1. Trypsinize the cells as per Subheading 3.1.2, steps 2–5, and
transfer the contents of each flask into 5 50 mL centrifuge
tubes (see Note 9).
2. Wash the surface of each flask with an additional 4 mL of MEF
medium to recover any remaining cells, and pool into the
5 50 mL tubes.
3. Pellet the cells by centrifugation at 400 g for 5 min.
4. Aspirate the supernatant from the pellets. Resuspend and pool
the pellets in 30 mL MEF medium, and transfer to a single
50 mL tube. Determine cell number using an automated cell
counter slide or hemocytometer (see Note 10).
5. Place the tube in a research irradiator, and expose to γ radiation
at a dose rate of 1.0 Gy/min for 30 min.
6. Pellet the resulting iMEFs by centrifugation at 500 g for
5 min.
7. Aspirate the supernatant, and resuspend the pellet in cryopreservation medium at 1 107 cells/mL by gentle pipetting (see
Note 11).
8. Using a serological pipette, dispense 500 μL aliquots
(5 106 cells) into cryovials.
9. Place cryovials in a controlled rate freezer or freezing containers (“Mr. Frosty”) at 80 C for 24 h (see Note 12). After
freezing, transfer the vials to liquid N2 cryo-storage (see
Note 13).
3.2
mESC Culture
In this protocol, mESCs (see Note 14) are maintained as monolayer
cultures on gelatine-coated culture vessels that have been seeded
with iMEF feeder cells (Fig. 2a). iMEFs can be obtained commercially (e.g., EmbryoMax® Primary Mouse Embryo Fibroblasts,
Strain CF1, Irradiated, passage 3 (Merk Millipore)) or generated
as described in the previous section. Feeder cells provide an additional attachment matrix as well as factors that support the maintenance of pluripotency in ESCs in addition to serum and LIF
[29, 30].
3.2.1 Gelatine Coating
of Culture Vessels
and Seeding of MEFs
1. Overlay the surface of a 25 cm2 culture flask with 2 mL 0.1%
gelatine solution. Incubate at room temperature for at least
20 min (see Note 15).
Mouse Embryonic Stem Cell Culture
39
Fig. 2 mESC culture. (a) Schematic overview depicting recovery of mESC from cryopreservation and their
routine maintenance. (b–f) mESC recovery from cryopreservation and expected morphology. (b) iMEFs are
seeded 12–24 h before thawing of mESCs. (c) Thawed mESCs are seeded at a density of 2.5 104 cells/cm2
and should reach 70% confluence after 3 days (d–f). Scale bar: 25 μm
2. Retrieve a vial of irradiated MEFs from cryo-storage. Thaw and
resuspend as per Subheading 3.1.1, steps 1–5. Determine cell
number using an automated cell counter slide or
hemocytometer.
3. Aspirate the gelatine solution from the culture flask prepared in
step 1 of this section. Transfer a volume of suspension containing 5 105 iMEF cells (2 104 cells/cm2) into the flask, and
add additional MEF medium to achieve a working culture
volume of 3 mL (see Note 16).
4. Incubate the flask at 37 C for 12–24 h to allow the feeders to
attach. The cells should cover 70–100% of the flask (Fig. 2b).
3.2.2 Thawing of mESCs
1. Retrieve a vial containing ~1 106 mESCs from cryo-storage.
Thaw and pellet as per Subheading 3.1.1, steps 1–4.
2. Aspirate the supernatant from the pellet, and resuspend in
1 mL mESC medium by gentle pipetting. Determine cell number using an automated cell counter slide or hemocytometer.
3. Aspirate MEF medium from the gelatine-coated flask with
feeders prepared in Subheading 3.2.1. Transfer a volume of
mESC suspension containing 6.25 105 cells
40
Jacob M. Paynter et al.
(2.5 104 cells/cm2) into the flask (Fig. 2c), and add additional mESC medium to achieve a final culture volume of
3 mL. Mix by gentle pipetting.
4. Incubate the flask at 37 C for 24 h under atmospheric oxygen
and 5% CO2.
5. Perform a media change to remove any dead cells and traces of
DMSO: Aspirate culture medium and gently overlay the cells
with 3 mL fresh mESC medium.
6. Incubate at 37 C for a further 48–72 h, changing medium
after 48 h. The cells will require passaging upon reaching ~70%
confluence (Fig. 2d–f) (see Note 17).
3.2.3 Routine Passaging
of mESCs
1. Prepare a fresh gelatine-coated 25 cm2 flask seeded with iMEFs
as per Subheading 3.2.1.
2. Aspirate culture media from the mESC culture vessel (25 cm2
flask), gently overlay with 2 mL DPBS, and aspirate to remove
traces of culture medium.
3. Dispense 1 mL Trypsin-EDTA solution into each flask, and
rock back and forth to ensure the solution is distributed evenly
over the cell layer.
4. Incubate the flasks at room temperature for 3–5 min.
5. Tap the flasks to dislodge the cells. Examine under a microscope to ensure all cells are fully detached.
6. Dispense 1 mL mESC medium into each flask to neutralize the
trypsin. Gently pipette up and down several times to homogenize the cell suspension.
7. Transfer the contents of the flask into a 15 mL centrifuge tube.
8. Pellet the cells by centrifugation at 400 g for 5 min.
9. Aspirate the supernatant from the pellet, and resuspend in
5 mL mESC medium by gentle pipetting (see Note 18). Determine cell number using an automated cell counter slide or
hemocytometer. A 70% confluent 25 cm2 flask should yield
between 7 106 and 1 107 cells.
10. Aspirate MEF medium from the gelatine-coated flask with
feeders prepared in step 1 of this section. Transfer a volume
of mESC suspension containing 1.25–2.5 105 cells
(0.5–1 104 cells/cm2) into the flask, and add additional
mESC medium to achieve a final culture volume of 3 mL (see
Note 19). Mix by gentle pipetting.
11. Incubate at 37 C for 72 h under atmospheric oxygen and 5%
CO2 until cultures reach ~70% confluence, changing medium
after 48 h.
Mouse Embryonic Stem Cell Culture
3.2.4 Cryopreservation
of mESCs
41
1. After counting, pellet mESCs obtained in Subheading 3.2.3,
step 9 by centrifugation at 500 g for 5 min.
2. Aspirate the supernatant, and resuspend the pellet in cryopreservation medium at 1 106 cells/mL by gentle pipetting. A
70% confluent 25 cm2 should yield 7–10 cryovials.
3. Using a serological pipette, dispense 1 mL aliquots
(1 106 cells) into cryovials.
4. Place cryovials in a controlled rate freezer or (“Mr. Frosty”)
freezing containers at 80 C for 24 h. After freezing, transfer
the vials to liquid N2 cryo-storage.
4
Notes
1. Certain batches of FCS can be detrimental to mESC cultures.
Hence, we recommend using embryonic stem cell-qualified
FCS which can be purchased from several vendors (e.g.,
Thermo Fisher, Merck, Applied StemCell). If this is undesirable due to financial constraints, batch testing of sera from
other vendors can be performed. For batch testing, we recommend culturing mESCs for several passages (~5) in the alternative sera and to only use batches that are able to maintain the
typical dome-shaped morphology and growth rate of mESCs.
2. Cells should be thawed as quickly as possible to maximize
recovery. This can be facilitated by using a circulating water
bath and periodically agitating the vial.
3. We recommend the use of low oxygen incubators for MEF
expansions. MEFs cultured in a low oxygen environment
show higher proliferation rates and delayed senescence compared to those cultured under atmospheric oxygen [41], thus
giving rise to higher quality feeders.
4. MEF cultures should not be allowed to exceed 90% confluency,
as this can induce growth inhibition and result in poor
expansion.
5. When changing medium or washing with PBS, never dispense
liquid directly onto the cells, as this can compromise the monolayer. Instead, eject gently down the side of the flask.
6. Prolonged trypsin exposure can compromise cell viability. We
do not advise exposing cells to trypsin for much longer than
5 min.
7. When passaging MEFs, culture flasks may be reused to reduce
financial costs.
8. The growth rate of MEFs slows down at later passages due to
the onset of senescence, and cultures may take longer to reach
confluence. Senescent MEFs have a flattened, spread out
42
Jacob M. Paynter et al.
(“fried egg”) morphology. Cultures dominated by overtly
senescent cells give rise to poor-quality feeders; hence we advise
not to expand MEFs beyond passage 3. If culturing for longer
than 72 h, cells should receive a media change every 48 h.
9. When handling high numbers of flasks, we recommend dividing the labor across subsets to avoid cells drying out during the
washing steps.
10. Due to the high cell density of the suspension at this stage, we
recommend taking a 20 μL aliquot of the concentrated suspension and diluting it 1:10 in MEF media before counting to
determine the cell number more accurately.
11. It is advisable to minimize exposure of cells to DMSO at
subfreezing temperatures due to its toxicity. Cell viability can
be increased by working quickly and keeping the cells at a low
temperature. Feeders should be resuspended in prechilled
cryopreservation medium and aliquoted promptly into prechilled cryovials.
12. A controlled cooling rate of 1 C per minute is optimal for
maximizing cell viability during freezing. While the use of “Mr.
Frosty” freezing containers is acceptable for cryopreserving
iMEFs, we recommend the use of a controlled rate freezer, as
the freezing process induces a localized spike in temperature. A
controlled rate freezer compensates for this and achieves a
more uniform cooling rate resulting in superior cell viability
after thawing.
13. High-quality feeders should have a high recovery rate after
thawing (>70%) and adopt a spindle-shaped morphology
(Fig. 1e).
14. The procedures outlined in this Methods chapter were established with R1 mESCs. While this protocol is compatible with
mESC lines from other mouse strains, growth rates may vary
slightly. Therefore, seeding densities may need to be optimized
for other mESC lines.
15. Culture vessels can be scaled up or down as needed, e.g., use
6 mL of 0.1% gelatine solution for a 75 cm2 flask.
16. Although less cost-effective, feeders can also be cultured in
mESC medium for convenience.
17. Culturing cells beyond 70% confluence may lead to spontaneous differentiation and poor cell viability. Always ensure that
the morphology of the colonies is predominantly dome-shaped
before using the cells for experiments. Immunostaining for
pluripotency markers such as Oct4, Nanog, Sox2, and SSEA1 can be used to indicate the quality of mESC cultures (Fig. 3).
Immunostaining can be performed as described previously
[42] using primary antibodies against OCT4 (mouse IgG2b,
Mouse Embryonic Stem Cell Culture
43
Fig. 3 Assessment of differentiation status of mESC cultures. mESC colony stained for pluripotency markers
(a) SSEA1 and (b) OCT4; (c) the DNA intercalating dye 40 ,6-diamidino-2-phenylindole (DAPI) was used to
visualize cell nuclei; Panel d depicts a bright-field image of the mESC colony. Scale bar: 25 μm
1:100 dilution, Clone: C-10, Santa Cruz Biotechnology) and
SSEA-1 (mouse IgM, 1:200 dilution, Clone: MC-480,
DSHB); secondary conjugated antibodies goat anti-mouse
IgG2b-AF 488 (1:400 dilution, Thermo Fisher Scientific)
and goat anti-mouse IgM-AF 555 (1:400 dilution, Thermo
Fisher Scientific); and nuclear stain 40 ,6-diamidino-2-phenylindole, dihydrochloride (DAPI) (1:1000 dilution, Invitrogen).
Cultures with excessive spontaneous differentiation can be
rescued by FACS isolation of SSEA-1 and EPCAM doublepositive cells and their use for re-culture [40, 43]. A more
stringent assay for pluripotency is the teratoma formation
assay entailing subcutaneous injection of mouse pluripotent
stem cells into the flanks of immune compromised mice to
assess their in vivo differentiation potential into derivates of
all three germ layers [44, 45].
18. Most differentiation protocols require the use of mESCs in the
absence of contaminating feeders. mESC suspensions (after
Trypsin/EDTA dissociation) can be depleted of feeders by
transferring the 5 mL suspensions obtained in Subheading
3.2.1, step 9 onto a non-gelatine-coated 25 cm2 culture flask
and incubating at 37 C for 30 min. After this period, the
feeders will have attached to the plate, while the mESCs remain
in suspension. The supernatant containing the mESCs can be
collected and used for downstream differentiation applications
as detailed previously [46].
19. Experienced users can perform a 1:10–1:20 split which should
approximate this seeding density.
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Chapter 4
Production of High-Titer Lentiviral Particles for Stable
Genetic Modification of Mammalian Cells
Michael R. Larcombe, Jan Manent, Joseph Chen, Ketan Mishra,
Xiaodong Liu, and Christian M. Nefzger
Abstract
Lentiviral gene transfer technologies exploit the natural efficiency of viral transduction to integrate exogenous genes into mammalian cells. This provides a simple research tool for inducing transgene expression or
endogenous gene knockdown in both dividing and nondividing cells. This chapter describes an improved
protocol for polyethylenimine (PEI)-mediated multi-plasmid transfection and polyethylene glycol (PEG)
precipitation to generate and concentrate lentiviral vectors.
Key words PEI transfection, PEG precipitation, Titration, Gene transfer
1
Introduction
Effective delivery and expression of exogenous genes in mammalian
cells is essential for the study of gene function. Viral transfer technologies are routinely used for transient and integrating gene delivery in vitro and, once biosafety concerns are addressed, have a vast
potential for clinical applications [1–3]. Unlike the transient expression achieved with adenoviral vehicles, lentiviral and retroviral vectors allow stable transgene integration for sustained and heritable
gene expression. This is particularly useful for generating transgenic
animals, reprogramming fibroblasts into induced pluripotent stem
cells (iPSCs), and creating stable cell lines overexpressing or silencing genes via RNA interference [4].
While both lentiviral and retroviral gene transfer methods integrate transgenes into the genome of targeted cells for continued
expression, lentiviruses transduce both replicating and
non-replicating cells, integrate away from cellular promoters,
allow for a larger genomic payload, and maintain transgene
Michael R. Larcombe, Jan Manent, and Joseph Chen contributed equally to this work.
Ivan Bertoncello (ed.), Mouse Cell Culture: Methods and Protocols, Methods in Molecular Biology, vol. 1940,
https://doi.org/10.1007/978-1-4939-9086-3_4, © Springer Science+Business Media, LLC, part of Springer Nature 2019
47
48
Michael R. Larcombe et al.
expression in pluripotent cells [5–9]. Thus, lentiviruses are a particularly widely used and a popular tool for the stable genetic modification of mammalian cells. Furthermore, advances have been made
to minimize risk and improve delivery efficiency via the design of
new generation lentiviral vectors [10]. These systems prevent the
generation of replication competent virus through deletion of nonessential viral genome components and separation of the remaining
elements. As such the second-generation system used in this study
requires three separate vectors encoding for the transgene (called
“transfer vector”), replication genes (including the rev transactivator), and envelope genes. Omission of unnecessary virulence factors
and isolation of the remaining components further reduce the risk
of homologous recombination that could lead to the generation of
unwanted replicating virus [11, 12]. Modern vector designs
improve infectivity of the generated particles by substitution of
viral coating proteins within the envelope vector for a broader
(vesicular stomatitis Indiana virus G protein [VSV-G], as used in
this protocol) [13, 14] or more specified tropism (human parainfluenza virus type 3 [HPIV3] for lung epithelial cells) [15, 16].
Numerous methods for producing lentiviral particles have been
established, predominantly using transient co-transfection of the
desired transfer vector with the accessory plasmids [17, 18]. This
can be achieved with a variety of reagents utilizing lipid-based,
polymer-based, or naked DNA delivery [19]. Nevertheless, price,
efficiency, scalability, and simplicity must be accounted for when
selecting transfection technique and reagents. Commercial products such as Lipofectamine and newer versions thereof offer a
simple and efficient way to achieve effective plasmid transfection;
however, especially for large-scale experiments, they can be cost
intensive. Since viral particle production is generally achieved using
highly transfectable human embryonic kidney 293T (HEK293T)
cells, lipofection-based approaches can be substituted with the
more cost-effective calcium phosphate (Ca-phosphate) or polyethylenimine (PEI) transfection systems. Ca-phosphate and PEI
methods are also reasonably simple and capable of producing high
viral titers from HEK293T cells [15, 20–22]. However transfecting
with Ca-phosphate is very sensitive to fluctuations in pH [23], and
therefore the resulting titers can differ largely between experiments.
Since generation of high viral titers is dependent on efficient transfection, we consider the PEI-based method more reliable for consistent results. PEI transfection operates by condensing DNA into
positively charged particles for delivery across the cell membrane.
DNA is then released in the cytoplasm and incorporated into the
nucleus during cell division for temporary expression [24], followed by the secretion of lentiviral particles into the growth
medium. The resulting supernatant can then be collected and
used for direct transduction of target cells or concentrated to
achieve higher titer viral preparations. Protocols for lentiviral
Lentiviral Particle Production and Concentration
49
concentration typically require expensive ultrafiltration units and
lengthy periods of ultracentrifugation [15, 25–27]. For laboratories without access to this high-end equipment and due to the
ease of use, polyethylene glycol (PEG) precipitation offers a cheap
and simple alternative to concentrate large volumes of supernatant
and efficiently recover the viral particles [25, 28]. PEG is a highly
hydrophilic polymer composed of repeating subunits of ethylene
oxide and is commercially available in varying molecular weights.
Generally, higher molecular weight PEGs are more efficient for
precipitation compared to lower molecular weight PEGS
[29]. Therefore we are using PEG (8000 Da) that has a relatively
high molecular weight and is therefore very effective as a crowding
agent in aqueous solution. PEG separates the viral particles from
the aqueous medium and forces them to aggregate through a
process called “steric exclusion” [30] which is supported by high
levels of salt such as NaCl, through a “salting-out” mechanism
[31]. This allows the virus to be pelleted with a simple benchtop
centrifuge without the need for ultracentrifugation [32–35].
Therefore, in this chapter, we describe an improved protocol
for the production and concentration of replication-deficient hightiter lentiviruses using PEI transfection and concentration by PEG
precipitation. In the context of this protocol, we also describe the
routine maintenance of the HEK293T producer cell line, a method
for determining viral titers (for viral inserts with and without a
fluorescent reporter gene) and the use of the viral concentrates to
infect a target cell type of interest.
2
Materials
2.1 Lentiviral
Production
and Titration
1. Lenti-X™ 293T cell line (HEK293T) (Clontech, 632180) or
293T cell line (ATCC® CRL-3216™).
2. Complete growth medium (for 293T cells and MEFs):
Dulbecco’s modified eagle medium (DMEM), containing
10% fetal calf serum (FBS) (v/v); 1% GlutaMAX™ Supplement
(100) (v/v); 1% MEM nonessential amino acid solution
(100) (v/v); 1% penicillin/streptomycin (100) (v/v);
100 μM sodium pyruvate; 55 μM β-mercaptoethanol.
3. Viral production culture medium: Advanced DMEM containing (Gibco®, 12491015) 2% FBS (v/v) (see Note 1); 1% GlutaMAX™ Supplement (100) (v/v); 1% MEM nonessential
amino acid solution (100) (v/v); 1% penicillin/streptomycin
(100) (v/v).
4. 15 mL centrifuge tubes.
5. Cryopreservation medium: FBS containing 10% DMSO (v/v).
6. Dulbecco’s phosphate-buffered saline (DPBS).
50
Michael R. Larcombe et al.
7. 0.25% trypsin-EDTA (1), phenol red (380 mg/L EDTA,
2500 mg/L trypsin).
8. 175 cm2, angled neck, vented cap cell culture flasks (T175).
9. 50 mL centrifuge tubes.
10. “Mr. Frosty” freezing container.
11. Linear polyethylenimine 25,000.
12. BSA fraction V (7.5%) (BSA).
13. Virkon® disinfectant cleaner.
14. UltraPure distilled water.
15. PAX2 plasmid (psPAX2 was a gift from Didier Trono
(Addgene, #12260)).
16. MD2G plasmid (pMD2.G was a gift from Didier Trono
(Addgene, #12259)).
17. OKSM plasmid (Merck Millipore, SCR512).
18. rtTA-GFP plasmid (designed and ordered from vector builder
[Cyagen Biosciences]; lentiviral plasmid with a EF1A promoter
driving the m2rtTA gene, followed by an internal ribosome
entry site and an eGFP reporter (LV-EF1A-m2rtTA-IRESeGFP)).
19. 0.45 μm, HV Durapore® membrane filter (Merck Millipore).
20. Polyethylene glycol (PEG) 8000.
21. 5M NaCl in dH2O, filtered with 0.22 μm.
22. Doxycycline hyclate diluted to 2 mg/mL in dH2O (1000
stock).
23. Polybrene infection/transfection reagent 10 mg/mL stock
(used at 1:1700 dilution).
2.2 Immunofluorescence and Viral
Titration Analysis
(See Note 18)
1. Paraformaldehyde (PFA).
2. Triton-X.
3. DAPI: 40 ,6-diamidino-2-phenylindole, dihydrochloride.
4. Mouse anti-Oct4 antibody (Santa Cruz Biotechnology,
sc-5279).
5. Goat anti-mouse IgG Alexa Fluor 555 (Life Technologies®,
A21422).
3
Methods
3.1 Recovery from
Cryopreservation
and Routine Culture
of HEK293T Cells
1. Collect cryovial containing 4–5 106 HEK293T cells from
liquid nitrogen storage, and thaw quickly in a 37 C water bath.
2. Transfer cells to a 15 mL conical tube along with 10 mL of
pre-warmed complete growth medium.
Lentiviral Particle Production and Concentration
51
3. Centrifuge at 500 g for 5 min, and then aspirate supernatant
containing toxic DMSO without disturbing cell pellet.
4. Resuspend cells in 4 mL of growth medium by gentle pipetting, and transfer 3.5 106 cells into a 175 cm2 flask (T175)
made to 20 mL with growth medium.
5. Incubate at 37 C in a 5% CO2 incubator. Replace media every
2–3 days (as required) with 20 mL of fresh growth medium
during routine culture until cells reach 80% confluence (see
Note 1).
6. To harvest cells, firstly remove the spent growth medium, and
then wash cells with 10 mL PBS. Apply DPBS to the side of the
flask, and gently tilt to spread, trying not to disturb the weakly
adherent cells.
7. Aspirate DPBS, and evenly distribute 5 mL trypsin-EDTA over
the cells to dissociate.
8. Incubate the cells at room temperature for 3–5 min, using the
microscope to verify when >90% of the cells have detached.
9. Neutralize the trypsin by adding 10 mL of growth medium.
Pipet up and down several times over the cell culture surface to
separate and collect remaining cells.
10. Transfer cells to a 50 mL centrifuge tube and pellet at 500 g
for 5 min.
11. Resuspend the cells in 5 mL of complete growth medium, and
remove a sample for counting.
12. In general, 80–90% confluent flasks can be passaged at a split
ratio of 1:5 up to 1:20. Depending on the split ratio, the new
flasks will become confluent in a 2–4-day time frame.
13. Create frozen stocks by resuspending the cells in cryopreservation medium at 2–6 106 cells/mL, and dispense 1 mL
aliquots into cryovials. Use a controlled rate freezer or “Mr.
Frosty” freezing container at 80 C for 24 h, and transfer
frozen cells to liquid nitrogen for long-term storage.
3.2 PEI Transfection
(See Note 2)
and Collection
of Primary Viral
Supernatant
1. Thaw an aliquot of 1 mg/mL PEI (see Note 3).
2. The day before transfection, plate HEK293T cells (that have
been passaged at least once after recovery from cryopreservation) in complete growth medium at a density of 1.5 107 to
2 107 cells per T175 flask (8.5 104 to 1.15 105 cells/
cm2). Depending on the growth rate of your HEK293T culture, adjust to have healthy, 80% confluent cultures for transfection the following day (Fig. 1) (see Note 4).
3. The next day in the late afternoon, 1 h prior to transfection,
change complete growth medium to viral production culture
medium (20 mL per T175 flask).
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Michael R. Larcombe et al.
Fig. 1 Schematic overview of viral particle production via PEI transfection
Table 1
Transfection mix for second-generation lentivirus production in HEK293T cells
T175
T75
T25
Vector
Concentration Ratios μg DNA Volume μg DNA Volume
Transfer DNA
100 ng/μL
3
15
150 μL 6.3
63 μL
2.1
21 μL
psPAX2
100 ng/μL
2
10
100 μL 4.2
42 μL
1.4
14 μL
pMD2.G
100 ng/μL
1
5
50 μL
21 μL
0.7
7 μL
120
120 μL 50.4
50.4 μL
16.8
16.8 μL
H2O
980 μL
423.6 μL
141.2 μL
Total volume
1.4 mL
0.6 mL
0.2 mL
PEI (4 μg/μg DNA) 1 mg/mL
2.1
μg DNA Volume
DNA concentrations are arbitrarily set at 100 ng/μL for the purpose of this example
If transfecting more than one T175 flask, the values in this table can be scaled up accordingly
4. Prepare transfection mix as shown in Table 1: per T175 to
transfect, mix 15 μg transfer DNA, 10 μg psPAX2, 5 μg
pMD2.G, and 120 μg PEI in water to a final volume of
1.4 mL (see Note 5).
Lentiviral Particle Production and Concentration
53
5. Vortex solution for 10 s, and incubate at room temperature for
15 min.
6. Pipet up and down gently 2–3 times, and add the solution to
the cells dropwise (Fig. 1).
7. Swirl flask gently to evenly distribute the DNA mix over the
cells and return to incubator.
8. Incubate overnight at 37 C, 5% CO2.
9. Early the next morning, discard transfection medium into
strong bleach or 1% Virkon® solution, and replace with
fresh viral production culture medium (Fig. 1).
10. 24 h later, collect supernatant for the first time, and replace
with fresh viral production culture medium. Keep supernatant
at 4 C if not processed right away (Fig. 1) (see Note 6).
11. 24 h later (i.e., 48 h after removal of transfection complexes),
collect supernatant for a second time. The flasks with any
remaining cells can be discarded at this point (Fig. 1).
12. Proceed with concentration.
3.3 Viral
Concentration
1. Filter the harvested medium through a 0.45 μm membrane,
and take note of total volume (Fig. 2) (see Note 7).
2. Adjust NaCl concentration to 400 mM with a 5 M NaCl stock
solution. This is achieved by adding a 1/17 volume of
5M NaCl to viral production culture medium (i.e for 10 mL
of viral supernatant, add 588 μL of 5M NaCl).
3. Add 50% PEG solution to a final concentration of 8.5% (see
Note 8).
4. Mix well and transfer mixture into 50 mL Falcon tubes. Make
sure to have balanced volumes in the tubes for centrifugation in
step 6 (Fig. 2).
5. Incubate the tubes at 4 C or on ice for 5 h, with regular
agitation (see Note 9).
6. Spin samples for 1.5 h at >4000 g in a centrifuge pre-cooled
to 4 C (see Note 10).
7. Gently discard supernatant (without disturbing viral pellet) in
bleach or 1% Virkon® solution, and centrifuge empty tube
again for 5 min (Fig. 2).
8. Gently remove any remaining supernatant with a P1000 pipette
without disturbing pellet (see Fig. 2 for picture of the pellet).
9. We recommend to resuspend the pellet in DPBS containing 1%
BSA (v/v) in a volume corresponding to 1/100th of the
amount of primary supernatant used for concentration (e.g.,
resuspend the pellet resulting from the concentration of 30 mL
supernatant in a volume of 300 μL; see Note 11).
54
Michael R. Larcombe et al.
Fig. 2 Schematic overview of viral particle concentration with PEG
10. Aliquot into 20–30 μL aliquots (see Note 12).
11. Freeze at 80 C.
3.4
Titration
3.4.1 Viral Titration of a
Lentiviral Vector Carrying
a Fluorescent Reporter
1. Thaw and recover a vial of mouse embryonic fibroblasts
(MEFs) from DMSO (see Note 13) into complete growth
media.
2. Seed MEFs in 12 wells (9 for rtTA-GFP titering and 5 control
wells (3 control wells and 2 wells for counting)) in a 24-well
plate format at 1 104 cells/cm2 at least 12–24 h before viral
transduction (Fig. 3a, c).
3. Count the cells of two control wells to obtain accurate cell
count at the time of transduction.
4. Thaw and aliquot rtTA-GFP viral concentrate (at 1:1000,
1:10000, 1:100000 dilutions; see Note 14) into complete
growth media containing polybrene (1:1700 dilution of
stock) in 1000 μL volumes.
5. Remove media from the wells previously seeded with MEFs,
and add 500 μL media of the respective serial dilutions onto
cells (Fig. 3a, c).
6. Spin plates at 750 g for 60 min at room temperature to
increase the efficiency of transduction.
7. The following day, replace media containing viral particles with
fresh complete growth media.
8. 72 h after transduction, harvest each of the wells by dissociating cells with appropriate dissociation reagent (0.25% Trypsin,
EDTA, etc.) for 5 min. Resuspend cells with FACS buffer (PBS
w/2% FBS) supplemented with DAPI (1 μg/mL), and transfer
cells into a FACS tube to determine the percentage of GFP
Lentiviral Particle Production and Concentration
55
Fig. 3 Titration of viral concentrates. (a) Schematic overview of rtTA-GFP viral titration. (b) FACS analysis of
rtTA-GFP viral titration. (c) Schematic overview of OKSM viral titration. (d) Immunofluorescence images for
GFP (green) and Oct4 (red) in MEFs transduced with OKSM and rtTA-GFP viruses after 48 h and quantification
of Oct4þ cells within the GFPþ population. All nuclei were counterstained with DAPI (blue). Scale
bar ¼ 25 μm
expressing cells by flow cytometry for the m2rtTA-GFP construct (Fig. 3b).
9. Calculate titers of viral concentrate using the following formula: (% of GFP+ cells [between 1% and 25%] number of
cells at the time of transduction dilution factor)/volume of
media for transduction (in mL). Titers are usually indicated as
transducing units per mL (TU/mL) (see Notes 15 and 16 for
example calculation).
56
Michael R. Larcombe et al.
3.4.2 Viral Titration of a
Lentiviral Vector Without
a GFP Fluorescent Marker
The titration of the OKSM virus (which does not have a fluorescent
reporter) follows a similar pattern as described in steps
3.4.1. steps 1–9 for the m2rtTA-GFP virus. However, there are
noteworthy differences. In particular, for calculating OKSM viral
titers, an immunofluorescence staining is required to determine the
percentage of cells expressing exogenous Oct4, Klf4, Sox2, or
C-Myc (Fig. 3c, d). Furthermore as the OKSM constructs is inducible, the titer determination will have to occur in the presence of
excess m2rtTA-GFP virus and doxycycline (2 μg/mL) to enable
OKSM expression (see Notes 17 and 18 for detailed information as
well as Fig. 3c, d).
3.5 Viral
Transduction of Target
Cell Type
1. MEFs (or any other cell type that has ideally also been used to
determine the viral titers) can be used for transduction in an
experimental context.
2. For infection seed MEFs at 1 104 cells per cm2 as in
Subheading 3.4.1., step 2, (albeit in a 6-well format in 2 mL
of media or scale to other well formats) 12–24 h before
transduction.
3. Prepare transduction mix as in Subheading 3.4.1., step 4 with
outlined modifications: it is important for the experimenter to
decide at what multiplicity of infection (MOI; see Note 17) to
perform transduction. As a reference if the experimenter decides for an MOI ¼ 1, only ~60% of cells will become transduced
as some cells become infected with more than one virus. Conversely infection with an MOI of 10 will result in infection of
>85–90% of cells with an average of 10 viral integrants per cell.
An example of how to calculate the amount of viral concentrate
to add to a 6-well with a set number of cells for a specific target
MOI is provided in Note 19.
4. Perform spin inoculation as described in Subheading
3.4.1. steps 5–7, using 2 mL of transduction mix per 6-well.
Please note that it will take 48–72 h before gene products
introduced with the viral inserts can be detected in the infected
cells.
4
Notes
1. HEK293T cells undergo contact inhibition of growth if
allowed to become confluent. This must be avoided as a key
parameter for efficient viral production is healthy, exponentially
growing cells. Gently move the freshly seeded flask back and
forth, left and right to evenly distribute the cells throughout
the flask. Try to recover low-passage stock (<25), if cells are
not growing well.
Lentiviral Particle Production and Concentration
57
2. Prior to undertaking any work with lentiviral vectors, ensure all
safety specifications provided by your institution or governing
body are upheld. All work must be completed by trained personnel inside class 2 or higher biosafety cabinets with all equipment and spills correctly decontaminated. As with all other
tissue cultures, it is essential to work in a sterile environment
and clean all items within the cabinet with 70% ethanol.
3. Create a stock of PEI at 1 mg/mL following manufacturer’s
instruction. Briefly, dissolve PEI powder in endotoxin-free
dH2O, adjust pH to 7.0 with HCl, and allow time to
completely dissolve. Adjust the total volume to a final concentration of 1 mg/mL, and filter through a 0.22 μm membrane
to remove undissolved PEI. Aliquot and store at 20 C. Each
thawed aliquot can be stored at 4 C for 2 months before
transfection efficiency is affected. If a sample precipitates, heat
to 37 C and vortex thoroughly before use. Discard and take a
new aliquot if necessary.
4. Cells should be in their exponential growth phase for transfection as indicated in Note 1. If the cells are too confluent, they
will not be transfected or produce virus less efficiently. This
protocol may be scaled according to your viral production
needs.
5. This protocol is compatible with both second- and thirdgeneration lentiviral packaging systems. If using a thirdgeneration system, we recommend a plasmid ratio of transfer
vector:VSV:RRE:REV at 3:1:2:2 while maintaining a mass ratio
of 4:1 of PEI to total DNA [36].
6. For transfer vectors carrying a fluorescent marker, transfected
HEK293T cells can be visualized under a fluorescent microscope to determine the extent of transfection. But be mindful
that strong agitation of the flask at this stage might result in
HEK293T cell detaching. Expect >70% of HEK293T cells to
express the fluorescent marker.
7. Using filters with a smaller pore size may result in retention of
viral particles.
8. Resuspend PEG powder in water, 50% weight/volume, and
filter (0.45 μm). The solution will be very viscous, so we
recommend resuspending overnight at 37 C with agitation.
9. We recommend constant mixing using a rotating wheel or
rolling table at 4 C; if this isn’t available, you can invert
tubes on ice at half-hourly intervals.
10. We have successfully been using a Heraeus Multifuge X3R
centrifuge (Thermo Fisher) at Vmax of 4700 rpm/4800 g.
58
Michael R. Larcombe et al.
11. The PEG/viral pellet is quite sticky. It helps to resuspend the
pellet and aliquot at room temperature but return to ice or
freeze down immediately afterward.
12. Depending on your titer and experimental requirements, you
may want to adjust the aliquot size in the next viral preparation.
Avoid freeze thaw cycles as the amount of infectious particles
drops dramatically with each additional freeze thaw cycle (ideally only have one freeze thaw cycle).
13. The transduction efficiency of a lentiviral vector varies between
cell types; therefore it is necessary to titrate against the cell type
intended for use in the planned experiments. Seed at least
duplicate wells for each viral titration (for five concentrations,
seed at least ten wells of cells), and include two control wells
that are not transduced with virus. MEFs are only used as an
example cell type in the context of this manuscript and should
be substituted for the experimenters’ cell type of need.
14. You may prepare these solutions in an empty 24-well plate by
adding viral concentrate into a 24 well at a 1:100 or 1:1000
dilution and then perform ten-fold serial dilutions into neighboring wells containing growth media supplemented with
polybrene.
15. Use the dilutions that give values in the range of 1–20% GFP
cells to calculate the MOI to avoid complications arising from
cells receiving multiple insertions.
16. Example calculation of viral titer:
(% of GFP+ cells [e.g., 5%, express as 0.05] number of cells at
the time of transduction [e.g., 2 104] dilution factor
[100000])/volume
of
media
for
transduction
(0.5 mL) ¼ 2 108 transduction units (TU)/mL
17. The multiplicity of infection or mean occurrence of infection
(MOI) is defined as the theoretical average number of viral
integrants per target cell.
18. To determine the titer of OKSM viruses, seed MEFs in 24-well
plates (Fig. 3c) at 1 104 cells/cm2 12–24 h before viral
transduction. Aliquot rtTA-GFP virus into complete growth
media containing polybrene (1:1700 dilution) at MOI ¼ 10.
(The MOI of 10 for rtTA-GFP is chosen to ensure most of the
cells are expressing the transactivator that is critical for doxycycline induction of OKSM expression (see Note 19 for example
calculation). As an alternative, OKSM lentivirus can be titered
using an rtTA-expressing stable cell line.) Separate out the
rtTA-GFP containing media into 1000 μL volume aliquots,
and establish 1:100, 1:1000, 1:10000, and 1:100000 dilutions
of OKSM virus. We recommend using at least four 10 serial
dilutions ranging from 1:100 to 1:100000 of OKSM virus to
get a more accurate titration of the virus. In our experience, a
Lentiviral Particle Production and Concentration
59
dilution factor of up to 1:1000000 can be required to determine the transducing units of high-titer viral preparations. Add
500 μL of media into respective wells (as indicated in Fig. 3c)
followed by spin inoculation as per Subheading 4.4.1.6. On
the following day, remove media from wells, and replace with
fresh media containing doxycycline (2 μg/mL). After 48–72 h,
fix cells and perform immunofluorescence on cells to determine infective units of OKSM virus preparation. Further
details on performing immunofluorescence can be referred
from Nefzger et al. [37] and Chen et al. [38]. We recommend
using an Alexa Fluor 555 secondary antibody as it does not
interfere with the 488 nm (GFP) channel or the DAPI channel
when analyzing expression of viral titration and a primary
antibody against Oct4 (see Subheading 2). Calculate the subset
of GFPþ cells that are positive for Oct4, Klf4, Sox2, or C-Myc
(rtTA-GFP transduced cells) to determine the percentage of
transduction (Fig. 3d). We perform cell quantification using
the particle analysis option of the ImageJ software (http://rsb.
info.nih.gov/ij/). We recommend counting cells from multiple images per replicate well to give a more accurate estimation
of the transduction rate of the OKSM virus. It is recommended
to take images from at least four fields of view per well for
analysis.
19. Example calculation for transduction at MOI ¼ 10:
Number of cells to be infected 2 104 MOI ½10
¼ TU required to transduce cells at MOI ¼ 10 2 105 TU
Volume of viral aliquot to use for transduction of 2 104 cells
at MOI ¼10
2 105 TU ðTU required to transduce cells at MOI ¼ 10Þ=
2 108 =mL ðTiter of viral concentrateÞ ¼ 0:001 mL or 1 μL
Be advised that MOIs that are higher than 50 can lead to varying
degrees of cell death due to cytotoxicity.
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Chapter 5
Generation of Mouse-Induced Pluripotent Stem Cells
by Lentiviral Transduction
Xiaodong Liu, Joseph Chen, Jaber Firas, Jacob M. Paynter,
Christian M. Nefzger, and Jose M. Polo
Abstract
Terminally differentiated somatic cells can be reprogrammed into an embryonic stem cell-like state by the
forced expression of four transcription factors: Oct4, Klf4, Sox2, and c-Myc (OKSM). These so-called
induced pluripotent stem (iPS) cells can give rise to any cell type of the body and thus have tremendous
potential for many applications in research and regenerative medicine. Herein, we describe (1) a protocol
for the generation of iPS cells from mouse embryonic fibroblasts (MEFs) using a doxycycline (Dox)inducible lentiviral transduction system; (2) the derivation of clonal iPS cell lines; and (3) the characterization of the pluripotent potential of iPS cell lines using alkaline phosphatase staining, flow cytometry, and the
teratoma formation assays.
Key words Mouse-induced pluripotent stem cells, Fibroblasts, Reprogramming, Lentiviral transduction, OKSM, Teratoma assay
1
Introduction
In 2006, Shinya Yamanaka and Kazutoshi Takahashi reported that
by overexpressing four transcription factors, namely, Oct4, Klf4,
Sox2, and c-Myc (OKSM), mature differentiated cells such as
mouse embryonic fibroblasts (MEFs) can be reprogrammed into
embryonic stem (ES) cell-like cells, which they termed induced
pluripotent stem (iPS) cells [1]. A year later, Yamanaka and colleagues generated human iPS cells using OKSM [2]. This discovery
opened up a new research field and gave rise to a number of
paradigms for the use of iPS technology in basic research and
medicine. For example, iPS cells can be generated from patients
with genetic disorders and then differentiated into a cell type of
interest to model and study disease processes and progression
Xiaodong Liu and Joseph Chen contributed equally to this work.
Ivan Bertoncello (ed.), Mouse Cell Culture: Methods and Protocols, Methods in Molecular Biology, vol. 1940,
https://doi.org/10.1007/978-1-4939-9086-3_5, © Springer Science+Business Media, LLC, part of Springer Nature 2019
63
64
Xiaodong Liu et al.
in vitro [3, 4]. In addition, these cells can be used as screening
platforms for the identification and development of novel therapeutic compounds [5]. Furthermore, basic mechanistic studies
during the reprogramming process are starting to provide us with
a basic framework to understand transcription factor-based reprogramming [6–8]. However, more studies are required to complete
this picture. For this purpose, mouse iPS cells remain a suitable and
useful model because very stringent functional assays to test their
potential such as gestational complementation, germline transmission, tetraploid complementation, and single-cell chimerism can be
readily assessed in the mouse system [9]. In addition, cell types like
MEFs are relatively easy to maintain and reprogram considerably
faster than their human counterparts (~2 weeks vs ~4 weeks,
respectively) [7, 10].
A variety of methods have been developed to generate iPS cells
since their discovery, and each method has advantages and disadvantages as reviewed by Robinton and Daley [3]. Among these
methods, using a lentiviral system is a cost-effective, robust, and
efficient approach for transgene delivery since lentiviruses can
transduce almost all mammalian cells, including dividing and nondividing cells [11]. Furthermore, using a doxycycline (Dox)inducible polycistronic cassette encoding the four reprogramming
factors OKSM, in combination with a Tet-on transactivator (rtTA)
[12], allows temporal control of the expression of the Yamanaka
factors to obtain transgene-independent bona fide iPS cells.
In order to determine if the cells that were reprogrammed are
indeed pluripotent, further analyses to verify pluripotency are
required. Analyses of iPS cell lines by flow cytometry and through
the alkaline phosphatase assay are good and quick screening methods to discard aberrant or differentiated lines. As such pluripotent
cell lines should be positive for pluripotent cell surface markers like
SSEA1 and EpCAM [6] and express the cytoplasmic enzyme alkaline phosphatase at high levels [13]. A widely accepted and more
stringent, albeit time-consuming, assay for a functional demonstration of pluripotency potential is the teratoma formation assay. This
assay entails the injection of the iPS cells into flanks of immunocompromised mice to assess their in vivo differentiation potential
[14]. The principle of this assay is to test whether the iPS or ES cell
lines are capable of generating derivatives of all three germ layers,
one of the crucial hallmarks of pluripotency [9].
In the context of this chapter, we describe (1) the generation of
iPS cells from MEFs using a Dox-inducible lentiviral transduction
system in the serum/LIF condition; (2) the subsequent isolation of
clonal iPS cell lines for downstream experiments; and (3) characterization techniques like alkaline phosphatase staining, flow cytometry, and the teratoma formation assay to verify that the resulting iPS
cell lines are pluripotent.
Mouse Induced Pluripotent Stem Cells
2
65
Materials
2.1 Mouse
Embryonic Fibroblasts
Mouse embryonic fibroblasts (MEFs), isolated from embryos of
any genetic background of interest, can be used for reprogramming
experiments. Isolation of MEFs from embryonic day (E) 13.5
mouse embryos is described in great detail in [15]. Alternatively,
primary MEFs can be purchased commercially from various companies such as Merck Millipore (PMEF-CFL-P1).
2.2 Reagents
for Lentiviral
Transduction
1. Generation and titer determination of lentiviral particles harboring the OKSM construct (OKSM plasmid (Millipore,
SCR513)) [12] and the m2rtTA (Ef1a-rtTA-GFP plasmid)
construct are described in detail in a preceding chapter in this
volume [16].
2. Polybrene infection/transfection reagent 10 mg/mL stock
(used at 1:1700 dilution).
2.3 General Cell
Culture Reagents
1. MEF culture medium (MEF media): Dulbecco’s modified
eagle medium (DMEM), containing 10% fetal bovine serum
(FBS) (v/v); 1% GlutaMAX supplement (100) (v/v); 1%
MEM nonessential amino acid solution (100) (v/v);
100 μM sodium pyruvate; 55 μM β-mercaptoethanol.
2. mESC/iPSC culture medium (iPSC media): KnockOut
DMEM containing 15% FBS (v/v) (see Note 1); 1% GlutaMAX
supplement (100) (v/v); 1% MEM nonessential amino acid
solution (100x) (v/v); 55 μM β-mercaptoethanol; 1000 unit/
mL recombinant murine LIF.
3. Dulbecco’s phosphate-buffered saline (DPBS) without calcium
and magnesium.
4. 0.1% gelatin solution (w/v) is prepared by mixing 1 g of gelatin
from porcine skin with 1 L ultrapure water (milli-Q). Autoclave
to dissolve and sterilise.
5. Cryopreservation medium: 90% FBS (v/v) and 10% dimethyl
sulfoxide (DMSO) (v/v).
6. 0.25% trypsin-EDTA.
7. Doxycycline hyclate diluted to 2 mg/mL in dH2O (1000
stock).
8. Mr. Frosty freezing containers.
2.4 Equipment
for iPS Cell Colony
Isolation
1. Dissection microscope.
2. Irradiated mouse embryonic fibroblasts (iMEFs) can be generated in house as described in a preceding chapter in this volume
[15]. Alternatively, iMEFs can be purchased commercially from
Merck Millipore (Merck Millipore, PMEF-CFX).
3. 24-well plates.
66
Xiaodong Liu et al.
2.5 Reagents
and Equipment
for Flow Cytometry
1. Anti-mouse BUV395 Thy1.2.
2. Anti-mouse BV421 EpCAM.
3. Anti-mouse SSEA1 Biotin.
4. Streptavidin Pe-Cy7.
5. DRAQ7 viability dye.
6. Tubes for flow cytometry analysis (5 mL Polystyrene RoundBottom Tube with Cell-Strainer Cap) and fluorescence-activated cell sorting (FACS) (5 mL Polypropylene RoundBottom Tube).
7. Flowcytometry instruments such as LSR-II analyzer (BD Biosciences) and BD Influx cell sorter.
2.6 Alkaline
Phosphatase Assay
1. 100 mM Tris–HCl pH 9.5.
2.7 Teratoma
Formation Assay
1. Dulbecco’s phosphate buffered saline (DPBS) with 1% bovine
serum albumin (BSA) (1% BSA/DPBS).
2. Vector Black Alkaline Phosphatase Substrate Kit II (Vector
Laboratories).
2. BD Luer-Lock Tip Syringe (1 mL).
3. 23G 1 1/4 in. needles.
4. Immunodeficient mice: NOD-SCID IL2Rgammanull (NSG).
5. Anesthesia unit.
6. Surgical station and tools.
7. Iodine solution.
8. 20, 70, 80, 90, and 100% ethanol.
9. Biopsy-processing cassettes and biopsy foam pads (see Note 2).
10. Chloroform.
11. 4% (w/v) paraformaldehyde (PFA) in PBS (4% PFA).
12. Tube rotator.
13. Shandon™ Paraffin (paraffin wax).
14. Paraffin bath.
15. Embedding molds.
16. Microtome (or cryostat).
17. Water bath at 56 C.
18. Superfrost™ Plus slides.
19. Hematoxylin and Eosin Stain Kit.
20. MIRAX scanner (or light microscope with camera).
Mouse Induced Pluripotent Stem Cells
3
67
Methods
3.1 Reprogramming
of MEFs into iPS Cells
1. Freshly derive MEFs, as described previously [15], or thaw low
passage (p0–p2) cryopreserved MEFs for reprogramming
experiments.
2. For thawing of cryopreserved MEFs, quickly transfer a cryovial
of MEFs (~2–3 million cells) from liquid nitrogen into a 37 C
water bath.
3. Once thawed, quickly transfer MEFs with a pipette into a
15 mL centrifuge tube containing 10 mL of pre-warmed
MEF media.
4. Pellet the cells by centrifugation at 450 g for 3 min.
5. Remove the supernatant, resuspend the cell pellet (1.5–3 106
cells on average) in 12 mL MEF media, and transfer to a T75
cell culture flask with a vented cap to allow the cells to recover
for 1–2 days before starting the reprogramming experiments.
6. One to two days after recovery, cellularize thawed or freshly
derived cells as follows: remove MEF media, wash cells once
with DPBS to remove traces of serum, and then add 3 mL of
0.25% trypsin-EDTA solution and incubate at 37 C for
3–5 min. Neutralize the enzymatic reaction by the addition of
3 mL MEF media, and pipette medium onto the surface of the
flask 3–5 times to dissociate the MEFs. Transfer the cell suspension into a 15 mL tube.
7. Perform cell counting using a hemocytometer or automated
cell counter.
8. It is recommended to seed cells at a range of
0.5–2.5 103 cells/cm2 in gelatin-coated 6-well plates (see
Note 3) containing MEF media (see Note 4) (Fig. 1).
Fig. 1 Schematic depicting MEF to iPSC reprogramming protocol
68
Xiaodong Liu et al.
9. Twenty-four hours later, perform lentiviral transduction as
follows: prepare viral mix by adding polybrene (1:1700),
lentivirus-m2rtTA (mean occurance of infection [MOI] of 2),
and lentivirus-OKSM (MOI of 2) (see Note 5 and Chap.4) in
2 mL iPSC media; following this aspirate culture media from
wells to be infected, and replace with the iPSC media containing the viral mix (Fig. 1).
10. Perform spin inoculation by transferring the plate(s) into a
centrifuge, and spin for 60 min at 750 g at room temperature. Afterward, transfer the plate(s) into a 37 C incubator
with 20% O2 and 5% CO2 (see Note 6).
11. On the next day, remove virus-containing media, and replace
with fresh iPSC media supplemented with doxycycline (2 μg/
mL) to initiate the reprogramming process (3 mL of media per
well of 6-well plates).
12. Perform media changes every other day using doxycyclinesupplemented iPSC media for the first 6 days of
reprogramming.
13. After 6 days, daily media changes are recommended due to
increased cell densities. Alternatively, add 6 mL of media into
one well of a 6-well plates if media changes can only be performed every other day.
14. Expected changes in cell morphology during reprogramming
are shown in Fig. 2. iPS cell colonies should be identifiable after
approximately 12 days, and it is recommended to transfer the
cells into doxycycline-free iPSC media for another 4 days to
remove aberrant iPS cell colonies that are still dependent on
forced transgene expression. After this period proceed to isolate clonal lines by colony picking.
3.2 Isolation
of Clonal iPS Cell Lines
by Colony Picking
1. Six hours to 1 day before colony isolation, prepare recipient plates by seeding iMEFs onto gelatin-coated 24-well plates
at a density of 2 104 cells/cm2 in 1 mL of iPSC media per
well (see Note 3).
Fig. 2 Timeline of reprogramming from MEFs to iPSCs. Representative brightfield images of reprograming cultures on days 0, 3, 6, 12, and 16. Scale
bar ¼ 25 μM
Mouse Induced Pluripotent Stem Cells
69
Fig. 3 Establishment of clonal iPSC lines. (a) (i) Identify dome-shaped colony in culture (Day 16 of reprogramming). (ii) Isolate colony with a pipette by removing cells surrounding the colony. (iii) Lift colony with pipette,
gently nudge side to detach cells from well plate. (iv) Dissociate cells by transferring colony into 1.5 mL tube
containing 0.25% trypsin. Gently pipette to dissociate colony further. After 2–4 min, transfer contents of
1.5 mL tube into a 24-well with iMEFs. (v) Bright field image of cells after 3 days in culture after transfer. (b)
Bright field image of iPSC colonies at passage 2. (c) FACS analysis of iPSC clonal line at passage 2. (d)
Alkaline-phosphatase staining of iPSC colonies at passage 2. Scale bar ¼ 25 μM
2. Rinse the 6-well plate(s) containing the reprogrammed cultures with DPBS, and then add 1 mL of warm DPBS into
each well. Colonies can be picked under an inverted light
microscope or a dissection microscope (see Notes 7 and 8).
3. Identify a reasonably isolated (i.e., not fused to other colonies)
iPS cell colony with characteristically dome-shaped morphology (Fig. 3a(i)).
4. Using a 20 μL pipette, draw a circle around the colony with a
sterile tip to detach the colony from surrounding fibroblasts
(Fig. 3a(ii)).
5. Nudge the colony with the tip to gently lift it from the underlying tissue culture plastic (Fig. 3a(iii)).
6. Aspirate the free-floating colony with the pipette in a 12.5 μL
volume, and transfer into a 1.5 mL tube containing 50 μL of
0.25% trypsin-EDTA (Fig. 3a(iv)).
7. After 2–4 min, gently further dissociate the transferred colony
by pipetting the medium within the tube several times.
8. Transfer the cell suspension from the tube directly into a well of
the prepared 24-well plate with the iMEFs (Fig. 3a(iv)).
70
Xiaodong Liu et al.
9. Repeat steps 3–8 with other colonies to generate more potential clonal lines (see Note 9).
10. Change media with fresh iPSC media 24 h after colony picking.
11. New, dome-shaped colonies should form in the recipient
24-wells after 2 days (Fig. 3a).
12. Expand new clones through routine passaging to propagate
the iPS cells (Fig. 3b) (see Notes 10, 11 and 12) as
described for mouse embryonics stem cells in a preceding
chapter in this volume [17].
3.3 Characterization
of Clonal iPS Cell Lines
3.3.1 Flow Cytometry
iPS cells can be analyzed by flow cytometry or purified through
FACS using positive markers associated with pluripotent cells and a
MEF marker they are negative for. For example, iPS cells can be
FACS depleted from Thy1.2-positive feeder cells and enriched for
SSEA1- and EpCAM-positive pluripotent cells (Fig. 3c). Only iPS
lines that express SSEA1 and EpCAM should be considered as
pluripotent. By extracting the SSEA1/EpCAM double-positive
population by FACS, undifferentiated iPS cells can be effectively
removed and the purified cells subsequently used for purposes such
as differentiation assays. Preparation of single-colour compensation
samples, antibody labelling process, and gating strategies were
described in great detail previously [15, 18, 19]; setting up the
voltages at the cell analyzer or sorter is to be performed by either an
experienced user or a dedicated FACS operator as described previously [15, 19].
3.3.2 Alkaline
Phosphatase Staining
Established clonal iPS cell lines (passaged 4–5 times to enrich for
pluripotent cells) can be submitted to an alkaline phosphatase assay.
For this assay, cells can be seeded in a 24-well or 12-well format.
When cultures are confluent (roughly 70% confluent colonies),
remove iPSC media from the wells, wash once with DPBS, and
then stain with alkaline phosphatase assay (Vector Black Alkaline
Phosphatase Substrate Kit II, SK-5200) according to manufacturer’s instructions. iPS cell colonies will stain black (Fig. 3d).
3.3.3 Teratoma
Formation Assay
iPS cells that have been established for at least 4–5 passages are
normally subjected to the teratoma assay. Upon injecting iPS cells
subcutaneously, they should start proliferating and differentiate
into the cell types of all three germ layers and thereby form a
growth, the so-called teratoma. If the resulting teratoma indeed
contains cells from the three germ layers (ectoderm, mesoderm,
and endoderm), the iPS cell line is deemed to be pluripotent. In
order to perform a teratoma formation assay, approval has to be
obtained from the Animal Welfare Committee or other regulatory
bodies before conducting any of these experiments.
Mouse Induced Pluripotent Stem Cells
71
1. Expand subcloned iPS cell lines into one T25 flask in the
presence of iMEF feeders.
2. When the cells are ready to be passaged, remove iPSC media,
then wash once with DPBS, and add 3 mL trypsin-EDTA
solution for 3–5 min at 37 C.
3. Neutralize the enzymatic reaction by the addition of 3 mL of
MEF media, and pipette up and down three to five times to
dissociate the iPS cells, and then transfer the cells to a
15 mL tube.
4. Pellet the cells by centrifugation at 450 g for 3 min, and then
resuspend the cell pellet in 12 mL iPSC media.
5. To purify and enrich iPS cells prior to the assay, an iMEF feeder
depletion step is performed. Transfer the cells onto a new
gelatin-coated T75 flask (see Note 3), and place the flask into
a 37 C incubator for 45–50 min. iMEF feeders and differentiated cells attach to the gelatin within 45 min of incubation
time, whereas iPS cells require 2–4 h to attach.
6. Transfer the supernatant (containing the nonadherent iPS
cells) to a 15 mL tube, pellet the cells by centrifugation at
450 g for 3 min, and then resuspend the cell pellet in
1–3 mL iPSC media to perform cell counting using a hemocytometer or automated cell counter.
7. Transfer ~1 106 cells into an Eppendorf tube, and pellet the
cells by centrifugation at 450 g for 3 min.
8. Resuspend the cell pellet in 200 μL prepared injection mix
containing 1% BSA/PBS or iPSC media, and keep on ice (see
Note 13).
9. Set up the anesthesia apparatus and surgical station in the
animal facility according to the instructions provided by the
manufacturer (see Note 14).
10. Before anesthetizing the mice, fill the anesthetic apparatus’
induction chamber by setting the oxygen flow rate at 4 L/
min and isoflurane at 4–5% for 1–2 min.
11. Once the induction chamber is filled, decrease the oxygen flow
rate and isoflurane to maintenance level (0.4 L/min flow rate
and 2–3% isoflurane).
12. Anesthetize the NGS mice using 2–3% isoflurane and 0.4 L/
min oxygen flow rate for anesthesia and its maintenance once
they are unconscious (Fig. 4a).
13. Remove one mouse from the induction chamber, and place a
nose cone on it to provide consistent anesthetic air flow.
14. Wipe the skin around the dorsal flanks of the mouse with
iodine solution and then 70% ethanol.
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Xiaodong Liu et al.
Fig. 4 Overview of teratoma assay. (a) Schematic depicting injection of iPSCs into NGS mouse and resulting
teratoma formation and isolation. (b) Hematoxylin and eosin (H&E) staining of teratoma sections and
visualization of three representative tissue types per germ layer. Scale bar ¼ 25 μM
15. Using a 1 mL syringe with 23-gauge needle, slowly draw
200 μL of the prepared cell suspension, remove bubbles from
the syringe, and proceed immediately to the next step.
16. Slowly inject 200 μL of the cell suspension subcutaneously into
the dorsal flanks of the NGS mouse. This can be done by
pinching a part of the mouse’s flank using the thumb and the
index finger (Fig. 4a) followed by inserting the needle between
Mouse Induced Pluripotent Stem Cells
73
the fingers. After ensuring that the needle is positioned subcutaneously, slowly inject the cell suspension.
17. After injection, keep the mouse anesthetized for ~10 min.
18. Monitor injected NGS mice at least once a week for 3–5 weeks
to track growth at the injection site.
19. When teratoma formation is evident at a stage when the
growth is still smaller than or around 1 cm3, it is recommended
(and an ethical requirement) to cull the mouse and excise the
teratomas (see Note 15).
20. Transfer the teratoma to a 50 mL tube, and submerge into an
ample volume of 4% PFA.
21. Fix tissue overnight at 4 C on a tube rotator.
22. On the following day, wash the teratomas with DPBS, and cut
into 2–5 slices using scalpels (the choice of the number of the
slices depends on the size of the teratoma and the personal
choice of the experimenter).
23. Place the teratoma slices into a labelled biopsy-processing cassettes with biopsy foam pads, and submerge in a container with
70% ethanol for sectioning.
24. Dehydrate tissue using graded alcohols (70, 80, 90, 100%) by
successively incubating in each solution (starting with 70%) for
at least 20 min, and then submerge in fresh chloroform solution twice for 1 h each.
25. Incubate cassettes twice in paraffin wax (molten at 56 C) for at
least 1 h (see Note 16).
26. After the tissue has been infiltrated with paraffin wax (step 25),
place cassette in a paraffin bath at 58 C for 15 min to melt
away residual wax.
27. Open cassette and pick tissue out of the cassette with a pair of
forceps. Transfer tissue onto embedding molds, and position it
preferably in the center of the depression of the mold. When
tissue is placed in the desired orientation, fill remaining portion
of mold with hot paraffin to desired volume. Place mold in
20 C freezer for at least 3 h before separating the tissue
block from the mold.
28. Section paraffin-embedded tissue block using a microtome
(or cryostat). Cut sections at 2–5 μm according to the manufacturer’s instructions (see Note 17).
29. Using a pair of forceps, transfer sections onto a 20% ethanol
bath (20% v/v ethanol in water), and then transfer sections
onto a heated water bath of 56 C. Collect sections onto
labelled slides, and leave to drain for 10 min (leave upright).
Leave slides to dry overnight on a slide rack in an oven at 40 C.
74
Xiaodong Liu et al.
30. Hematoxylin and eosin staining should be performed according to histology facility’s or manufacturer’s instruction. Refer
to Nelakanti et al. [20] for a detailed protocol of the staining
technique.
31. To obtain high-quality images and identify tissues of all three
germ layers, use a MIRAX scanner (or any other comparable
slide scanners).
32. Score the images for the presence of derivatives of the three
germ layers. Examples of representative tissues of each germ
layer are provided in Fig. 4b.
4
Notes
1. It is important to note that the batch and quality of FBS are
crucial to support pluripotent stem cell culture and reprogramming. Not all FBS batches are suited for reprogramming. If
batch testing is not possible, procure ES-qualified FBS
(in general more expensive).
2. Contact local histology platforms in advance for submission of
specimens for subsequent processing of teratoma assays.
3. Sterile 0.1% gelatin solution (w/v) is used to coat the plates or
flasks to provide attachment support to the MEFs. It is recommended to add adequate 0.1% gelatin to cover the surface of
the plates or flasks (e.g., 2 mL for a well of 6-well plate) and
incubate for 30 min or more at 37 C to coat.
4. Starting cell density has a dramatic impact on the reprogramming efficiency. We recommend trying a range of cell densities
to determine the optimal density for reprogramming experiments for that particular cell line.
5. MOI can be calculated based on the target cell number to be
transduced multiplied by the number of infective viral particles
per microliter of viral concentrate. When handling viruses,
please ensure that transduction is performed in a class II hood
and the user is double-gloved for safety and protection. Refer
to the previous chapter [16] for further details.
6. This reprogramming protocol is optimized for culture in normoxic conditions.
7. It is recommended to pick iPSC colonies using a dissection
microscope to derive clonal lines. Colony picking should be
performed in a sterile condition (e.g. inside a hood; and the
surface and the hood, and all tools and equipment should be
cleaned thoroughly with 80% ethanol before colony picking). It
is easiest to pick colonies a minimum format size of a 6-well
plate.
Mouse Induced Pluripotent Stem Cells
75
8. Before picking colonies, seed iMEFs in 24-well plates 24 h in
advance. For better visualization of colonies (and to facilitate
dissociation of colonies after picking later), remove iPSC
media, and wash cells with PBS. Aspirate and add 2 mL of
PBS in each 6-well plate (or 10 mL in a 10 cm dish). When
selecting a colony to pick, avoid colonies with a flattened/
differentiated appearance.
9. In order to derive 3–5 clonal lines, we advise picking a minimum of 20 colonies as not all colonies will expand after this
process.
10. Three to five days after establishing the clonal lines, cells can be
expanded into larger formats (from 24-well into a 6-well plate)
with iMEF feeders seeded 24 h prior to expansion.
11. It might be necessary to passage new clonal lines for a few times
(at least 2–3 passages) to get rid of cells that are not fully
reprogrammed/partially differentiated and enrich for true
iPSC colonies with dome-shaped morphology.
12. For cryopreservation and routine passaging of iPSCs, it is
recommended to follow the protocol described in a previous
chapter in this volume [17].
13. Matrigel diluted 1:3 in DMEM/F12 can be used to increase
teratoma formation as it enhances cell engraftment after injection [21]. Keep thawed Matrigel on ice at all times to prevent
solidification.
14. Depending on the equipment, delivery method and time of
anesthetic exposure to the animal can vary. It is recommended
that this part of the protocol be performed based on the facility’s preferences and equipment’s instruction manual.
15. Extraction of the formed teratomas is explained in detail by
Nelakanti et al. [20].
16. It is important to take note of the temperature of molten wax as
high temperatures and prolonged exposure to molten wax may
destroy antigens in tissue. Keep the wax at the lowest temperature possible in its molten state. Preferably, maintain this temperature at 2 C above the melting point of 56 C.
17. The operation of the microtome (or cryostat) and sectioning of
slides should be performed by trained personnel in accordance
to manufacturer’s instructions for the machines.
References
1. Takahashi K, Yamanaka S (2006) Induction of
pluripotent stem cells from mouse embryonic
and adult fibroblast cultures by defined factors.
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2. Takahashi K, Tanabe K, Ohnuki M, Narita M,
Ichisaka T et al (2007) Induction of pluripotent stem cells from adult human fibroblasts by
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3. Robinton DA, Daley GQ (2012) The promise
of induced pluripotent stem cells in research
and therapy. Nature 481:295–305
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wnt stimulation enhances differentiation of
pluripotent stem cells independent of β-catenin-mediated T-cell factor signaling. Stem
Cells. 36:822–833.
5. Avior Y, Sagi I, Benvenisty N (2016) Pluripotent stem cells in disease modelling and drug
discovery. Nat Rev Mol Cell Biol 17:170–182
6. Polo JM, Anderssen E, Walsh RM, Schwarz
BA, Nefzger CM et al (2012) Molecular roadmap of reprogramming somatic cells into iPS
cells. Cell 151:1617–1632
7. Nefzger CM, Rossello FJ, Chen J, Liu X,
Knaupp AS et al (2017) Cell type of origin
dictates the route to pluripotency. Cell Rep
21(10):2649–2660
8. Knaupp AS, Buckberry S, Pflueger J, Lim SM,
Ford E et al (2017) Transient and permanent
reconfiguration of chromatin and transcription
factor occupancy drive reprogramming. Cell
Stem Cell 21:834–845
9. De Los Angeles A, Ferrari F, Xi R, Fujiwara Y,
Benvenisty N et al (2015) Hallmarks of pluripotency. Nature 525:469–478
10. Liu X, Nefzger CM, Rossello FJ, Chen J,
Knaupp AS et al (2017) Comprehensive characterization of distinct states of human naive
pluripotency generated by reprogramming.
Nat Methods 14:1055–1062
11. Sakuma T, Barry Michael A, Ikeda Y (2012)
Lentiviral vectors: basic to translational. Biochem J 443(3):603–618
12. Sommer CA, Stadtfeld M, Murphy GJ,
Hochedlinger K, Kotton DN, Mostoslavsky G
(2009) iPS cell generation using a single lentiviral stem cell cassette. Stem Cells 27:543–549
13. Singh U, Quintanilla RH, Grecian S, Gee KR,
Rao MS, Lakshmipathy U (2012) Novel live
alkaline phosphatase substrate for identification
of pluripotent stem cells. Stem Cell Rev
8:1021–1029
14. Wesselschmidt RL (2011) The teratoma assay:
an in vivo assessment of pluripotency. Methods
Mol Biol 767:231–241
15. Nefzger CM, Alaei S, Knaupp AS, Holmes ML,
Polo JM (2014) Cell surface marker mediated
purification of iPS cell intermediates from a
reprogrammable mouse model. J Vis Exp
(91):e51728.
16. Larcombe MR, Manent J, Chen J, Mishra K,
Liu X, Nefzger CM (2019) Production of high
titer lentiviral particles for stable genetic modification of mammalian cells. Methods Mol Biol
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17. Paynter JM, Chen J, Liu X, Nefzger CM
(2019) Propagation and maintenance of
mouse embryonic stem cells. Methods Mol
Biol 1940:33–45
18. Nefzger CM, Alaei S, Polo JM (2015) Isolation
of reprogramming intermediates during generation of induced pluripotent stem cells from
mouse embryonic fibroblasts. Methods Mol
Biol 1330:205–218
19. Nefzger CM, Jarde T, Rossello FJ, Horvay K,
Knaupp AS et al (2016) A versatile strategy for
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20. Nelakanti RV, Kooreman NG, Wu JC (2015)
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Stem Cells 31:1498–1510
Chapter 6
Gene Editing of Mouse Embryonic and Epiblast Stem Cells
Tennille Sibbritt, Pierre Osteil, Xiaochen Fan, Jane Sun, Nazmus Salehin,
Hilary Knowles, Joanne Shen, and Patrick P. L. Tam
Abstract
Efficient and reliable methods for gene editing are critical for the generation of loss-of-gene function stem
cells and genetically modified mice. Here, we outline the application of CRISPR-Cas9 technology for gene
editing in mouse embryonic stem cells (mESCs) to generate knockout ESC chimeras for the fast-tracked
analysis of gene function. Furthermore, we describe the application of gene editing directly to mouse
epiblast stem cells (mEpiSCs) for modelling germ layer differentiation in vitro.
Key words CRISPR-Cas9, Embryonic stem cells, Epiblast stem cells
1
Introduction
Conventional methods to perform genome editing in embryonic
stem cells (ESCs) such as gene targeting by homologous recombination are inefficient and time-consuming, taking up to months to
a year to generate a stock of genetically modified mice for experimental studies of gene function. Recently, advances in genetic
manipulation technology have enabled the quick and efficient generation of edited genomes. Nucleases fused to specific
DNA-binding domains, such as transcription activator-like effector
nucleases (TALENs) and zinc-finger nucleases (ZFNs), have facilitated highly specific gene editing that can be achieved expeditiously,
but the utility and the cost-effectiveness of these technologies
remain challenging [1, 2]. CRISPR-Cas9 technology has recently
arisen to the fore as the most amenable technique to perform
genome editing [3]. In addition to producing the desired genetic
modification within a shorter time frame, this technology is efficient, is relatively straightforward in design, and can be applied to
both cell lines and whole organisms.
Here, we describe the use of CRISPR-Cas9 for genome editing
in mouse (m) ESCs and EpiSCs. Several resources are available to
design the single-guide RNAs (sgRNAs) to target the gene of
Ivan Bertoncello (ed.), Mouse Cell Culture: Methods and Protocols, Methods in Molecular Biology, vol. 1940,
https://doi.org/10.1007/978-1-4939-9086-3_6, © Springer Science+Business Media, LLC, part of Springer Nature 2019
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Tennille Sibbritt et al.
interest, such as the Zhang Lab website (http://crispr.mit.edu/) or
the Broad Institute’s Genetic Perturbation Platform (https://
portals.broadinstitute.org/gpp/public/). Commonly, a combination of these resources is used to select the optimal sgRNAs that
target within the first 200 bp of the start codon in the coding
region and have high specificity with low off-target effects. Furthermore, the sgRNA must be followed on the 30 end by the 3 bp NGG
protospacer adjacent motif (PAM). The cloning of the sgRNA for
targeting the gene of interest into plasmids containing the Cas9
nuclease, a selection marker (puromycin or GFP), and a sgRNA
scaffold (pSpCas9(BB)-2A-Puro or pSpCas9(BB)-2A-GFP) can be
ready within a week for use in the electroporation of stem cells
[3]. Within another 3 weeks, mESCs or mEpiSCs harboring target
mutations can be established, which can be taken further for clone
selection by puromycin resistance or GFP expression. The
subsequent expansion of positive clones for cryopreservation and
genotyping usually takes at most another 4 weeks.
We have previously generated chimeric embryos derived predominantly from the targeted genome-edited ESCs by microinjecting the stem cells into eight-cell preimplantation mouse embryos
which are transferred to pseudopregnant mice for further intrauterine development [4, 5]. In addition to the capability to generate
mid-gestation embryos of the edited genotype for the phenotypic
analysis and elucidation of gene function, this approach has also
been employed to derive genome-edited mEpiSCs from the chimeric embryos [4]. The generation of ESC-derived chimeras enables
the transition of the stem cells through the development of the
inner cell mass to an epiblast state from which authentic mEpiSCs
can be derived. This approach bypasses the technical barrier that the
transition from mESCs to mEpiSCs of a proper epiblast-like state
has not been achieved in vitro. However, derivation of mEpiSCs
with an edited genotype via chimeric embryo production is timeconsuming and laborious. Moreover, the efficiency of derivation of
mEpiSCs is lower than that of mESCs, resulting in increased animal
usage. This technically demanding protocol and the low efficiency
of mEpiSC derivation could be replaced by a genome editing
protocol that applies directly to mEpiSCs. Efforts to perform
gene editing directly on mEpiSCs have been hampered by the
technical difficulty in maintaining the mEpiSCs (1) without feeders
(that interferes with the selection of edited cells) and (2) as single
cells for clonal selection of editing events. These hurdles are
reputed to be the different requirements for maintaining the
mEpiSCs due to their primed pluripotent state and the poising of
cell differentiation. We have overcome these issues with the use of
conditioned mEpiSC medium obtained from culture with mouse
embryonic fibroblasts (MEFs) and chemical reagents that enhance
the viability of single mEpiSC cells during clonal selection in vitro.
Gene Editing Embryonic and Epiblast Stem Cells
79
Gene editing of mESC and mEpiSCs by these approaches is
efficient. In the case of puromycin selection, 80–100% of clones
that successfully grow contain a mutation on at least one allele of
the gene of interest. The success rate of generating biallelic frameshift mutations varies from 35% to 100%. With GFP selection, we
generally find mutations in 60–80% of the sequenced clones, with
10–20% of these containing biallelic frameshift mutations. For
several genes that have been edited by CRISPR-Cas9 technology,
we were able to confirm the absence of protein for the biallelic
frameshift mutations in mESC clones and the reduction of protein
in some monoallelic mESC clones. For mEpiSCs, sequencing analysis of a mixed population of clones revealed that desirable editing
events have taken place.
In this chapter, we describe procedures for generating frameshift mutations in mESCs and directly in mEpiSCs (Fig. 1).
2
Materials
Prepare and store all reagents at room temperature unless indicated
otherwise in protocol or packaging of reagent. Diligently follow all
safety and waste disposal regulations when performing
experiments.
2.1 CRISPR Plasmid
Components
1. pSpCas9(BB)-2A-Puro (PX459) V2.0 (Addgene plasmid
#62988) or pSpCas9(BB)-2A-GFP (PX458) (Addgene plasmid #48138) (gift from Feng Zhang) with the sgRNA ligated
into the plasmid according to the protocol described in [3].
2.2 Cell Culture
Components
1. 100 nucleosides: 0.16 g adenosine, 0.146 g cytidine, 0.17 g
guanosine, 0.146 g uridine, 0.048 g thymidine in 200 mL
H2O. Swirl at 37 C for several hours or overnight. Filter
before use and store aliquots at 4 C. Warm to 37 C prior
to use.
2. Mouse embryonic stem cell (mESC) culture medium: DMEM,
12.5% heat-inactivated fetal calf serum (FCS), 1000 U/mL
leukemia inhibitory factor, 0.1 mM β-mercaptoethanol, 1
nonessential amino acids, 1 nucleosides. Store at 4 C.
3. Mouse embryonic fibroblast (MEF) culture medium: DMEM,
10% FCS, 0.1 mM β-mercaptoethanol. Store at 4 C.
4. Mouse epiblast stem cell (mEpiSC) medium: Knockout Serum
Replacement, 1 nonessential amino acids, 1 GlutaMAX™,
0.1 mM β-mercaptoethanol. Store at 4 C. Media is supplemented with 10 ng/mL recombinant human FGF2 and
20 ng/mL recombinant Human/Mouse/Rat Activin A
extemporaneously.
PX459 V2.0
PX458
2A
2A
Pu
ro R
CRISPR
plasmid
FP
CRISPR
plasmid
Electroporate
mESCs or mEpiSCs
U6
EG
U6
(add ROCKi to mEpiSCs)
20mer oligo
cloning site
20mer oligo
cloning site
Select GFP positive
clones ~3-4 days
after transfection
Rest for 24 h
Puromycin select
(48 h mESCs, 24 h mEpiSCs)
Select clones
(add ROCKi to mEpiSCs for 24 h)
mESCs
Freeze clones & grow on gelatin to
extract genomic DNA for genotyping
mEpiSCs
(conditioned medium + ROCKi)
PCR amplify, Sanger sequence & analyse mutation using TIDE
mEpiSCs
mESCs
*
PAM
*
PAM
Clone PCR products into pGEM®-T Easy Vector & Sanger sequence
PAM
Wild-type
e.g. mESCs
*
PAM
*
PAM
Positive
clone
Allele 1: 7 bp deletion
Allele 2: 2 bp deletion
Thaw, expand and freeze positive clones
Validate clones
Sanger sequence for
final validation
mRNA expression analysis
(e.g. RT-qPCR)
Protein expression analysis
(e.g. western blot)
Fig. 1 Workflow for genome editing in mESCs and mEpiSCs using PX459 V2.0 or PX458. PX459 V2.0 and
PX458 contain the S. pyogenes Cas9 nuclease ORF and a cloning backbone for the sgRNA. PX459 V2.0
contains the 2A-Puro ORF directly downstream of the Cas9 ORF, while PX458 contains the 2A-EGFP ORF
Gene Editing Embryonic and Epiblast Stem Cells
81
5. Conditioned mEpiSC medium: mEpiSC medium is incubated
overnight at 37 C on MEFs at a density of 9 104 cells/cm2.
The following day, the medium is filtered. The conditioned
medium can be stored at 20 C for up to 1 month. Use only
for cell culture without MEFs, and supplement with Activin A
and FGF2 as mentioned in step 4.
6. 2 freeze medium: 50% heat-inactivated FCS, 30% culture
medium, 20% DMSO. Make up fresh each time and keep at
4 C.
7. TrypLE™ Select.
8. Collagenase IV: Make a stock of 2 mg/mL in mEpiSC
medium.
9. ROCK inhibitor (Y-27632, TOCRIS): ROCKi is used only
when mEpiSCs are seeded in a single cell suspension to
improve cell viability. Add to mEpiSC media to a final concentration of 10 μM only for 24 h following TrypLE™ Select
treatment.
10. Dulbecco’s (D)PBS.
11. Puromycin (10 mg/mL).
12. 0.1% gelatin: Mix 1 g gelatin from bovine skin with 1 L
H2O. Autoclave and filter before use.
13. MicroTube Rack System™ Tubes.
2.3 Electroporation
Components
1. Neon® Transfection System.
2. Neon® Transfection System 100 μL kit.
ä
Fig. 1 (continued) directly downstream of the Cas9 ORF, allowing for puromycin or EGFP selection, respectively. After electroporation of the plasmids containing the sgRNA of interest, clones are left to grow. In the
case of mEpiSCs, clones are grown in mEpiSC medium supplemented with 10 μM ROCKi for 24 h, while
mESCs are grown in mESC medium. Once established, clones are selected and grown in a 96-well plate
containing MEFs. One-third (mESCs) or one-half (mEpiSCs) of the cells are cryopreserved, while the remainder
are grown on 0.1% gelatin for several passages to remove MEFs for genotyping of the mutation. In the case of
mEpiSCs grown on gelatin, clones are passaged in conditioned medium with 10 μM ROCKi added after each
passage for 24 h. Clones are lysed and subjected to PCR and Sanger sequencing, followed by analysis by TIDE
to decompose the mutations on each allele. Examples of the chromatograms indicating gene editing are
shown for mESCs and mEpiSCs; overlapping peaks in the sequence of several bases upstream of the PAM
indicate mutations in at least one allele. PCR products are then cloned into pGEM®-T Easy Vector for
confirmation of the gene editing event. An example of a confirmed biallelic frameshift mutation is shown
for mESCs. Final validation of the mutation involves thawing out the cryopreserved clones of interest,
expanding them from a 48-well plate to a 6-well plate over several weeks, and repeating the process of
lysis, PCR, cloning, and Sanger sequencing. Final validation of the knockout is confirmed by mRNA expression
analysis (RT-qPCR) using primers that overlap the indel or protein expression analysis (Western blot). *
indicates the indel site
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2.4 Cell Lysis
Components
1. Cell lysis buffer: 50 mM Tris–HCl, pH 8.0, 1 mM EDTA, 0.5%
Tween-20. Freshly add 200 μg/mL Proteinase K.
2.5 PCR
Amplification
Components
1. BioMix™ (Bioline).
2.6 Agarose Gel
Electrophoresis
and PCR Purification
Components
2. Forward and reverse primers spanning the mutation site
(desired product size ~700 bp, with sufficient sequence flanking the mutation).
1. Agarose.
2. RedSafe™ Nucleic
Biotechnology).
Acid
Staining
Solution
(iNtRON
3. HyperLadder™ 100 bp (Bioline).
4. 1 TAE buffer: Make up 50 TAE buffer by combining 424 g
Tris base, 57.1 mL acetic acid, and 100 mL 0.5 M EDTA
(pH 8.0), and make up to 1 L in H2O. To make 1 TAE
buffer, combine 40 mL 50 TAE buffer with 1.96 L H2O.
5. Wizard SV Gel and PCR Clean-Up System (Promega).
2.7 Ligation,
Bacterial
Transformation,
and Plasmid
Purification
Components
1. pGEM®-T Easy Vector System (Promega).
2. α-Select Silver Competent Cells (Bioline).
3. Luria-Broth (LB): Combine 4 g Tryptone (vegetable), 2 g
Bacto™ Yeast Extract, and 4 g NaCl, and add 400 mL
H2O. Autoclave.
4. LB ampicillin: As for LB, add ampicillin to a final concentration
of 100 μg/mL once cooled enough to touch.
5. Combine 4 g Tryptone (vegetable), 2 g Bacto™ Yeast Extract,
and 4 g NaCl, and add 400 mL H2O. Autoclave.
6. LB ampicillin agar plates: As for LB, add 6.5 g Bacto™ Agar.
Autoclave. Cool to the point that you can touch. Add ampicillin to a final concentration of 100 μg/mL. Under sterile conditions pour ~20 mL into bacterial plates. Store at 4 C for up
to 1 month.
7. LB ampicillin agar plates for bacterial transformations using
pGEM®-T Easy Vector: As for LB ampicillin agar plates, add
fresh 0.5 mM IPTG and 50 μg/mL X-Gal to the plates after
the agar has set. Spread across the plate, and hold with the lid
off in a 37 C incubator until plates have dried (~20 min).
8. ISOLATE II Plasmid Mini Kit (Bioline).
3
Methods
Carry out all procedures at room temperature unless otherwise
specified. Warm all cell culture media components to room
Gene Editing Embryonic and Epiblast Stem Cells
83
temperature or 37 C before use. Unless otherwise stated, all cell
culture is done under sterile conditions in a laminar flow hood.
Follow all waste disposal regulations diligently when disposing
waste materials.
3.1 Electroporation
of mESCs with PX459
v2.0 or PX458
1. 24 h before electroporation, seed a 10 cm plate (two plates for
PX458) with 1.5 106 MEFs in MEF medium, and place into
the 37 C incubator.
2. On the day of electroporation, set up and save the following
parameters on the Neon® Transfection System:
Pulse voltage, 1200 V; pulse width, 20 ms; pulse number, 2
3. Set up the electroporation equipment according to the manufacturer’s instructions.
4. Remove 5 μg of PX459 V2.0 or 3 μg PX458 with the specific
sgRNA ligated in from the stock tube, and aliquot into a
1.5 mL tube (see Note 1). Put aside.
5. Change the medium on the pre-seeded MEFs to 10 mL mESC
medium. Label the plate with all the required details for the
electroporation. Place back into the incubator until required.
6. Take the mESCs out of the 37 C incubator, aspirate the
medium, and rinse with 4 mL DPBS.
7. Add 3 mL TrypLE™ Select to the mESC plate, and place back
into the incubator for 5 min.
8. Take the plate out of the incubator and add 6 mL mESC
medium. Inactivate and break into a single cell suspension (see
Note 2).
9. Centrifuge cells at 1000 rpm (200 g) for 5 min.
10. Once spinning has completed, remove the medium leaving
only the cell pellet, and then resuspend in 10 mL DPBS.
Count the cells.
11. For electroporation with pX459 V2.0, remove 5 106 cells,
and pipette into a new 15 mL tube. For electroporation with
pX458, remove 1 106 cells, and pipette into a new 15 mL
tube (see Note 3).
12. Centrifuge at 1000 rpm (200 g) for 5 min. During this time,
add 3 mL E2 buffer into a Neon® Tube, and place into the
Neon® Pipette Station.
13. When spinning has completed, remove all DPBS leaving only
the cell pellet.
14. Resuspend cells in 120 μL R buffer until they are in a single cell
suspension.
15. Transfer the cells to the 1.5 mL tube containing the plasmid
from step 4. Mix well by gentle pipetting.
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16. Insert a 100 μL Neon® tip into the Neon® Pipette, and collect
the cell and DNA mixture. Ensure there are no bubbles (see
Note 4).
17. Move the Neon® Pipette containing the cells and DNA to the
Neon® Pipette Station.
18. Load the saved parameters on the Neon® Transfection System
from step 2. Press “start.” Once completed, the unit will
display “complete.”
19. Transfer the electroporated cells onto the pre-seeded MEF
plate. Evenly distribute the cells by rocking back and forth
and side-to-side (not swirling).
20. If there are still substantial numbers of cells left in the 1.5 mL
tube, repeat steps 14–19.
21. Return the plate to the incubator.
3.2 Electroporation
of mEpiSCs
with PX459 v2.0
1. 24 h before electroporation, seed 2 6 cm plates with 1 106
MEFs in MEF medium, and place into the 37 C incubator.
2. Repeat steps 2–4 from Subheading 3.1.
3. Rinse the plates containing pre-seeded MEFs with 2 mL DPBS,
and then add 4 mL of mEpiSC medium supplemented with
10 μM ROCKi. Label the plate with all the required details for
the electroporation. Place back into the incubator until
required.
4. Add 2 mL of Collagenase IV to the plates containing mEpiSCs,
and place back into the 37 C incubator for 10 min.
5. Take the mEpiSC plates out of the incubator, and add 2 mL
mEpiSC medium to detach the colonies from the feeder layer.
6. Spin the clumped suspension at 1000 rpm (200 g) for 30 s.
7. Resuspend the cell pellet in 1 mL TrypLE™ Select, and incubate at room temperature for 2 min.
8. Break the cell clumps using a P1000 tip and add 2 mL mEpiSC
medium.
9. Centrifuge cells at 1000 rpm (200 g) for 5 min.
10. Remove the medium leaving only the cell pellet, and then
resuspend in 2 mL DPBS. Count the cells.
11. Proceed with step 11 from Subheading 3.1 for electroporation
of mEpiSCs.
3.3 Puromycin
Selection of mESCs
and mEpiSCs
Transfected
with pX459 v2.0
1. 24 h post-electroporation, feed the electroporated mESC or
mEpiSC media containing 2 μg/mL or 1 μg/mL puromycin,
respectively (see Note 5). Repeat this the following day for
mESCs with fresh mESC media (see Note 6).
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85
2. 72 h post-electroporation of the mESCs (48 h postelectroporation for mEpiSCs), wash the cells twice with
DPBS, and feed with mESC or mEpiSC media.
3. For the electroporated mESCs, seed 1.5 106 MEFs in mESC
medium onto the plate. For mEpiSCs, seed 1 106 MEFs in
mEpiSC medium onto each plate (see Note 7).
4. Continue DPBS wash and feed daily.
3.4 Clone Picking
of pX459 v2.0Transfected mESCs
and mEpiSCs
1. The day before colonies are ready to be picked, seed 8.42 104
MEFs for mESCs or 6.24 104 MEFs for mEpiSCs into the
wells of a 96-well plate (see Note 8). The number of wells to
seed depends on the number of clones to be picked. If possible,
try to pick 20–30 clones. Place the plate into the 37 C
incubator.
2. On the day of picking the mESCs, change the media in the
96-well plate to 80 μL mESC media. For mEpiSCs, rinse once
with 100 μL DPBS to remove any trace of FCS before adding
80 μL mEpiSC media with 10 μM ROCKi. Label the wells with
the clone number (i.e., 1, 2, 3, etc.) and return to the
incubator.
3. In another 96-well plate, add 30 μL TrypLE™ Select to the
same series of wells seeded with MEFs on the plate prepared in
step 2. Label the wells with the clone number (i.e., 1, 2,
3, etc.).
4. Take the electroporated cells out of the incubator, and examine
the clones on the plate. Mark on the plate which clones are
suitable for selection based on morphology (see Note 9).
5. Aspirate the medium from the plate, and rinse the mESCs or
mEpiSCs with 4 mL or 2 mL DPBS, respectively.
6. Add 7 mL DPBS to the mESCs or 2 mL to the mEpiSCs (see
Note 10), and under a microscope, start picking clones, with
each being placed in a single well of the 96-well plate containing TrypLE™ Select (see Note 11). This can be done by setting
a P20 pipette to 4 μL (see Note 12).
7. After 5 min, inactivate TrypLE™ Select by adding 70 μL
mESC medium or mEpiSC medium supplemented with
10 μM ROCKi to each well.
8. Using a multichannel pipette, pipette each well up and down to
dissociate into single cells.
9. Transfer the cells to the 96-well plate containing the MEFs,
and place back into the incubator.
10. Feed the clones daily with 200 μL mESC medium or mEpiSC
medium.
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3.5 Clone Picking
of pX458-Transfected
mESCs
1. Repeat steps 1–2 from Subheading 3.4.
2. Check growth of cells 3 days after electroporation for size and
the presence of fluorescence to decide the best time for clone
picking. A balance needs to be made between strong GFP
expression and a sufficient number of cells per clone (see Note
13).
3. Under a fluorescence microscope, pick clones uniformly
expressing GFP using a mouth pipette, and place directly into
the 96-well plate containing pre-seeded MEFs (see Note 14).
4. Feed the clones daily with 200 μL mESC media until most
clones are large enough to be passaged. As the clones that were
picked were small, they will not grow to fill the entire well.
5. 24 h before passaging the clones, repeat step 1 from Subheading 3.4. On the day of passaging, repeat step 2 from Subheading 3.4.
6. Passage the entire clones onto the 96-well plate that was
pre-seeded with MEFs 24 h earlier (see Note 15).
3.6 Passaging
and Cryopreservation
of mESC Clones
Once clones are almost confluent, it is necessary to cryopreserve a
proportion of the cells as a stock as well as grow the remaining cells
on gelatin to remove MEFs for genomic DNA (gDNA) extraction
and genotyping of the mutation. Not all clones grow at the same
rate, and some clones may not grow at all, so it is necessary to
exercise a compromise between the expansion of cells by enhancing
growth and preventing cell differentiation.
1. Remove mESC media from each well.
2. Rinse each well with 100 μL DPBS and add 30 μL TrypLE™
Select. Place in the incubator for 5 min.
3. During the incubation, coat wells of a fresh 96-well plate with
100 μL 0.1% gelatin, aspirate off gelatin after a few minutes,
and immediately add 160 μL mESC media. The number of
wells to coat is equivalent to the number of clones that has been
picked.
4. Inactivate the TrypLE™ Select by adding 90 μL mESC media
to each well.
5. Using a multichannel pipette, pipette each well up and down to
dissociate into single cells.
6. Transfer 40 μL of the cells onto the plate containing gelatin.
Label the plate accordingly and place into the incubator.
7. To the remaining 80 μL cells, slowly add 80 μL 2 freeze
media, and gently pipette up and down.
8. Transfer each clone to a separate tube of the MicroTube Rack
System™. Attach the lids and label the tubes with the necessary
details.
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9. Place the MicroTube Rack System™ Tubes on ice, and transfer
to a 80 C freezer.
10. Feed the cells on gelatin daily with 200 μL mESC media and
passage when cells are confluent. Generally, three passages on
gelatin are sufficient to remove all MEFs (see Note 16).
3.7 Passaging
and Cryopreservation
of mEpiSC Clones
1. Remove mEpiSC media from each well.
2. Rinse each well with 100 μL DPBS, add 30 μL Collagenase IV,
and place back into the 37 C incubator for 10 min.
3. During the incubation, coat the wells of a new 96-well plate
with 100 μL 0.1% gelatin for 20 min, then aspirate off gelatin,
and immediately add 140 μL conditioned mEpiSC medium
supplemented with 10 μM ROCKi.
4. Add 30 μL mEpiSC medium to the dissociated cells, and
detach the colonies from the feeder layer. Transfer into
1.5 mL tubes.
5. Spin the clump suspension at 1000 rpm (200 g) for 30 s.
6. Resuspend the cell pellet in 120 μL conditioned mEpiSC
medium supplemented with 10 μM ROCKi. Try to dissociate
the clumps as much as possible.
7. Transfer 60 μL of the cell suspension to the plate containing
gelatin. Label the plate accordingly, and place into the
incubator.
8. To the remaining 60 μL cells, add 60 μL 2 freeze media, and
gently pipette up and down.
9. Repeat steps 8 and 9 of Subheading 3.6.
10. Feed the cells daily with 200 μL conditioned mEpiSC medium,
and, when cells are confluent, passage at a 1:2 ratio onto a new
gelatin-coated 96-well plate. Add 10 μM ROCKi after each
passage for 24 h. Three passages through gelatin-coated culture are sufficient to remove all MEFs; however, the mEpiSCs
may take up to 2 weeks to recover.
3.8 Cell Lysis, PCR
Amplification,
and Sanger
Sequencing
for Genotyping
1. Once all MEFs are removed and cells are confluent, aspirate off
media, and rinse with 100 μL DPBS.
2. Add 100 μL lysis buffer with Proteinase K to each of the clones,
and incubate overnight at 56 C (see Note 17).
3. Transfer the lysates to eight-strip tubes, and inactivate Proteinase K by incubating at 95 C for 10 min in a thermocycler (see
Note 18).
4. PCR amplify the region surrounding the mutation for each of
the clones using the conditions in Tables 1 and 2. It will be
necessary to perform the same PCR reaction on a sample that
has not been edited in the same region (see Note 19).
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Table 1
PCR master mix for the amplification of gDNA surrounding the mutation
site using BioMix™
Reagent
Volume for one reaction (μL)
BioMix™
10
Forward primer (10 μM)
1
Reverse primer (10 μM)
1
Template gDNA
2
Sterile deionized water
6
Table 2
PCR cycling conditions for the amplification of gDNA surrounding the
mutation site using BioMix™
Stage
Temperature ( C)
Time
Initial denaturation
95
5 min
35 cycles
95
60
72
30 s
30 s
1 min/kb
4
Forever
Hold
Denaturation
Annealing
Extension
5. Make a 2% agarose gel in 1 TAE that is enough to fill a large
casting tray (~200 mL).
6. Once cool, add 15 μL of RedSafe™ and gently swirl into the
solution. Carefully pour the agarose solution into a gel cast tray
with combs already inserted without introducing air bubbles to
the gel.
7. Load 5 μL of the PCR products into each well. Add 6 μL of the
appropriate ladder into a lane next to your samples (see Note
20). Electrophorese gel at 100 V for 2.5 h to allow for sufficient separation of the bands indicating potential biallelic
mutations.
8. After imaging, select samples in which the fragment size is
relatively close to the expected size, and purify samples from
the remaining 15 μL PCR product using Wizard® SV Gel and
PCR Clean-Up System according to manufacturer’s instructions (see Note 21).
9. Perform Sanger sequencing of the PCR products, including
the unedited control, with the forward primer used for the
PCR amplification.
10. Once the sequencing results have returned, use TIDE
(https://tide-calculator.nki.nl/) [6] to predict the insertions/
Gene Editing Embryonic and Epiblast Stem Cells
89
deletions (indels) of the samples compared to the unedited
control sample (see Note 22).
11. Determine which clones are worth pursuing for further characterization based on the predicted indels, % of sequences with
that indel, and total efficiency of the prediction. The desired
indels should result in a biallelic frameshift mutation (number
of base deletions not divisible by 3), with ~50% sequences with
each indel at a high efficiency.
3.9 Decomposition
of Mutations: Ligation,
Transformation,
and Extraction
of Plasmid DNA
1. Ligate 2 μL of purified PCR product of the selected clones
from Subheading 3.8 into the pGEM®-T Easy Vector according to manufacturer’s instructions (see Note 23).
2. Thaw a vial of the α-Select Silver Competent Cells on ice for
~5 min. Label a 1.5 mL tube for every ligation sample, and
aliquot out 2 μL of the sample into the tubes, and pre-chill on
ice, while competent cells are thawing.
3. Once completely thawed, gently mix the competent cells by
pipetting up and down once before adding 50 μL of cells to
each pre-chilled ligation mix. Pipette up and down gently once
to mix.
4. Incubate mixture on ice for 30 min.
5. Heat shock the cells at 42 C for 30 s in a water bath.
6. Immediately place on ice for 2 min.
7. Add 350 μL of LB to the transformation mix.
8. Incubate transformation mix at 37 C for 1 h, shaking at
200 rpm.
9. Plate 200–300 μL of the transformation mix carefully onto a
pre-warmed LB ampicillin agar plate with IPTG and X-gal (see
Note 24). Incubate overnight at 37 C.
10. Seal transformation plates and place at 4 C the following
morning (see Note 25).
11. Aliquot 4 mL of freshly made LB ampicillin into 8–12 labelled
14 mL Falcon™ Round-Bottom Polypropylene Tubes.
12. Pick 8–12 pure white colonies from the transformation
plate using a yellow 200 μL pipette tip, suspend the colony
into the LB ampicillin, and discard the tip within the tube
(see Note 26).
13. Culture the picked subclones at 37 C overnight, shaking at
200 rpm.
14. Isolate the plasmid using the ISOLATE II Plasmid Mini Kit as
per manufacturer’s instructions. Elute in 30 μL of H2O.
15. Send isolated plasmid samples off for Sanger sequencing using
the M13 reverse primer.
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16. Use https://www.ebi.ac.uk/Tools/msa/muscle/ [7, 8] to
compare sequenced results to the reference sequence (see
Note 27).
3.10 Thawing
and Validation
of Confirmed mESC
and mEpiSC Clones
1. Seed one well of a 48-well plate 1 day prior to thawing confirmed clones with MEFs.
2. On the day of thawing, remove MEF media, and replace with
400 μL mESC medium or mEpiSC medium supplemented
with 10 μM ROCKi.
3. Cut the selected clones from the strips of the MicroTube Rack
System™, and immediately put tube(s) on ice (see Note 28).
4. Hold the tube in the 37 C water bath until almost thawed.
5. Sterilize the outside of the tube with ethanol, and slowly add
200 μL of pre-warmed mESC or mEpiSC media into the tube.
6. Move the thawed cells to a 15 mL tube. Do not discard the
original tube.
7. Add 500 μL of pre-warmed mESC or mEpiSC media into the
original tube to clean up the leftover cells, and move all the
contents into the 15 mL tube.
8. Add 1.5 mL of pre-warmed mESC or mEpiSC media slowly
into the tube with the cells (see Note 29).
9. Centrifuge the cells at 1000 rpm (200 g) for 5 min.
10. Aspirate off the supernatant leaving about 200 μL of media.
11. Carefully resuspend the cells in another 200 μL of pre-warmed
mESC or mEpiSC media supplemented with 10 μM ROCKi.
12. Transfer the cells into the single well of the 48-well plate
pre-seeded with MEFs.
13. Add another 200 μL of pre-warmed mESC medium or
mEpiSC medium supplemented with 10 μM ROCKi into the
15 mL tube to collect the remaining cells, and transfer them
into the well of the 48-well plate (see Note 30).
14. Expand the clones, and freeze down vials of cells whenever
possible until enough cells can be seeded onto a full 6-well
plate (see Note 31).
15. After getting to the 6-well plate stage, also passage cells onto
0.1% gelatin in a 6-well plate for at least three passages. This
plate will be used to extract gDNA from for validation of
mutations.
16. Cryopreserve the 6-well plate of cells on MEFs once it becomes
confluent. Each well is enough to cryopreserve into two vials
that can be thawed onto a single well of a 6-well plate.
17. Once the cells on gelatin have been passaged three times and
are confluent, rinse each well with 1 mL DPBS.
Gene Editing Embryonic and Epiblast Stem Cells
91
18. Add 1 mL of DPBS into each well and scrape cells thoroughly
(see Note 32). Resuspend cells and transfer each well to a
separate 1.5 mL tube.
19. Rinse each well with another 0.5 mL of DPBS, and collect into
the same 1.5 mL tube.
20. Centrifuge samples at 1000 rpm (200 g) for 5 min.
21. Aspirate excess DPBS, removing as much as possible without
disrupting the cell pellet (see Note 33).
22. Snap freeze samples in liquid nitrogen, and either proceed to
the next step or store at 80 C until required.
23. Extract gDNA from one cell pellet by adding 100 μL of lysis
buffer with fresh Proteinase K and incubating overnight at
56 C.
24. Repeat steps 3–8 from Subheading 3.8, and then repeat Subheading 3.9 (see Note 34).
25. Confirm gene knockout by RT-qPCR and Western blotting
(see Note 35).
4
Notes
1. It is best if <10 μL plasmid is used. Aim for a plasmid stock
concentration of ~1 μg/μL.
2. It is very important that mESCs are in a single cell suspension
to reduce the number of chimeric colonies that may originate
from two or more edited cells.
3. Less cells are required for electroporation with PX458 as the
clones will be selected upon GFP expression, but negative
clones will not be eliminated. Too many cells may result in
high confluence after a few days of culture.
4. It is important that there are no air bubbles in the Neon®
pipette tip as this can cause arcing, possibly resulting in the
failure of the transfection.
5. We have determined these concentrations of puromycin to be
the best for antibiotic selection; however, it may be necessary to
perform a kill curve for different batches of puromycin.
6. mESCs require 48 h puromycin treatment, and mEpiSCs
require only 24 h puromycin treatment.
7. After puromycin treatment, many of the MEFs die which may
adversely affect mESC clone growth. To rectify this, once
puromycin treatment is complete, add additional MEFs to the
plate. This step is absolutely critical for mEpiSCs.
8. It is easier to dilute the MEFs to the required number and
volume and adding the required amount to each well rather
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than adding the cells and topping up the volume with media as
this causes uneven distribution of the cells along the bottom of
the well.
9. Colonies that have smooth edges and grow in a dome shape are
deemed suitable for selection. Colonies that don’t have smooth
edges and contain large cells that give a “rocky” appearance are
most likely differentiated and not suitable for selection. Try to
avoid selecting colonies that are substantially large, as these
may be derived from cell clumps that weren’t successfully dissociated into single cells and thus may contain multiple
mutations.
10. DPBS causes the cells to dissociate from the surface of the plate
over time, so it is important not to hold the cells in DPBS for
too long. Selecting colonies is best performed in batches. The
number of clones to pick per batch depends on how quick the
process takes. Selecting 6–8 clones per batch is a good starting
point. Between batches, DPBS is removed and replaced with
mESC or mEpiSC media and placed back in the incubator to
allow the cells to recover for 5 min. The process can then be
repeated.
11. We have selected colonies under non-sterile conditions. This is
possible if the room in which this is performed is thoroughly
cleaned and free of contamination; otherwise selecting colonies
under a microscope in a laminar flow hood would be preferred.
If performing this under non-sterile conditions, try and keep
the lid of the plates on as much as possible.
12. We normally pick clones under the 4 objective on the microscope. Find the clone you want to pick, and use the P20 pipette
to scrape it, being careful not to contaminate it with other
clones. The clones stick to the MEFs and can be difficult to
aspirate, so quite a bit of scraping may be necessary.
13. On day 3, GFP expression is quite high, but it is possible that
there aren’t enough cells to pick. By day 4, GFP expression
starts to decline but is still sufficient for visualizing the clones
for selection. Day 4 would be the last practical time point for
clone picking.
14. A mouth pipette can be generated by placing the boundary
between the thin and wide part of a 9 in. glass Pasteur pipette
under a hot Bunsen flame until it just begins to melt and pulled
such that the diameter of the tip is no larger than 1–2 mm. A
diamond cutter can be used to remove the excess thin part of
the glass. Cut 5–6 cm from the part that was pulled, and check
under a microscope whether the cut is straight. To generate the
mouth pipette, attach the tip of a filtered P1000 tip to one end
of a clear silicon tube of ~4 mm in diameter and 70 cm long and
Gene Editing Embryonic and Epiblast Stem Cells
93
a mouth piece to the opposite end. The pulled pipette can then
be attached to the P1000 tip.
15. As the clones that are selected using PX458 are small, these will
not grow to fill an entire 96-well plate. Therefore, it is necessary to trypsinize and replate the entire contents of each well
onto a new 96-well plate containing MEFs when the clones are
large enough to ensure they fill the well and do not
differentiate.
16. The morphology of the cells when grown on gelatin is not
important as they will be used to genotype the mutation only
and will not be used for future experiments. It is important that
all MEFs are removed prior to this. The best way to assess this
is by checking wells where mESC growth is minimal; if no
MEFs are visible in these wells, then most wells containing
mESCs should be clear.
17. We have previously used a hybridization oven for the overnight
incubation. We placed the 96-well plate in a plastic container
containing several wet paper towels on the bottom before
closing the lid to humidify and minimize condensation on the
top of the plate.
18. There will be condensation on the top of the plate despite the
box being humidified. Do not centrifuge the plate as this may
cause cross-contamination of samples.
19. To perform the genotyping analysis using TIDE, a PCR must
be performed on gDNA from cells that have either not been
edited or have been edited in a different region, as a wild-type
control. This allows the indels to be decomposed by comparing
the edited sequence to the unedited sequence. It is important
to ensure that the primers amplify a product that’s ~700 bp in
order for TIDE to work well. However, we have successfully
used TIDE using primers that amplify a product of ~500 bp.
20. We use HyperLadder 100 bp of which the PCR products are
within the range.
21. Sample size on gels can be up to ~50 bp different from
intended amplified fragment due to large indels. It is safe to
assume amplification is successful based on the presence of a
band within 100 bp of your intended fragment.
22. We recommend adjusting indel size range to maximum
(2–50 bp) and leaving other parameters as they are. The size
range can be adjusted if TIDE shows up an error.
23. For a standard 10 μL ligation reaction, we typically use 25 ng of
~700 bp PCR product. Ligation at room temperature for 2 h
will suffice.
24. We found that plating the entire transformation mixture may
lead to overcrowding of colonies occasionally and reduction to
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Tennille Sibbritt et al.
~250 μL reduces the chances of this occurring but also provides more than sufficient colonies to pick. It is important to
gently pipette the transformation mix out onto the plate without mixing as it will separate the already formed colonies into
small colonies or single cells; this can lead to micro- to small
colonies forming on the plate, which are difficult to pick.
25. Blue colonies tend to become more vibrant after sitting at 4 C
and therefore are more easily differentiated from the pure white
colonies.
26. Select colonies with a white background to select pure white
colonies as some colonies with small blue foci may appear
white. We find that leaving the used pipette tips in the culture
greatly improved yield of bacterial cultures the following day.
Start the mini-culture toward the end of the day to avoid
prolonged growth, which improves the quality of culture.
27. Sequences are aligned to the PCR amplicon sequence to confirm indels predicted by TIDE and/or validation of the mutation. Some sequences are the reverse complement of the
amplicon and need to be reverse-complemented before
alignment.
28. Scissors or a razor can be used to excise the required tube from
the strip; however caution should be taken as the lids are prone
to popping out when isolated.
29. Avoid agitating or moving the cell suspension onto the sides of
the tubes to ensure it settles toward to bottom for ease of
recovery.
30. It is advised to check the well under a microscope to ensure that
there are cells in the well and not in the tubes.
31. Passage from a 48-well plate to a 24-well plate and then to a
12-well plate. The first passage from the 48-well plate to a
24-well plate may not be for a week. During each set of
passage, cryopreserve cells as backups. A single confluent well
can be separated into two cryovials with each vial capable of
being thawed onto a single well of the same size it was frozen
from. The mEpiSCs take longer to recover from the thaw than
mESCs.
32. Scraping the 6-well plate may cause splatter and loss of cells.
Avoid this by gripping lower down the cell scraper and using
less force. Make sure to scrape the edges of the wells, which are
difficult. Wells can be checked under a microscope following
scraping to identify missed areas. This does not have to be
performed in a completely sterile environment and can be
performed on the lab bench.
33. We find that if we place up to two 1.5 mL tubes into a 50 mL
tube before spinning, the cell will pellet at the bottom of the
Gene Editing Embryonic and Epiblast Stem Cells
95
tube rather than along the side. This makes it easier when
aspirating DPBS and also when extracting gDNA. Tilting the
1.5 mL tube and aspirating allow for the least amount of DPBS
remaining while also avoiding disruption of the cell pellet.
34. The agarose gel electrophoresis and TIDE sequencing steps
can be skipped as the amplified product and indel are confirmed in the initial stages. If there are inconsistencies between
the confirmation and validation, a 2% agarose gel can be run
before purification and Sanger sequencing to check fragment
size and troubleshoot any changes.
35. A RT-qPCR can be performed to check the level of edited
mRNA. Design one of the primers such that they overlap the
indel, but the primers themselves are not edited. The qPCR
should show a reduction of unedited mRNA in the edited
sample. If an antibody is available, perform a Western blot to
confirm that the gene of interest is knocked out.
Acknowledgments
Our work was supported by grants from the Australian Research
Council (DP 160103651, DP 160100933), the National Health
and Medical Research Council of Australia (1127976), and the late
Mr. James Fairfax (Bridgestar Foundation). PPLT is a NHMRC
Senior Principal Research Fellow (Grant 1110751).
References
1. Urnov FD, Rebar EJ, Holmes MC, Zhang HS,
Gregory PD (2010) Genome editing with engineered zinc finger nucleases. Nat Rev Genet
11:636–646
2. Hsu PD, Zhang F (2012) Dissecting neural
function using targeted genome engineering
technologies. ACS Chem Neurosci 3:603–610
3. Ran FA, Hsu PD, Wright J, Agarwala V, Scott
DA, Zhang F (2013) Genome engineering using
the CRISPR-Cas9 system. Nat Protoc
8:2281–2308
4. Osteil P, Studdert J, Wilkie E, Fossat N, Tam PP
(2016) Generation of genome-edited mouse
epiblast stem cells via a detour through ES cellchimeras. Differentiation 91:119–125
5. Fossat N, Ip CK, Jones VJ, Studdert JB, Khoo
PL et al (2015) Context-specific function of the
LIM homeobox 1 transcription factor in head
formation of the mouse embryo. Development
142:2069–2079
6. Brinkman EK, Chen T, Amendola M, van Steensel B (2014) Easy quantitative assessment of
genome editing by sequence trace decomposition. Nucleic Acids Res 42:e168
7. Edgar RC (2004) MUSCLE: multiple sequence
alignment with high accuracy and high throughput. Nucleic Acids Res 32:1792–1797
8. Edgar RC (2004) MUSCLE: a multiple sequence
alignment method with reduced time and space
complexity. BMC Bioinformatics 5:113
Chapter 7
Identification of Circulating Endothelial Colony-Forming
Cells from Murine Embryonic Peripheral Blood
Yang Lin, Chang-Hyun Gil, and Mervin C. Yoder
Abstract
Human umbilical cord blood contains highly proliferative circulating endothelial colony-forming cells
(ECFC). These cells have promising therapeutic potential for various cardiovascular diseases by possessing
robust in vitro clonal expansion potential and the ability to form functional blood vessels in vivo upon
transplantation into recipient immunodeficient mice. However whether similar cells also exist in murine
blood remains unresolved, which impedes the study of circulating ECFC biology using murine models.
Here we describe a method to identify and culture murine embryonic peripheral blood-derived circulating
ECFC through co-culture with OP9 stromal cells. Using this method, embryonic circulating ECFC can be
identified by the formation of sheet-like or network-like endothelial colonies upon OP9 stromal cell
monolayers.
Key words Endothelial colony-forming cells, OP9 stromal cells, Circulating endothelial cells, Mouse
embryo, Peripheral blood, Endothelial progenitor cells
1
Introduction
Upon being replated on type 1 rat-tail collagen-coated plates and
cultured in complete EGM-2 medium, human circulating endothelial colony-forming cells (ECFC) from umbilical cord blood and
peripheral blood can attach to the tissue culture plates and form
robust endothelial colonies [1, 2]. Cells from circulating ECFC
colonies possess a hierarchy of clonal proliferative potential and can
form long-term functional blood vessels in vivo after transplantation into immunodeficient mice [1, 2]. Thus, circulating ECFC
hold great promise as a cell therapy for the treatment of cardiovascular diseases. However, whether an equivalent cell type also exists
in murine peripheral blood has not been resolved, which hinders
the study of circulating ECFC using various transgenic mouse
models and thus leaves many questions about the origin, production, and function of circulating ECFC unanswered. Though some
earlier studies have reported the identification of circulating cells
Ivan Bertoncello (ed.), Mouse Cell Culture: Methods and Protocols, Methods in Molecular Biology, vol. 1940,
https://doi.org/10.1007/978-1-4939-9086-3_7, © Springer Science+Business Media, LLC, part of Springer Nature 2019
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Yang Lin et al.
that exhibit some endothelial cell (EC) features in vitro and named
these cells “endothelial progenitor cells” (EPC) [3, 4], it was soon
discovered that essentially all of the bone marrow EPC identified in
these studies were in fact a heterogeneous mixture of hematopoietic
stem and progenitor cells that may resemble endothelial cell phenotypes upon in vitro culture with growth factors but lack the
ability to form functional blood vessels in vivo [5–7]. To our
knowledge, there has only been one prior report that used stringent
circulating ECFC culture methods to grow murine circulating
blood-derived proliferative EC [8]. However this article pointed
out that murine circulating ECFC were so rare that only 1 ECFC
could be detected from the combined blood volume of over
50 adult mice [8].
By applying an OP9 stromal cell co-culture system that has
been widely used to grow murine EC colonies [9–12], we have
developed a method to identify circulating ECFC from murine
embryonic peripheral blood. Using this method, murine embryonic circulating ECFC can be identified by the formation of EC cell
surface marker-expressing, sheet-like, or network-like EC colonies
on OP9 stromal cell monolayers after 7–10 days of culture.
2
Materials
Prepare all medium and solutions using deionized ultrapure water.
All culture medium should be stored at 4 ˚C and replaced by fresh
medium every 4 weeks. All reagents should be stored according to
the manufacturer’s instructions.
2.1
Equipment
1. 1 L glass beaker.
2. pH meter.
3. 1 L graduated cylinder.
4. Tissue culture hood.
5. Pipettes.
6. 0.22 μm filter membranes.
7. Vacuum pressure system.
8. Humidified 37 ˚C incubator in room air with 5% CO2.
9. Cell culture centrifuge.
10. T25 culture flasks.
11. 14 mL and 50 mL conical tubes.
12. 6-well culture plates.
13. 12-well culture plates.
14. 37 ˚C water bath.
15. Surgical scissors.
Culture of Circulating ECFC
99
16. 6 cm petri dish.
17. Dissection microscope.
18. 70 μm nylon filter.
2.2
Reagents
1. Basal Minimum Essential Medium Alpha medium (MEMα):
For 1 L basal medium, dissolve 10 g Minimum Essential
Medium Alpha medium powder (Gibco) in 950 mL water in
a beaker at room temperature. Add 2.2 g sodium bicarbonate
to the solution; stir the mixture until all the powders are dissolved. Adjust pH to 7.2–7.4 with 1 N HCL or 1 N NaOH.
Adjust the final volume to 1 L with water in a graduated
cylinder (see Note 1). Filter the medium through a 0.22 μm
filter membrane into a sterile container using a vacuum pressure system.
2. OP9 medium: Mix 79.5% basal MEMα medium with 20% fetal
bovine serum (FBS; see Note 2) and 0.5% 10, 000 U/mL
penicillin streptomycin (Pen Strep) solution (final concentration: 50 U/mL). Filter the medium through a 0.22 μm filter
membrane.
3. 0.1 M 2-mercaptoethanol stock solution: Add 70 μL
2-mercaptoethanol to 10 mL basal MEMα medium. Filter
the mixture through a 0.22 μm filter membrane (see Note 3).
4. EC culture medium: Mix 89.5% basal MEMα medium, 10%
FBS (see Note 4), 0.5% Pen Strep, and 1: 2000 0.1 M
2-mercaptoethanol stock solution (final concentration:
5 10 5 M). Filter the medium through a 0.22 μm filter
membrane.
5. Heparin stock solution: Dissolve heparin sodium salt powder
into the water to make a 2000 U/mL stock solution. Filter the
solution through a 0.22 μm filter membrane (see Note 5).
6. Blood collection buffer: For every 100 mL buffer, add 0.5 mL
Pen Strep solution, 1 mL FBS, and 100 μL 2, 000 U/mL
heparin stock solution (final concentration: 2 U/mL) into
sterile PBS (see Note 6).
7. Phosphate buffered saline (PBS): PBS can be purchased (e.g.,
Sigma-Aldrich or Gibco) or can be formulated as follows. Add
900 mL water to a 1 L beaker. Add 8 g sodium chloride (NaCl,
final concentration 137 mM). Add 0.2 g potassium chloride
(KCl, final concentration 2.7 mM). Add 1.44 g disodium
hydrogen phosphate (Na2HPO4, final concentration
10 mM). Add 0.24 g monopotassium phosphate (KH2PO4,
final concentration: 1.8 mM). Adjust the pH to 7.4 with HCl,
and then transfer the solution in the beaker to a graduated
cylinder. Adjust the volume to 1 L with water. Aliquot the PBS
into 500 mL bottles and autoclave.
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Yang Lin et al.
8. 0.25% trypsin-EDTA.
9. 70% ethanol.
10. Red blood cell lysis buffer.
2.3 Cells
and Animals
3
1. OP9 stromal cells.
2. Male and female C57/BL6 mice to prepare day 10.5–day 12.5
embryos.
Methods
OP9 stromal cells and circulating ECFC should be cultured in a
humidified 37 ˚C incubator in room air with 5% CO2. Unless
otherwise noted, all experimental procedures should be carried
out at room temperature.
3.1 OP9 Stromal Cell
Maintenance
Maintain OP9 stromal cells by culturing on T25 culture plates in
OP9 medium (4 mL medium/T25 plate; see Note 7 and Fig. 1).
When the cultures reach 80~90% confluence, the cells need to be
passaged following these steps in a tissue culture hood:
1. Wash the cultured cell monolayers two times with sterile PBS
(see Note 8).
2. Add 1 mL 0.25% trypsin-EDTA to the T25 flasks. Keep the
culture in a humidified 37 ˚C incubator in room air with 5%
CO2 for 3 min.
3. Take out the culture from the incubator. Add 3 mL OP9
medium to neutralize the trypsin. Pipette 5–10 times to wash
Fig. 1 Morphology of OP9 stromal cells. (a) Normal OP9 culture with 80–90% confluency that is ideal for
co-culture of endothelial cells. Cells show a polygonal morphology. (b) When OP9 stromal cells are cultured for
a longer period of time or cultured in suboptimal conditions, the cell growth rate starts to increase, and cells
start to transform into an elongated morphology, and the cells will overgrow and become intensely packed;
OP9 cultures should be discarded upon reaching this state. Scale bar, 100 μm
Culture of Circulating ECFC
101
off the cells from the bottom of the flasks. Transfer the mixture
of cells, medium, and trypsin-EDTA from each T25 flask into a
14 mL conical tube.
4. Centrifuge the cells at 300 g for 5 min.
5. Re-suspend the cells from each 80–90% confluent T25 flask in
12 mL OP9 medium. Plate the cells into three new T25 flasks
(4 mL/T25). Shake the flasks well to evenly disperse the cells
(see Note 9). Culture the cells in a humidified 37 ˚C incubator
in room air with 5% CO2.
3.2 Preparing OP9
Plates for EC
Co-culture
OP9 plates should be prepared 24 h or 48 h before the co-culture.
1. Collect and centrifuge OP9 stromal cells as previously
described.
2. Re-suspend OP9 stromal cells in OP9 medium, and replate the
cells in 6-well or 12-well culture plates. If the plates were
prepared 24 h before the EC co-culture experiment, OP9
stromal cells from one 80–90% confluent T25 flask should be
re-suspended in 6 mL medium and plated in 3 wells of a 6-well
plate (2 mL/well) or 6 wells of a 12-well plate (1 mL/well). If
the plates were prepared 48 h before the initiation of the
co-culture, OP9 stromal cells from one T25 flask should be
re-suspended in 12 mL medium and replated in one 6-well
culture plate (2 mL/well) or one 12-well culture plate (1 mL/
well).
3. Tap the plates to evenly disperse the cells. Culture the cells
in a humidified 37 ˚C incubator in room air with 5% CO2
(see Note 10).
3.3 Collection
of Embryonic Blood
This procedure can be performed on a lab bench. Surgical tools
should be sterilized (autoclaved) before the experiment to reduce
the chance of contamination. Euthanized pregnant mice need to be
sprayed with 70% ethanol before opening the skin.
1. Pre-warm PBS and blood collection buffer at 37 ˚C in a water
bath (see Note 11). Prepare 10 mL buffer for each embryo
collected.
2. Euthanize day 10.5–day 12.5 pregnant C57/BL6 female mice
through cervical dislocation (see Note 12).
3. Pinch the abdominal skin with a pair of forceps, and make a
lateral incision at the midline with surgical scissors. Pull the skin
to expose the peritoneum. Open the peritoneum with scissors
to expose the peritoneal cavity.
4. Remove the uterus from the peritoneal cavity. Transfer the
uterus into warmed PBS in a 6 cm petri dish.
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Yolk Sac
Placenta
Umbilical
Cord
Embryo
Proper
Vitelline
Cord
Fig. 2 Illustration of an embryonic day 12.5 embryo. Yolk sac should be opened
along the indicated straight line. Dashed line indicates the site to cut umbilical
and vitelline vessels
5. Under a dissection microscope, remove the muscle layer of the
uterus to collect each embryo (see Note 13).
6. Wash off maternal blood from each embryo by briefly rinsing
the embryos in PBS. Place individual embryos in a 6 cm petri
dish with 4–6 mL warm blood collection buffer. Make an
incision on the yolk sac (cut along the straight line indicated
on Fig. 2) to expose the vitelline and umbilical blood vessels.
Cut the cord to open vitelline and umbilical vessels (cut at the
dashed line in Fig. 2). Embryonic blood will be pumped out
from the embryo with each heartbeat (see Note 14) until the
embryo is exsanguinated and turns white.
7. Transfer the blood containing petri dishes to a tissue culture
hood. Aspirate the blood solution using a 5 mL pipette, and
pass through a 70 μm nylon filter into a 14 mL conical tube.
3.4 Circulating ECFC
Culture
In this step, murine embryonic mononuclear cells will be plated on
OP9 stromal cells to culture embryonic circulating ECFC.
1. Centrifuge the blood cells collected from murine embryos
(Subheading 3.3) at 300 g for 5 min. Re-suspend the cells
from each embryo in 5 mL red blood cell lysis buffer (see Note
15). Incubate the cells for 5 min at room temperature. Add
10 mL EC culture medium to dilute the red blood cell lysis
buffer. Centrifuge the cells at 300 g for 5 min. Re-suspend
the blood mononuclear cell pellet from each embryo into 2 mL
EC culture medium.
2. Carefully remove the medium from OP9 plates prepared in
Subheading 3.2. Disperse the suspension of blood mononuclear cells collected from each embryo onto OP9 stromal
cells in 1 well of a 6-well plate or 2 wells of a 12-well plate
(see Note 16). Gently tap the plates to evenly disperse the cells.
Culture of Circulating ECFC
103
Fig. 3 A network-like (left panel) and a sheet-like (right panel) embryonic day 12.5 murine circulating ECFCderived endothelial colony after 10-day co-culture with OP9 stromal cells. The cultures were fixed with 4%
paraformaldehyde and stained with rat anti-mouse CD31 primary antibody and Alexa Fluor 488-conjugated
anti-rat IgG secondary antibody. Scale bar, 100 μm
Incubate the co-cultured cells in a 37 ˚C incubator in room air
with 5% CO2.
3. Gently (see Note 16) remove the spent medium, and add fresh
medium after 24 h to remove nonadherent cells. Change
medium every 24 h afterward. After 7–10 days culture,
sheet-like or network-like circulating ECFC-derived colonies
can be identified in the cultures (Fig. 3). Cultured murine
embryonic circulating ECFC colonies can be maintained for
up to 3 weeks (see Note 17). After fixing the cultures with 4%
paraformaldehyde at room temperature for 10 min and washing the fixed cultures three times with PBS, these colonies can
be visualized by immunohistochemistry/immunofluorescence staining using antibodies against EC surface markers
(Fig. 3; see Note 18).
4
Notes
1. If the MEMα powder contains phenol red, the color of freshly
made medium should be red. The turning of color into yellow
or pink is an indication that the pH of the medium has
changed. In that case, new medium should be prepared, and
old medium should be discarded. We use MEMα powder from
Gibco to prepare MEMα medium, while other forms of MEMα
medium from other vendors may also be suitable for this
experiment. If you use other suppliers, please follow the suppliers’ instructions while preparing MEMα medium.
2. FBS selection is crucial for culturing OP9 stromal cells. Different lots of FBS from different vendors should be tested by
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making test OP9 medium and comparing with known optimal
FBS for culturing OP9 stromal cells. If a medium contains
optimal FBS, OP9 can be passaged for 10 passages in
3–4 weeks while retaining their polygonal shape (Fig. 1a).
Currently we are using FBS from Atlanta Biologicals (cat.
S11550. Lot. L14148). (OP9 stromal cells were obtained
from Dr. Yoshimoto Kobayashi at the University of Texas
Health Science Center Houston, Houston, TX).
3. 2-Mercaptoethanol is crucial for the culture of murine
EC. Fresh 0.1 M 2-mercaptoethanol stock solution should be
made every 3–4 weeks and stored at 4 ˚C. 2-Mercaptoethanol
has strong smell and is hazardous in case of inhalation and skin
or eye contact. Always open pure 2-mercaptoethanol bottle in a
fume hood to make a stock solution.
4. Compared with the FBS for OP9 culture, the choice of FBS for
EC co-culture is less stringent. Currently we are using defined
FBS from HyClone (cat. SH30070.03. Lot. AWC10533).
5. Heparin is important for preventing the blood from coagulating. Heparin stock solution should be kept at 4 ˚C, and new
stock solution should be made every 4 weeks. We make heparin
stock solution by dissolving heparin salt powder from SigmaAldrich with water, while commercially available ready-made
heparin stock solutions can also be purchased.
6. For every experiment, blood collection medium needs to be
freshly made at the same day of the blood collection.
7. The quality of OP9 stromal cells is crucial for the co-culture of
murine EC. Normal OP9 stromal cells display a polygonal
morphology (Fig. 1a) and a controlled proliferation rate
(after being split at 1:3 ratio, the cells in culture reach 90%
confluence at day 3). When OP9 stromal cells transform into an
elongated spindle shape (Fig. 1b) and start growing at an
accelerated rate, the cells need to be discarded, and new stromal
cells should be thawed and cultured. Normally, a freshly
thawed OP9 culture can be passaged for up to 4 weeks without
compromising their quality. If OP9 stromal cells are grown in
medium with suboptimal FBS, they can be passaged for a
shorter period of time before their morphology starts to
change (2–3 weeks). While culturing OP9 stromal cells, the
color of medium should be closely monitored. Whenever the
medium turns pink instead of red, the medium in the OP9
cultures should be replaced by fresh medium.
8. FBS in the culture medium can neutralize trypsin activity. Thus
the cultures need to be washed with PBS to remove residual
FBS containing medium prior to effective release of the adherent cells. When washing OP9-cultured cells, add PBS gently
against the wall of the tissue culture flasks, briefly rinse the cells,
Culture of Circulating ECFC
105
and then carefully aspirate the PBS with glass pipettes
connected to a vacuum source to avoid inadvertent aspiration
of the cells.
9. While replating OP9 stromal cells, the cell suspension should
be evenly distributed to prevent differentiation caused by cell
aggregation. The normal growth of OP9 stromal cells requires
proper air ventilation. If the caps of the T25 culture flasks are
not filtered, loosen the caps during culture to allow air
exchange.
10. It is recommended to co-culture EC with 80–90% confluent
OP9 stromal cells. OP9 plates that have reached 90% confluency may still be usable for EC co-culture after 1–2 days,
but the size of EC colonies might be affected.
11. To avoid blood coagulation, embryonic blood needs to be
collected in warm PBS buffer rather than cold buffer.
12. Other than C57/BL6, other mouse strains like SV129, CD1,
and FVB can also be used. To prevent embryo death in utero
and blood coagulation, it is crucial to perform the blood
collection process as quickly as possible. Thus, euthanizing
the pregnant dams through cervical dislocation is preferred
compared with other slower euthanization methods like using
a CO2 chamber.
13. Due to the small size of murine embryos, it is recommended to
perform the following steps under a dissection microscope and
use two pairs of sharp tip watchmaker’s forceps (one with
straight tip, one with bent tip) and a pair of straight sharp tip
iris scissors.
14. To collect more blood from the embryos, it is important to
perform the previous dissection steps quickly so the embryonic
murine hearts can keep beating as the major driver to cause
exsanguination. If the umbilical and vitelline vessels are
clogged by coagulation during the collection process, a second
cut can be made proximal toward the embryonic side of
the cord.
15. Red blood cells can affect the growth of OP9 stromal cells and
EC. So it is important to remove them from the culture. If red
blood cells are successfully lysed, the buffer will turn from
transparent to a light red color. After the treatment of red
blood lysis buffer, some red blood cells, especially enucleated
primitive erythroid, will still remain but will be removed by the
repeated medium change steps after establishing the
co-cultures.
16. While adding medium to a culture plate with OP9 stromal
cells, place the pipette tip against the wall of each well, and
add the medium drop by drop to avoid breaking the OP9
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monolayer. Do not leave the OP9 stromal cell monolayer to
dry for more than 30 s, or this will result in loss of support for
the EC colonies, and there is a greater chance for release of the
entire OP9 monolayer from the plate.
17. Some embryonic hematopoietic progenitor-derived colonies
will also grow on the OP9 monolayer. However EC colonies
show unique sheetlike or network-like morphology (Fig. 3),
express EC-specific markers like CD31 (Pecam1) and CD144
(VE-cadherin), and do not express hematopoietic specific markers like CD45 and CD11b and thus can be distinguished from
hematopoietic colonies via immunohistochemistry or immunofluorescent staining (Fig. 3). Normally, 1–3 circulating
ECFC colonies can be derived from the blood volume of
each embryonic day 10.5–11.5 murine embryo, while 10–20
circulating ECFC colonies are expected to be identified from
each embryonic day 12.5 embryo. After prolonged culture,
some OP9 stromal cells will differentiate into oval shape adipocytes. However these adipocytes will not affect the growth
of EC colonies.
18. Even after fixation, the ECFC-OP9 co-culture should still be
handled gently because the cell monolayers are easy to peel off
from the bottom of the culture plates. It is recommended to
always add buffer or solution drop by drop to the center of the
wells while processing the cultures.
References
1. Ingram DA, Mead LE, Tanaka H, Meade V,
Fenoglio A et al (2004) Identification of a
novel hierarchy of endothelial progenitor cells
using human peripheral and umbilical cord
blood. Blood 104:2752–2760
2. Javed MJ, Mead LE, Prater D, Bessler WK,
Foster D et al (2008) Endothelial colony forming cells and mesenchymal stem cells are
enriched at different gestational ages in
human umbilical cord blood. Pediatr Res
64:68–73
3. Asahara T, Kawamoto A, Masuda H (2011)
Concise review: Circulating endothelial progenitor cells for vascular medicine. Stem Cells
29:1650–1655
4. Asahara T, Murohara T, Sullivan A, Silver M,
van der Zee R et al (1997) Isolation of putative
progenitor endothelial cells for angiogenesis.
Science 275:964–967
5. Rehman J, Li J, Orschell CM, March KL
(2003) Peripheral blood “endothelial progenitor cells” are derived from monocyte/
macrophages and secrete angiogenic growth
factors. Circulation 107:1164–1169
6. Yoder MC (2013) Endothelial progenitor cell:
a blood cell by many other names may serve
similar functions. J Mol Med 91:285–295
7. Medina RJ, Barber CL, Sabatier F, DignatGeorge F, Melero-Martin JM et al (2017)
Endothelial progenitors: a consensus statement
on nomenclature. Stem Cells Transl Med
6:1316–1320
8. Somani A, Nguyen J, Milbauer LC, Solovey A,
Sajja S, Hebbel RP (2007) The establishment
of murine blood outgrowth endothelial cells
and observations relevant to gene therapy.
Transl Res 150:30–39
9. Hirashima M, Kataoka H, Nishikawa S,
Matsuyoshi N, Nishikawa S (1999) Maturation
of embryonic stem cells into endothelial cells in
an in vitro model of vasculogenesis. Blood
93:1253–1263
10. Hashimoto K, Fujimoto T, Shimoda Y,
Huang X, Sakamoto H, Ogawa M (2007)
Culture of Circulating ECFC
Distinct hemogenic potential of endothelial
cells and CD41+ cells in mouse embryos.
Develop Growth Differ 49:287–300
11. Naito H, Kidoya H, Sakimoto S,
Wakabayashi T, Takakura N (2012) Identification and characterization of a resident vascular
107
stem/progenitor cell population in preexisting
blood vessels. EMBO J 31:842–855
12. Naito H, Wakabayashi T, Kidoya H,
Muramatsu F, Takara K et al (2016) Endothelial side population cells contribute to tumor
angiogenesis and antiangiogenic drug resistance. Cancer Res 76:3200–3210
Chapter 8
Imaging and Analysis of Mouse Embryonic Whole Lung,
Isolated Tissue, and Lineage-Labelled Cell Culture
Matthew Jones and Saverio Bellusci
Abstract
Research on lung development and disease frequently utilizes mouse models to conduct in vitro experiments. Such experiments involve multiple methodologically distinct stages, from careful consideration of
mouse models used to obtain biological samples, to the culturing and imaging of those samples, and finally,
to post-imaging analysis. Here, we detail basic protocols to assist with each of these stages. First, we discuss
harvesting and preparing biological samples; second, we focus on culturing embryonic whole lung explants
and isolated mesenchyme and epithelium; third, we specify the basics of obtaining still and live images; and
finally, we bring these methods together by considering and briefly analyzing a lineage-labelling
experiment.
Key words Lung explant, Isolated epithelium, Isolated mesenchyme, Organ culture, Tissue culture,
Lineage tracing, Still imaging, Live imaging, Image analysis
1
Introduction
Research on the molecular mechanisms regulating lung development, homeostasis, and disease often involves using wild-type and
genetically modified mouse models and conducting in vitro experiments to test tentative ideas or to compliment in vivo findings.
While results obtained in vitro must always be interpreted with care,
a well-designed, properly conducted in vitro experiment does permit the researcher to confidently address particular hypotheses that
would be prohibitively difficult, or impossible, in vivo. See refs.
[1–3] for reviews on the use of in vitro models to study lung
development and disease.
For instance, questions concerning the local effects of a molecule of interest, such as a pharmacological inhibitor, are routinely
tested using mouse-based in vitro models. In vitro pharmacological
intervention, often using a reporter-based model to label and trace
cells of interest, combined with image analysis to quantify morphological effects, can lead to innovative hypotheses. These hypotheses
Ivan Bertoncello (ed.), Mouse Cell Culture: Methods and Protocols, Methods in Molecular Biology, vol. 1940,
https://doi.org/10.1007/978-1-4939-9086-3_8, © Springer Science+Business Media, LLC, part of Springer Nature 2019
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Matthew Jones and Saverio Bellusci
may eventually transfer to research in vivo and the testing of drugs
and other molecules to treat disease. See refs. [4–6] for reviews on
the use of in vitro models in the biomedical field and the difficulties
in translating in vitro to in vivo studies.
In vitro experiments exploring lung development and disease
involve multiple methods: obtaining biological samples (such as
whole lung explants or primary cells) from embryonic mice of
known age; culturing samples, often in chemically defined medium
supplemented with molecules (such as growth factors), the
biological roles of which are under investigation; acquiring valuable
phenotypic data via high-quality imaging of samples, which enables
one to capture large-scale phenotypic differences between samples
over time (such as changes in morphology); and finally, the postacquisition analysis and quantification of images. Furthermore,
fluorescence live imaging is frequently used to identify and quantify
the expression of fluorescent proteins in living tissue, providing
insight into the activity of cells and proteins in real time.
Given that in vitro experiments routinely involve multiple
methodological stages, it is critical to have a well-developed, precise, and concise set of protocols to follow. Clearly, errors and poor
practice at any stage from obtaining biological samples to analyzing
experimental data will negatively impact results and interpretation.
In this chapter, we describe the basic steps routinely employed
in our lab to conduct in vitro experiments, from obtaining
biological samples to analyzing data in the form of images. These
steps are organized into four sections: first, obtaining and preparing
biological samples from mice; second, the culturing of whole lung
explants and isolated mesenchyme and epithelium; third, the essentials of taking good, high-quality images, both still and live; and
finally, a brief section applying these steps to an actual experiment:
the labelling, live imaging, and analysis of cells in embryonic whole
lung explants.
2
Materials
Prepare, aliquot, and store all reagents according to manufacturer’s
instructions. Prepare fresh culture medium for each experiment,
and avoid repeated freeze-thaw cycles for all reagents.
2.1 Mice
and Euthanasia
1. Timed-pregnant mice (wild-type or genetically modified) sacrificed at desired postcoitum embryonic stage (E), where E0.5 is
assumed to be noon on the day a vaginal copulation plug is
found.
2. Narcoren® pentobarbital sodium (16 g/100 mL) dissolved in
0.9% sterile sodium chloride to make a 25% working solution
(40 mg/mL).
Embryonic Whole Lung and Tissue Culture: Imaging and Analysis
111
3. BD™ 24-26G Microlance™ three needles and Braun™
0.01–1 mL Injekt™-F syringes.
4. 50 mL Falcon® tubes.
5. Sterile phosphate-buffered saline (PBS) (1), pH 7.2.
2.2 Embryo
Harvesting and Lung
Dissection
1. Dissecting tools: Student fine scissors, student vannas spring
scissors, Dumont® #5 and #5CO stainless steel forceps, and a
small Moria® perforated spoon.
2. Culture medium used to cover embryo during lung dissection
and to incubate lungs once dissected (see Note 1). Medium
contains Dulbecco’s Modified Eagle Medium (DMEM) (1),
supplemented with D-glucose, L-glutamine, HEPES, pyruvate,
and phenol red, 10% fetal bovine serum, and 1% penicillin
(10,000 units/mL)-streptomycin (10 mg/mL).
3. 60 and 92 mm polystyrene Petri dishes.
4. Leica MZ 125 stereoscopic dissecting microscope.
5. Laminar airflow workstation.
6. 40 μL and 100 μL calibrated micropipets with aspirator tube
assembly.
2.3 Separation
of Mesenchyme
and Epithelium
1. Dispase, a neutral metalloprotease derived from Bacillus polymyxa. Aliquot and store at 20 C.
2. Fetal bovine serum (FBS) (ATCC, Wesel, Germany). Sterile,
not heat inactivated. Aliquot and store at 20 C.
3. Culture medium used to incubate separated epithelium and
mesenchyme (see Note 1). Medium contains Dulbecco’s Modified Eagle Medium (DMEM) (1), supplemented with Dglucose, L-glutamine, HEPES, pyruvate, and phenol red, 10%
fetal bovine serum, and 1% penicillin (10,000 units/mL)-streptomycin (10 mg/mL).
4. Sterile polystyrene culture dishes (e.g., 24-well culture plate).
5. Tungsten carbide microdissection needles with a tip diameter
of 0.001 mm and length of 1.2 cm.
6. Leica MZ 125 stereoscopic dissecting microscope.
7. Laminar airflow workstation.
8. 40 μL and 100 μL calibrated micropipets with aspirator tube
assembly.
2.4 Whole Lung
Explant and Isolated
Epithelium
and Mesenchyme
Culture
1. General-purpose culture medium: Medium contains Dulbecco’s Modified Eagle Medium (DMEM) (1), supplemented
with D-glucose, L-glutamine, HEPES, and pyruvate, with or
without phenol red (see Note 2), 10% fetal bovine serum, and
1% penicillin (10,000 units/mL)-streptomycin (10 mg/mL).
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Matthew Jones and Saverio Bellusci
2. Serum-free, chemically defined culture medium: Medium contains Dulbecco’s Modified Eagle Medium/Nutrient Mixture
F-12 (DMEM/F-12) (1), supplemented with L-glutamine,
HEPES-free, with or without phenol red (see Note 2), and 1%
penicillin (10,000 units/mL)-streptomycin (10 mg/mL).
3. Stainless steel metallic grids, constructed with 0.75 mm thick,
0.38 mm diameter stainless steel wire mesh (Cat. # FE248710,
Goodfellow) (see Note 3 for details on constructing grids).
4. Sterile polystyrene culture dishes (e.g., 24-well culture plate).
5. Sterile phosphate-buffered saline (PBS) (1), pH 7.2.
6. Graefe forceps.
7. Whatman® Track-Etch™ membranes, 13 mm in diameter with
an 8.0 μm pore size.
8. Laminar airflow workstation.
9. Leica MZ 125 stereoscopic dissecting microscope.
10. 40 μL calibrated micropipets with aspirator tube assembly.
11. Heracell™ 150 incubator.
12. Matrigel® growth factor reduced basement membrane matrix,
phenol red-free. Store at 20 C.
2.5 Still Imaging
Using Bright-Field
Microscopy
1. Leica MZ 125 stereoscopic dissecting microscope, equipped
with a Schott KL 1500 LED light source and a Spot™
Insight™ 2.0 Mp Color Mosaic camera.
2. Spot™ 4.5.9 imaging software for Mac computers.
2.6 Live Imaging
Using Fluorescent
Microscopy
Leica AF6000 Integrated System for Live Cell Imaging and Analysis, which includes a DM6000B fluorescent inverted microscope,
CTR6000 electronics box, DFC 305FX camera, heating and CO2
control units, climate control chamber, and a designated computer
and software.
2.7 Lineage
Labelling and Tracing
Using a CreERT2/LoxPSTOP-LoxP Reporter
System
Only materials that differ from those previously mentioned are
included here.
1. Mouse Strains
(a) A CreERT2 driver line where Cre is under transcriptional
control of a gene of interest. In our example, we use
Fgf10CreERT2/+ mice generated in our lab, as described in
El Agha et al. [7].
(b) An inducible LoxP-STOP-LoxP reporter line that will
express a fluorescent protein after recombination with Cre
recombinase. In our example, we use TomatoRFP flox/flox
mice obtained from Jackson Laboratories (Stock #
007908, Jackson Laboratory), which expresses a red fluorescent protein (RFP) after recombination [8].
Embryonic Whole Lung and Tissue Culture: Imaging and Analysis
113
2. Induction of System to Label Cells: Tamoxifen stock solution
dissolved in corn oil at room temperature to make a 20 mg/mL
working solution. Aliquot working solution (approximately
500 μL per aliquot) and store at 20 C.
3. Harvesting, Culturing, and Manipulating Samples: Supplementary molecules added to medium. In our example we add
vivo-morpholinos to inhibit the activity of target mRNA.
4. Imaging and Image Analysis: Leica MM AF image analysis
software powered by MetaMorph®. ImageJ 1.50i open-source
imaging software (to download desired version, see http://
imagej.nih.gov/ij).
3
Methods
3.1 Preparation
of Biological Samples
All steps involving embryo harvest, lung dissection, and mesenchymal and epithelial isolation should be performed in sterile conditions, in a laminar flow workstation, using a dissecting stereoscope.
All steps are carried out at room temperature, unless otherwise
noted.
1. Consideration of Mice and Embryonic Stage: Mice should be
carefully selected for each experiment, taking into account
factors such as genetic background and age, which may have a
bearing on the experimental outcome (see Note 4). Furthermore, while the following steps have been described in general
terms, we are most experienced using these methods with
E12.5-E16.5 embryos with a C57BL/6J genetic background.
The steps, therefore, may need to be optimized for other
mouse strains and models.
2. Euthanasia and Harvesting Embryos: The established method
to euthanize animals may differ from the one detailed here.
Make sure to follow all local and national laws concerning the
ethical use of animals for scientific purposes.
(a) Timed-pregnant females are sacrificed by an overdose
of pentobarbital administered intraperitoneally (0.1 mL
working solution/10 g mouse weight). Confirm
death by ensuring absence of pupil response to light and
absence of leg reflexes by pinching between the toes of
the foot.
(b) To remove the uterus containing the embryos, lay the
female on her back, spray her abdomen with 70% ethanol,
make an incision at the base of her abdomen, pull the skin
upward while holding the hind legs, open the peritoneal
cavity, and dissect and remove the uterus.
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(c) Wash the uterus by placing it in a 50 mL Falcon® tube
containing sterile PBS and gently rocking it for approximately 2 min.
(d) To remove the embryos, place the rinsed uterus in a Petri
dish containing sterile PBS. Under a dissecting stereoscope, incise the uterine wall using forceps or fine scissors.
Gently remove the embryos from the uterus, and sever
their umbilical cords. Using a small perforated spoon,
transfer each embryo to a new Petri dish containing culture medium.
3. Whole Lung Dissection: Under a dissecting stereoscope, harvest lungs at room temperature in Petri dishes containing
culture medium.
(a) To immobilize embryo, lay the embryo on its right flank
(for left-handed researchers, reversing the orientation
described herein might be helpful). Completely cover
the embryo with medium (Fig. 1a) (see Note 5). Working
with fine-tipped forceps in the left hand, pin the embryo
with one of the forceps tips through its abdomen at the
base of its hind limb, and pin its head with the other tip.
Avoid piercing any internal organs. Apply gentle pressure
against the bottom of the dish, and maintain a steady
hand. In this way, the embryo will be immobilized during
the dissection (Fig. 1b).
(b) To harvest lungs, use fine-tipped forceps in the right hand;
remove the fore and hind limbs at their base on the left
side of the embryo (Fig. 1c). Using the tips of the forceps
like scissors (or spring scissors for late-stage embryos),
gently cut through the skin and the ribs of the embryo
from the midline of the lower abdomen to the site of the
removed hind limb, to the base of the rib cage (where ribs
meet the spine), up the spine toward the site of the
removed forelimb, through the clavicle, and finally
through the top of the trachea (see Note 6). This semicircular incision will allow the separation of the internal
organs from the embryonic skeleton and will grant easy
access to the lungs, which are located on the dorsal side of
this internal mass (Fig. 1d).
(c) To separate the internal organs, it is easiest to work with
both sets of forceps. With the tips of the forceps, gently
peel and pull apart the two halves of the incision (Fig. 1e).
Ideally, the lung—often still attached to other organs such
as the heart—should be completely removed from the
other organs and the rest of the body at this stage
(Fig. 1f).
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Fig. 1 Overview of embryonic lung dissection using E16.5 embryo. (a) Embryo placed on its right flank and
covered with medium. (b) Placement of forceps to immobilize embryo, making sure to avoid internal organs
(white bar). (c) Removal of hind and forelimbs (white arrows). (d) Semicircular incision revealing internal
organs (black dashed line). (e) Separation of organs from the body. Note the location of the left lung lobe (white
arrow). (f) Removal of internal organs. (g) Removing unwanted organs from lungs, such as the heart (white
arrow). (h) Removing excess and unwanted tissue. (i) Cleaned lung with trachea intact, ready for culturing.
Scale bars, 2.5 mm (a–h); 2 mm (i)
(d) Once isolated from the rest of the body and the majority
of the other internal organs, the lung can be cleaned of
any unwanted organ tissue (such as the heart) and of any
residual connective tissue (Fig. 1g). To remove unwanted
tissue, use both sets of forceps to gently remove tissue
attached to the lung mesothelium and trachea, making
sure not to pierce the lobes. It is easiest to accomplish this
step by gently pinning the lung to the bottom of the Petri
dish by the trachea and pulling excess tissue away from the
lobes (Fig. 1h). Once free from residual tissue, the dissected lungs are ready to be cultured (Fig. 1i) (see Note 7).
4. Separation of Epithelium and Mesenchyme: The following
protocol is modified from the protocol described by del
Moral and Warburton [9].
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(a) Take a suitably sized culture dish (we use a 24-well culture
plate), and add 500 μL undiluted dispase to each well,
which will contain dissected lung samples.
(b) Transfer the isolated lungs to the dispase, and incubate on
ice for 20–30 min (the time needs to be optimized here as
the dispase activity may vary depending on experimental
and lab conditions).
(c) Prepare empty wells (on the same plate if possible) by
adding 500–1000 μL pure FBS to each well. Transfer
the dispase-digested samples to the pure FBS, and incubate on ice for 15 min. This blocks the dispase enzymatic
activity.
(d) Transfer samples to new wells containing culture medium,
and keep on ice.
(e) To mechanically dissociate epithelium from mesenchyme,
take a single lung, and transfer it to a Petri dish containing
culture medium. Under a dissecting stereoscope, use
tungsten microdissection needles to gently separate the
mesenchyme from the epithelium.
(f) Use a calibrated micropipet and aspirator tube to mouth
pipet the separated epithelium and mesenchyme to
respective wells containing culture medium on ice. The
tissue explants are now ready for culturing.
3.2 Whole Lung
Explant and Isolated
Epithelium
and Mesenchyme
Culture
The following steps must be performed under sterile conditions,
including properly sterilized tools (i.e., autoclaved), as contamination is the main concern with culturing.
1. Whole Lung Explant Culture
(a) Depending on number of samples, the culture dish used
will vary. Prepare a suitable culture dish by adding to each
sample-containing well a metallic grid with a hole
punched in the middle (see Note 8). Then add an amount
of culture medium suitable to barely cover the grid. In the
other wells, add sterile PBS to maintain proper humidity
in the dish once the dish is closed.
(b) Using a pair of sterile forceps, place a Track-Etch™ membrane, shiny side down, atop the medium of each sample
well. Avoid trapping bubbles under the membrane.
(c) Using a calibrated micropipet, transfer dissected lungs to
the center of the membrane. Try to avoid transferring
excess medium with the lung, and make sure the lung is
properly oriented (see Note 9 for suggested technique to
orient the lung). See Fig. 2a for an overview of the experimental setup used to culture lung explants.
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Fig. 2 Experimental setup to culture whole embryonic lung explants. (a) Dissected whole lungs are properly
positioned on filter membranes placed on wire meshes designed for suitable culture dishes. (b) Example of a
cultured E12.5 lung. Note that the wire mesh is not visible. Scale bar, 1000 μm
(d) Incubate plate at 37 C and 5% CO2, or, if samples are
intended for live imaging, put it directly in the calibrated
live imaging chamber (see Subheading 3). Prior to imaging, lungs should be allowed to settle on the membrane
for approximately 30 min. See Fig. 2b for an example of a
cultured whole lung explant.
2. Isolated Epithelium and Mesenchyme Culture
In experiments involving the culture of isolated endoderm
and mesenchyme, it is common for investigators to use
serum-free, chemically defined media supplemented with predetermined amounts of compounds, such as growth factors.
Therefore, the following steps assume additional modifications
to the culture medium will be required to meet one’s particular
research question.
(a) Make a working solution of Matrigel® by diluting it 1:1
with DMEM/F-12 serum-free culture medium, supplemented with the desired compounds. Keep on ice.
(b) Prepare a suitably sized culture dish by adding a thin layer
of this diluted Matrigel® to the bottom of each samplecontaining well. Allow to harden for 1 min at room
temperature.
(c) Add enough diluted Matrigel® to each sample-containing
well to form a dome.
(d) Using a calibrated micropipet, quickly transfer samples to
the middle of the dome of Matrigel®.
(e) Place culture dish with properly positioned samples in an
incubator at 37 C and 5% CO2 for approximately
20–30 mins or until the gel polymerizes.
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Fig. 3 Activity of FGF7 and FGF10 in cultures of isolated E11.5 lung epithelium. (a) Epithelium grown for 48 h in
control medium, without supplemental FGFs. Note how most cells have died. (b) Culture with 250 ng/mL FGF7
supports the growth of epithelium and its expansion into a cyst-like structure absent any obvious buds. (c)
Culture with 250 ng/mL FGF10 results in epithelium that undergoes significant primary and secondary
branching. Scale bar, 250 μm. (Modified, with permission of Company of Biologists, from Bellusci et al. [14])
(f) Gently cover the polymerized Matrigel® with the same
medium used to dilute the gel. Place the dish back in the
incubator for the duration of the experiment, or, if live
imaging, put it directly in the calibrated live imaging
chamber. See Fig. 3 for an example of isolated epithelium
cultured in Matrigel® and media supplemented with various fibroblast growth factors (FGFs).
3.3 Still and Live
Imaging
After the lung samples have been properly obtained, prepared, and
positioned on filter membranes or in Matrigel® in suitable culture
plates provided with medium, one can obtain valuable phenotypic
data via imaging of the samples. Images should be taken at the start
of the experiment and at the end of the experiment, at a minimum.
All samples should be imaged at the same magnifications using
comparable brightness and contrast settings. The following section
provides a general overview of imaging samples using bright-field
microscopy to obtain still pictures and by using fluorescent microscopy during live imaging.
For all imaging steps, which involve the transport and manipulation of samples, wear gloves and a lab coat, and ensure that the
working area is clean to avoid sources of contamination. Furthermore, ensure the microscope used is properly adjusted to take
optimal images (simple adjustments, such as Koehler illumination,
may be performed by the researcher; a specialized technician should
be consulted for more advanced adjustments).
1. Still Images Using Bright-Field Microscopy
(a) Remove culture plate from incubator, and place it under
dissecting stereoscope.
Embryonic Whole Lung and Tissue Culture: Imaging and Analysis
119
(b) Remove lid of culture dish and keep it top-down on a
sterile surface.
(c) Set the microscope to the bright-field position, and turn
on the light source, adjusting the brightness so that it is
optimal while looking through the oculars. Turn on the
camera and the imaging software.
(d) Focus on a portion of the well so that only culture
medium is present in the field of view, along with the
overlying filter or Matrigel®. In the software, click on
“white balance”. This will adjust any background color
(such as phenol red) to white in the images.
(e) Find the sample. Adjust its position by rotating the plate,
and focus on it at the desired magnification (see Note 10).
Often, the focus while looking through the oculars differs
from that seen on the computer screen. Keep in mind the
picture captured will have the same focus as on the computer. Adjust contrast and brightness by using the software to achieve optimal images (see Note 11).
(f) Repeat steps 4 and 5 for each sample, keeping in mind to
use the same magnifications for each.
(g) Save images in a format compatible with post-processing
software, avoiding formats that reduce image quality (e.g.,
JPEG) (see Note 12). Figure 4 shows an example of still
images taken at different magnifications.
Fig. 4 Still images of a wild-type E12.5 whole lung explant cultured for 24 h. (a–c) The lung at time 0 h at three
different magnifications. (d–f) The same lung taken 24 h later at same magnifications. Scale bars, 1000 μm
(a, d); 500 μm (b, e); 250 μm (c, f)
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2. Live Imaging Using Fluorescent Microscopy
Before proceeding with live imaging, make sure to have the
live imaging equipment turned on and the climate control
chamber calibrated to 37 C and 5% CO2 well beforehand
(e.g., the night before imaging begins). Also, ensure there is
adequate water in the designated container to guarantee proper
humidity throughout the live imaging.
(a) Turn on all necessary equipment including the microscope and the designated computer, and open the live
imaging software. Make sure to properly set up the microscope to obtain the desired fluorescent channels.
(b) Fit the microscope with the proper stage to hold the
culture dish, and place samples cultured in phenol
red-free medium into the live imaging chamber. Keep
the lid of the culture dish on.
(c) Start the imaging software, and select the fluorescent
channels to be imaged, including the bright-field channel
if desired.
(d) Working in bright-field, find the sample to be imaged
through the oculars, and then switch the view to the
camera to see the image on the computer monitor.
(e) Focus on the sample at desired magnification, and then
switch to each applicable fluorescent channel, and use the
gauges in the software to make adjustments to the exposure time, intensity, and gain in each (see Note 13). If
quantifying the fluorescence intensity of multiple samples,
it is essential to apply the same settings to each sample, so
that valid comparisons can be made.
(f) Input the duration of the experiment and the time
between capturing an image (e.g., 30 min). If imaging
multiple samples, it is necessary to find and focus on each
sample and then store the coordinates of each sample in
the software. In this way, during each image capture, the
software will automatically move the microscope stage to
the stored coordinates and take an image.
(g) Start the live imaging experiment. Make sure microscope
room is kept dark for the duration of the experiment.
(h) Once the live imaging has been complete, save images in a
format compatible with post-processing software, avoiding formats which reduce image quality (e.g., JPEG). Not
only can each image be analyzed individually, but also the
series of images for each sample can be made into a timelapse movie. Figure 5 shows a series of images taken of an
E12.5 lung expressing red fluorescent protein (RFP) in
the epithelium.
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121
Fig. 5 Epithelial expression of red fluorescent protein (RFP) in a mouse E12.5 whole lung explant. Lungs were
cultured on a nucleopore membrane in phenol red-free culture medium and live imaged for 24 h. (a) Still
image at time 0 h showing the orientation of the lung, with the bright-field and red fluorescent channels
overlayed. (b–h) Still images taken at 4-h intervals showing the same lung, but only with the red channel
active. Note the extent of branching that occurs (see white arrows for examples). Scale bar, 250 μm
3.4 Lineage
Labelling and Tracing
Using a CreERT2/LoxPSTOP-LoxP Reporter
System
A powerful research model often employed to label and trace cells
involves creating genetically modified mice that express an inducible CreERT2 recombinase under the transcriptional control of a
gene of interest (such as a gene expressed by a specific cell type), as
well as a reporter construct downstream of a STOP sequence
flanked by two LoxP sites (a LoxP-STOP-LoxP reporter construct). CreERT2 is activated upon administration of tamoxifen
and functions to recombine the DNA flanked by target sequences
called LoxP sequences. In LoxP-STOP-LoxP reporter constructs,
recombination removes the STOP sequence, leading to reporter
expression. There are a variety of reporters, including green, red,
and yellow fluorescent proteins. The CreERT2/LoxP reporter
system is therefore a robust model to label specific cells at precise
times (for reviews, see refs. [10, 11]).
In this section we draw on the previous three sections to outline
an approach to conducting in vitro experiments using a CreERT2/
LoxP-STOP-LoxP-TomatoRFP mouse model to label and trace
cells. We supplement the general methods discussed with reference
to work conducted in our lab and recently published [12].
1. Crossing Mice and Induction at Desired Time Point:
Set up mating between an inducible CreERT2 driver mouse
line and a LoxP-based reporter line to obtain timed-pregnant
females (E0.5 at noon on the day a vaginal plug is seen). In our
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example, we crossed Fgf10CreERT2/+ mice with TomatoRFP flox/flox
mice to obtain Fgf10CreERT2/+/TomatoRFP flox/+ experimental
embryos (hereafter called Fgf10RFP).
(a) Inject intraperitoneally an optimal volume of tamoxifen working solution approximately 1 or 2 days before
harvesting the embryos (see Note 14). In our example
experiments, we administered a single injection of
0.075 mg of tamoxifen per gram body weight 2 days
before harvesting.
2. Harvesting, Culturing, and Manipulating Samples: See Subheadings 3.1 and 3.2 above for harvesting and culturing protocols (see Note 15).
(a) Add supplementary molecules at optimal concentrations
directly to culture medium. Mix by pipetting. It is often
necessary to conduct optimization experiments to determine the concentration required to produce a phenotypic
change. In our example, vivo-morpholinos, used to inhibit
the activity of a target microRNA (miRNA), were added to
the culture medium at 1–4 μM.
3. Imaging and Analysis: Pictures are taken as described in Subheading 3.3 (Still and Live Imaging) above.
(a) To analyze still images, save images in a format compatible
with analyzing software, avoiding formats that reduce
image quality (e.g., JPEG). Conduct morphometric analyses using software such as ImageJ. Various morphometric parameters of whole lung explants can be calculated,
including mesenchymal and epithelial area, length of
branches, and branching complexity. For cultured cells,
parameters such as cell number, average cell size, and cell
distribution can be quantified (see Note 16). Figure 6
shows an example of a morphometric analysis applied to
whole lung explant cultures.
(b) Analyze live imaging data using software such as MetaMorph®. Export images in a compatible format. MetaMorph® can calculate a number of parameters, including
cell morphometry and number, fluorescence intensity, and
motion analysis (see Note 16). Figure 7 shows an example
of Fgf10RFP whole lung explants live imaged for 48 h,
along with an analysis of cell motility and cell proliferation
(see figure for a description of the procedure to conduct the analysis).
Embryonic Whole Lung and Tissue Culture: Imaging and Analysis
123
Fig. 6 Morphometric analysis of E11.5 wild-type lungs cultured for 48 h. (a–d) Lungs cultured with vivomorpholino targeting a scrambled sequence (scra) (e–h) or mir-142-3p (mo142). After 48 h, (i) number of
buds, branching points and branches, (j) length of each branch order, and (k) epithelial and mesenchymal area
were quantified. Scale bars, 250 μm (a, c, e, g); 50 μm (b, d, f, h). Data are means s.d. (Modified, with
permission of Company of Biologists, from Carraro et al. [12])
4
Notes
1. Some experiments might require serum-free, chemically
defined culture medium. Other experiments, especially those
involving fluorescent imaging, require medium without phenol
red (e.g., DMEM/F-12 medium without phenol red). The
general-purpose medium we use most frequently in our lab,
though chemically incompletely defined, is optimal for the
growth and maintenance of tissue explants and cells over a
period of a few days.
2. If samples are intended for fluorescent imaging, make sure to
use culture medium free of phenol red, as this causes extreme
autofluorescence.
3. Cut the wire mesh to dimensions that will fit into the wells of
the culture dish used in the experiment. For example, for a
24-well dish with wells of 15 mm in diameter, cut circular
pieces of the metal mesh with a diameter of approximately
13–14 mm. When cutting the mesh, make sure to leave four
tabs sticking from the circular piece (one large tab approximately 5–6 mm in length and three smaller ones, positioned
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Matthew Jones and Saverio Bellusci
Fig. 7 Cell motility and proliferation analyses of Fgf10RFP whole lung explants and isolated mesenchymal cells.
(a, b) E11.5 lungs cultured with scrambled, and (c, d) mo142 vivo-morpholinos. (h–k) Isolated mesenchymal
cells electroporated with plasmids containing GFP (control) or APC (experimental) and cultured for 45 h. (e–n)
Using MetaMorph® software, cell motility and proliferation were analyzed. LIF files were imported to the
software, and random fluorescent cells were marked at 0 h using the “track object” tool. These cells were
then automatically tracked over the duration of the live imaging experiment. It was then possible to determine
how much each tracked cell moved (e, f and l, m) and, as a measure of proliferation, how many divisions each
cell underwent (g and n). Scale bars, 250 μm (a, c); 50 μm (b, d); 75 μm (h–k). Data are means s.d.
(Modified, with permission of Company of Biologists, from Carraro et al. [12]).
equidistant around the circumference of the circle, approximately 2–3 mm in length). Using a suitable tool, such as a
pair of pliers, bend the large tab 90 upward, and bend the
smaller tabs 90 in the opposite direction. The large tab now
serves as a “handle” to easily insert, position, and remove the
metallic grid from the well; the three smaller tabs serve as
“feet” to raise the mesh off the bottom of the well. Finally,
use a tool such as a hole punch to create a hole in the middle of
the grid. Make the hole large enough so that the grid will not
be visible as the magnification pictures are taken. Store
Embryonic Whole Lung and Tissue Culture: Imaging and Analysis
125
completed metallic grids in 100% ethanol. See Fig. 2a for an
example of a metallic grid used for culturing.
4. A common genetic background for experimental and wild-type
mice is C57Bl/6J. However, a variety of other backgrounds
exist, and might be more useful for one’s experiment.
CD1-IGS mice, for example, tend to produce large litters.
Thus, if a large number of embryos are required, this background might be a better choice.
5. There is no need to immobilize embryos by pinning them. It is
easiest to work on early-stage embryos while they are
submerged in medium. However, some researchers might
find that pinning later-stage embryos (i.e., E18.5) makes
them easier to dissect. We describe the dissection of lungs
using unpinned samples.
6. In some experiments it might be necessary to have a completely
intact trachea. In those cases, dissecting the larynx along with
the trachea is advisable. This can be accomplished by enlarging
the semicircular incision to include the pharynx and dissecting
through the region above the larynx.
7. Steps 3 and 4 carry a high risk of damaging the lung, especially
in early-stage embryos. Utmost care, hand steadiness, and
adequate practice can help ensure lungs remain undamaged.
8. The metallic grid ensures the membrane and the sample do not
shift too much during the manipulation and movement of the
plate necessary to take pictures. The central hole allows one to
take clear pictures. The grid is particularly important for live
imaging experiments, as manual manipulation of the plate is
not possible before pictures are taken.
9. It is essential all lungs have similar orientation on the membrane so that comparable pictures can be taken. Positioning the
lungs ventral side up, for example, with the left lobe on the
right or bottom edge of the field of view, is an ideal orientation
to image branching epithelium in E11.5-E14-5 lungs. Furthermore, the trachea of each sample should be straight, thus
ensuring each lung has similar internal partial pressure.
Proper orientation can be achieved by the following technique: take a lung from the medium so that the lung occupies
only the end of the pipet; ensure the trachea enters the pipet
first and the orientation of the lung is known (e.g., dorsal side
facing down). Transfer the lung to the filter by gently placing
the tip of the micropipet at a 45 angle on the filter. Slowly
expel the lung as the tip of the pipet is drawn across the middle
of the filter. In this way, the posterior of the lung will stick to
the filter, and as the pipet is drawn away, the lung will be
properly positioned. It may be necessary to use sterile forceps
to make minor adjustments to the lung after it has been placed
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Matthew Jones and Saverio Bellusci
on the filter; however, this carries the risk of damaging
the lung.
10. Each image should have the same orientation for comparison
purposes. It is advisable to manipulate the culture dish to
achieve this, and not to insert anything into the wells to rotate
the filters directly.
11. A common problem during imaging is that details of the
sample are masked by overly bright backgrounds or extreme
contrast. This can be avoided by adjusting the brightness and
gain gauges directly on the software.
12. File formats such as JPEG can drastically reduce image quality
due to compression processes. When possible, save files in the
native format provided by the company of the microscope and
software (e.g., Leica “.LIF” files).
13. A common problem with taking fluorescent images is oversaturation, which occurs when the intensity of the signal is above
the maximum recognized by the camera. A technique to avoid
this problem is to toggle the displayed colors in the software to
show minimum and maximum saturations (the toggle is called
“QuickLUT” in Leica software). Adjust the gauges to find the
saturation point of the signal. Slightly reducing the signal
below this point will provide maximum signal intensity without
oversaturating the image.
14. The optimal amount may differ between driver lines, tissue,
and cell of expression and needs to be determined either from
the literature or in the lab. Also, it takes time for tamoxifen to
achieve peak efficiency. In our experience, injecting a couple of
days before harvesting early-stage embryos provides maximum
recombination efficiency (e.g., inject at E10.5 to work with
E12.5 embryos).
15. In the example paper, aside from whole lung explant cultures,
we also cultured primary mesenchymal cells. This method was
not included in Subheading 2. Mesenchymal cells were
obtained by a differential adhesion protocol (for details, see
[13]). Briefly, whole lungs were dissected and subjected to
trypsin digestion. A single-cell suspension was produced and
passed through mesh filters. The remaining cells were plated,
and mesenchymal cells obtained by differential adhesion to the
plate bottom.
16. The particular steps involved in analyzing the images are
beyond the scope of this chapter. Consult the user manual of
the software for details on how to conduct morphometric
calculations.
Embryonic Whole Lung and Tissue Culture: Imaging and Analysis
127
Acknowledgments
We wish to thank Salma Dilai for her contribution to the manuscript. S.B. was supported by grants from the Deutsche Forschungsgemeinschaft (DFG; BE4443/1-1, BE4443/4-1,
BE4443/6-1, KFO309 P7, and SFB1213-projects A02 and
A04), Landes-Offensive zur Entwicklung Wissenschaftlich-Ökonomischer Exzellenz (LOEWE), UKGM, Universities of Giessen and
Marburg Lung Center (UGMLC), DZL, and COST (BM1201).
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11. Abe T, Fujimori T (2013) Reporter mouse
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12. Carraro G, Shrestha A, Rostkovius J,
Contreras A, Chao CM et al (2014)
miR-142-3p balances proliferation and differentiation of mesenchymal cells during lung
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13. Lebeche D, Malpel S, Cardoso WV (1999)
Fibroblast growth factor interactions in the
developing lung. Mech Dev 86:125–136
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Chapter 9
Mouse Hematopoietic Stem Cell Modification and Labelling
by Transduction and Tracking Posttransplantation
Benjamin Cao, Songhui Li, Claire Pritchard, Brenda Williams,
and Susan K. Nilsson
Abstract
The tracking of the hematopoietic potential of genetically manipulated fluorescent hematopoietic stem cells
(HSC) in the bone marrow (BM) allows the assessment of regulatory processes involved in the
re-establishment of hematopoiesis posttransplant. Herein, we describe the means to assess the consequence
of expressing specific genes in HSC on their engraftment potential posttransplant.
Key words Hematopoietic stem cells, Lentiviral transduction, Bone marrow transplantation
1
Introduction
As bone marrow (BM) hematopoietic stem cells (HSC) are ultimately responsible for the production of all blood cells, they are
routinely used in the clinic to reconstitute hematopoiesis following
transplantation. The process of reconstitution requires multiple
steps including homing and lodgment in a specialized microenvironment within the BM termed the stem cell “niche” [1–4], followed by proliferation and differentiation to produce the required
circulating blood cells. A large body of data has now been accumulated identifying key molecules involved in various aspects of this
process (reviewed in [5–7]). However, less than 30% of transplanted HSC actually home to the BM, with the majority of cells
being sequestered elsewhere in the body, in organs such as the liver
[8], resulting in highly variable engraftment levels. An ability to
improve BM engraftment posttransplant would have significant
clinical implications.
Herein we describe a method to assess the effects of the expression of specific genes in HSC on their engraftment potential posttransplant. We demonstrate that lentiviral transduction of purified
murine HSC with a gene of interest as well as a fluorescent protein
Ivan Bertoncello (ed.), Mouse Cell Culture: Methods and Protocols, Methods in Molecular Biology, vol. 1940,
https://doi.org/10.1007/978-1-4939-9086-3_9, © Springer Science+Business Media, LLC, part of Springer Nature 2019
129
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provides an easy method of tracking long-term multi-lineage
engraftment posttransplant using flow cytometry and hence provides an efficient and effective way of assessing the role of exogenously added genes.
2
Materials
2.1 Isolation of Bone
Marrow
and Preparation of HSC
1. Adult C57Bl/6J (Ly5.2) mice 6–8 weeks old (see Note 1).
2. Sterile #11 surgical blade and #3 handle.
3. Phosphate buffered saline (PBS): pH 7.2, 310 mOsm/L (see
Note 2) supplemented with 2% serum (PBS 2% Se): Defined
bovine calf serum, iron supplemented.
4. Sterile 50 mL conical tubes for the collection of BM.
5. Sterile porcelain mortar and pestle for grinding bones.
6. Sterile 40 μm nylon cell strainers.
7. 4 mg/mL solution of dispase II and 3 mg/mL solution of
collagenase I in phosphate buffered saline (PBS): pH 7.2,
310 mOsm/L.
8. 37 C orbital shaker, for example, Eppendorf Thermomixer C
(Eppendorf, Hamburg, Germany).
9. Hemocytometer and microscope equipped with phase contrast
or an automated cell counter. We use a CELL-DYN Emerald
18 (Abbott).
10. NycoPrep™ 1.077 Animal (Axis-Shield, Oslo, Norway).
11. Cannulas, for example, Unomedical (Unomedical, Mona Vale,
NSW, Australia) attached to 20 mL syringes.
12. Lineage depletion antibody cocktail: Purified rat anti-mouse
antibodies recognizing the cell surface antigens: B220 (lymphoid), GR-1 and MAC-1 (myeloid), and TER119 (erythroid); we used antibodies from BD Pharmingen, San
Diego, USA (see Note 3).
13. PBS supplemented with 2 mM EDTA and 0.5% (w/v) fraction
V bovine serum albumin (BSA, Sigma-Aldrich, St. Louis, MO,
USA), pH 7.4 (PBS EDTA 0.5% BSA).
14. Dynabeads for magnetic labelling of the cells: Sheep anti-rat
IgG beads 4.5 μm diameter, 4 108 beads/mL (Dynal Biotech ASA, Oslo, Norway).
15. MPC-L magnet for a 1 mL to 8 mL sample (Dynal Biotech).
16. Suspension mixer: Allowing both tilting and rotation at 4–8 C
for Dynabead incubation step (we use a MACSmix Tube Rotator in a fridge).
17. Sterile 14 mL polypropylene round-bottom tubes.
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18. 5 mL polystyrene round-bottom tube with cell strainer cap.
19. A cocktail of antibodies for HSC isolation: Rat-anti-mouseSca-1-PECy7 (Ly-6A/E, clone E13-161.7), rat-anti-mouse-ckit-BUV395 (CD117, clone 2B8), rat-anti-mouse-CD48FITC (CD48, clone HM48), and rat-anti-mouse-CD150-PE
(CD150, clone TC15-12F12.2)-conjugated antibodies. Antibody concentrations are pre-titred and all 1 μg/mL (we use
BD Pharmingen or BioLegend; see Note 4).
20. A solution of 0.05 μg/mL propidium iodide (PI) in PBS 2% Se
for determining cell viability.
2.2 HSC
FluorescenceActivated Cell Sorting
1. Flow cytometer with sorting capability (we use a BD Influx cell
sorter equipped with five solid-state lasers (355, 405, 488, 561,
and 628 nm). Band-pass filter settings for the detection of
fluorescence for FITC, PECy7, BUV395, PE, and PI are
528 19, 750 60, 362 34, 575 15, and 655 20,
respectively. HSC are sorted using a 70 μm nozzle at 30 psi,
drop delay frequency of 61 kHz. Sorting speed 25,000 cells/s.
2. Sterile 5 mL polypropylene tubes for cell collection postsorting.
3. Hemocytometer.
4. Light microscope fitted with phase contrast.
2.3
Titration of Virus
1. Lentivirus containing the RFP reporter and gene of interest
generated using a protocol such as that described in a preceding
chapter in this volume [9].
2. FDC-P1 cell line (ATTC CRL-12103).
3. WEHI-3 conditioned medium (CM) as a source of interleukin3 for FDC-P1 cell maintenance.
4. DMEM media supplemented with 5% Se and 25% WEHI-3
CM to maintain cells.
5. DMEM media supplemented with 5% Se, 2 mM of GlutaMAX,
25% WEHI-3 CM, and 4 μg/mL of Polybrene.
6. Sterile 1.5 mL microtubes.
7. 24-well cell culture plates.
8. 12-well cell culture plates.
9. A solution of 0.125 μg/mL 40 ,6-diamidino-2-phenylindole
(DAPI) in PBS 2% Se for determining cell viability.
10. Flow cytometric analyzer (we use a BD LSRII analyzer
equipped with five solid-state lasers (355, 405, 488, 561, and
628 nm). Band-pass filter settings for the detection of fluorescence for DAPI and RFP are 450 25 nm and 575 19,
respectively.
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2.4 Transduction
of Murine HSC
1. Lentivirus containing the RFP reporter and gene of interest.
2. Non-treated 48-well cell culture plates.
3. 96-well cell culture plates.
4. Recombinant human fibronectin fragment RetroNectin
(Takara).
5. PBS 310 mOsm/L 2% BSA.
6. PBS 310 mOsm/L 0.5% BSA.
7. HSC tissue culture media (we use StemSpan SFEM II media,
Stemcell Technologies).
8. 10% CO2 tissue culture incubator.
9. L-Glutamine (we use 200 mM GlutaMAX, Thermo Fisher
Scientific).
10. Recombinant mouse stem cell factor (rmSCF) and recombinant human Flt3 ligand (rhFlt3L).
11. Low-oxygen triple-mix tissue culture incubator (5% O2, 10%
CO2 in N2).
12. Hemocytometer and light microscope fitted with phase
contrast.
13. A solution of 0.125 μg/mL DAPI in PBS 2% Se for determining cell viability.
14. Flow cytometric analyzer (we use a BD LSRII analyzer
equipped with five solid-state lasers (355, 405, 488, 561, and
628 nm). Band-pass filter settings for the detection of fluorescence for DAPI and RFP are 450 25 nm and 575 19,
respectively.
2.5 HSC
Transplantation
1. 200,000 total bone marrow cells per recipient irradiated with
15 Gy as fillers.
2. 1 mL syringe with a 27 gauge needle.
3. Heat lamp.
4. 75% ethanol made in distilled water.
5. Kleenex tissue.
6. Apparatus to immobilize mouse during injection.
2.6 HSC Transplant
Analysis
1. Heparinized capillary tubes.
2. K2-EDTA microtainer tubes for blood collection.
3. 14 mL polypropylene round-bottom tubes.
4. 5 mL polystyrene round-bottom tubes with cell strainer caps.
5. Hemocytometer and microscope equipped with phase contrast
or an automated cell counter. We use a CELL-DYN Emerald
18 (Abbott).
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133
6. Antibodies: Rat-anti-mouse-CD3e-BV510 (clone 17A2),
rat-anti-mouse-CD45R-BV510 (B220, clone RA3-6B2),
rat-anti-mouse-CD45R-AF647, rat-anti-mouse-Gr-1-AF647
(Ly6G, clone RB6-8C5), and rat-anti-mouse-Mac-1-AF647
(CD11b, clone M1/70) are all BD Pharmingen or BioLegend.
Antibodies are optimally pre-titred and used at less than 1 μg/
mL.
7. NH4Cl lysis buffer composed of 150 mM NH4Cl, 10 mM
NaHCO3, 1 mM disodium EDTA, and distilled water, pH 7.4.
8. Flow cytometric analyzer (we use a BD LSRII analyzer
equipped with five solid-state lasers (355, 405, 488, 561, and
628 nm). Band-pass filter settings for the detection of fluorescence for RFP, BV510, AF647, and DAPI are 575 19,
500 50, 650 20, and 450 25 nm, respectively.
3
Methods
3.1 Sampling Murine
Bone Marrow
1. Kill mice by cervical dislocation and dissect iliac crests, femurs,
and tibias, and remove the muscle and connective tissue.
2. Store bones in 10 mL PBS 2% Se in a 50 mL Falcon tube.
3. Decant bones into a sterile mortar.
4. Grind bones with the pestle until the marrow cavity is open to
expose it to enzymatic digestion. Be careful not to pulverize the
bones.
5. Thoroughly mix cell and crushed bone suspension by pipetting
the supernatant up and down, then remove the cell supernatant, and filter through a 40 μm nylon cell strainer into a 50 mL
conical tube.
6. Rinse with PBS 2% Se and further crush bone fragments until
they become white.
7. Collect and filter the supernatant as indicated in step 5. Top up
the tube to 50 mL with PBS 2% Se and set aside on ice until
step 12.
8. Transfer the crushed bone fragments into a new 50 mL conical
tube containing the collagenase I/dispase II enzymatic suspension (1 mL per crushed bones of 1 mouse), and shake at 37 C
in an orbital shaker, 750 rpm for 5 min.
9. Add 20 mL neat PBS to the digested bone fragments and shake
vigorously for 20 s.
10. Filter the cell suspension through a 40 μm nylon cell strainer
into another 50 mL conical tube.
11. Repeat steps 9 and 10 and filter cells into the same 50 mL
conical tube. Top up the tube to 50 mL with PBS 2% Se.
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12. Centrifuge the cell suspension tubes (from step 7–11) at
400 g for 5 min at 4 C.
13. Discard supernatant and resuspend cell pellet in 10 mL PBS 2%
Se. Perform a cell count, and store cells on ice for HSC
pre-enrichment by density depletion and immunomagnetic
separation.
3.2 HSC
Pre-enrichment
1. Dilute the cell suspension to approximately 2 108 cells/
20 mL with PBS 2% Se.
3.2.1 Density Gradient
Separation
2. Divide 20 mL aliquots of cell suspension over an even number
of 50 mL centrifuge tubes.
3. Underlay each gradient with 10 mL NycoPrep 1.077A using a
cannula attached to a 20 mL syringe.
4. Centrifuge the gradients at 600 g for 20 min at room
temperature with no de-acceleration.
5. Collect the mononuclear cells from the interface between the
PBS layer and the NycoPrep solution into a 50 mL centrifuge
tube using a cannula attached to a 10 mL syringe. Collect the
mononuclear cells of two gradients into a 50 mL falcon tube,
and fill the tube with PBS 2% Se.
6. Centrifuge the tubes at 400 g for 5 min at 4 C.
7. Decant the supernatant, and resuspend the pooled cell pellets
in 50 mL PBS 2% Se.
8. Perform a cell count.
3.2.2 Immuno-labelling
Cells with a Cocktail
of Lineage Antibodies
1. Centrifuge cells at 400 g for 5 min at 4 C.
2. Stain cells at 1 107 cells/mL in the cocktail of antibodies
directed against hematopoietic lineage markers on ice for
20 min (see Note 5).
3. Wash cells with PBS 2% Se by centrifuging at 400 g for 5 min
at 4 C to remove unbound antibodies.
3.2.3 Immunomagnetic
Cell Separation
1. Resuspend cells in 2 mL PBS 2 mM EDTA 0.5% BSA, and
transfer into a 5 mL sterile polypropylene tube. Perform a cell
count and set aside on ice until step 7.
2. Resuspend Dynabeads.
3. Calculate the volume of Dynabeads needed based on the cell
number. We use a Dynabead/cell ratio of 1:3 repeated in two
steps (see Note 6).
4. Dispense the volume of Dynabeads required for both steps into
individual 1.5 mL microtubes.
5. Wash away the azide in the Dynabeads by adding 1 mL PBS
2 mM EDTA 0.5% BSA to each microtube and mixing well.
HSC Tracking Post Transduction
135
Place the tubes in the magnet for 1 min, remove and discard
supernatant, and then remove microtubes from the magnet.
6. Repeat step 5.
7. Resuspend each aliquot of washed Dynabeads in 250 μL of PBS
2 mM EDTA 0.5% BSA.
8. Add the first aliquot of Dynabeads to the cells and mix well.
9. Incubate for 5 min at 4 C with gentle tilting and rotation.
10. Place the mixture in the magnet for 2 min.
11. Without removing the tube from the magnet, transfer the
supernatant containing the unbound cells to a new 5 mL
polypropylene tube.
12. In order to collect any residual unbound cells, rinse the beadbound cells with 1 mL PBS 2 mM EDTA 0.5% BSA, and place
in magnet for 1 min.
13. Transfer the supernatant to the tube from step 11.
14. Add the second aliquot of Dynabeads to the collected cells and
mix well.
15. Incubate for 10 min at 4 C with gentle tilting and rotation.
16. Repeat steps 10–12.
17. Transfer the supernatant to a new 14 mL polypropylene tube.
18. Make up the volume of the negative cell suspension (unbound
cells) to 10 mL with PBS 2% Se, and perform a cell count.
3.3
HSC Isolation
3.3.1 HSC Labelling
1. Pellet cells by centrifuging at 400 g for 5 min at 4 C.
2. Stain cells at 1 108 cells/mL in an optimally pre-titred HSC
antibody cocktail, and incubate light protected on ice for
20 min.
3. Wash cells in 3 mL PBS 2% Se by centrifuging at 400 g for
5 min at 4 C to remove unbound antibody. Discard
supernatant.
4. Resuspend cells at 30–40 106 cells/mL in solution of PI, and
filter the cell suspension through a cell strainer into a new 5 mL
polypropylene tube prior to fluorescence-activated cell sorting
(see Note 7). Place on ice until sorted.
3.3.2 HSC Sorting
1. To set up the HSC sort, the following samples are required.
(a) 0.5–1 106 unstained total marrow cells to set the voltages for forward scatter, side scatter, FITC, BUV395,
PECy7, and PE.
(b) Individual tubes containing 0.5–1 106 cells stained with
appropriate FITC, PECy7, BUV395, and PE antibody
conjugate for compensation controls (see Note 8).
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Fig. 1 Representative flow cytometric plot of Lineage-Sca1+ckit+CD150+CD48+
(LSKSLAM) HSC for FACS sorting
2. Run the cell samples stained with HSC antibodies, and sequentially gate through FSC-H versus FSC-A, SSC-A versus FSC-A,
FSC-H versus PI, sca-PeCy7 versus c-kit-BUV395, and CD48FITC versus CD150-PE (Fig. 1).
3. Sort cells at predetermined optimal input speed, and collect
into 5 mL polypropylene tubes containing 200 μL PBS 2% Se.
4. Perform reanalysis of a 10 μL aliquot of sorted cells.
5. Perform a manual
hemocytometer.
3.4 HSC
Transduction
3.4.1 Virus Titration
viability
cell
count
using
the
1. Seed 4 wells of a 12-well plate with 2 105 FDC-P1 cells in
0.5 mL of DMEM 5% Se, 2 mM of GlutaMAX, 25% WEHI
CM, and 4 μg/mL of Polybrene.
2. Thaw virus stock from 80 C.
3. Make 1:10 and a 1:100 dilutions of the virus in the media from
step 1.
4. Add 10 μL of the undiluted and diluted virus to three wells of
FDC-P1 cells. The fourth well of cells is used as control.
Incubate the cells at 37 C overnight in a standard tissue
culture incubator (10% CO2).
5. Collect 50 μL of cells from each well into a 1.5 mL microtube.
6. Wash three times with 1 mL of PBS 2% Se by centrifuging at
4 C, 400 g for 5 min, and removing the supernatant.
7. Resuspend cells in 2 mL of media from 1 without Polybrene,
and culture in a 12-well plate for 3 days.
8. Collect cells from the plate, and wash once with 3 mL of PBS
2% Se by centrifuging at 4 C, 400 g for 5 min, and removing
the supernatant.
9. Resuspend cells in 200 μL of PI solution except for the control
cells which are resuspended in PBS 2% Se.
HSC Tracking Post Transduction
137
10. Analyze the cells by flow cytometry, and determine the percentage of RFP+ cells.
11. Calculate the virus titer using the dilution closest to 10% RFP+
cells using the following formula:
Virus titer ¼ 2 105 (percentage of RFP+ cells/100)/(10 dilution factor/1000) transduction units (TU)/mL.
3.4.2 HSC Transduction
1. Coat the required number of wells of a non-treated 48-well
culture plate with 200 μL of 50 μg/mL RetroNectin in PBS,
and incubate at 4 C overnight or at room temperature for 2 h.
2. Remove RetroNectin from wells, and block each well with
200 μL of PBS 2% BSA at room temperature for 30 min.
3. Remove BSA and wash each well with 300 μL of PBS.
4. Add 200 μL of SFEM II to the first RetroNectin-coated well,
then the estimated amount of virus required to give a multiplicity of infection (MOI) equal to or greater than 50 to each
subsequent well plus the necessary volume of SFEM II to make
the total volume equal 200 μL (see Note 9). Incubate at 37 C,
10% CO2 until HSC (LSKSLAM) are sorted.
5. Place 3000 HSC for the control and up to 500,000 HSC into
separate 5 mL polypropylene tubes, centrifuge at 400 g 4 C
for 5 min, and dry pellet.
6. Resuspend the 3000 control HSC in the media from the first
well, and place them back into the same well.
7. Resuspend up to 500,000 HSC with media containing virus
from the second well, and place them back into the same well.
8. Repeat for any remaining wells.
9. Add 2 μL of GlutaMAX to each well to have a final concentration of 2 mM and 10 ng/mL rmSCF and rhFlt3L. Mix well,
and incubate at 37 C in a triple-mix incubator (5% O2, 10%
CO2 in N2) overnight.
10. Collect transduced cells from each well, and wash twice with
1 mL of PBS 0.5% BSA by centrifuging at 400 g for 5 min at
4 C and removing the supernatant.
11. Resuspend cells with 1 mL of PBS 0.5% BSA, and perform a
viability cell count using phase contrast and a hemocytometer.
12. Transfer 3000 HSC from each sample of transduced cells to a
new tube for culture.
13. Add 200,000 filler cells per mouse to the residual transduced
HSC, and centrifuge at 400 g 4 C for 5 min prior to
resuspending each in PBS 0.5% BSA to allow for 200 μL per
transplant recipient.
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Benjamin Cao et al.
Fig. 2 Representative flow cytometric plot of FACS-purified LSKSLAM that has been mock-transduced
(no lentivirus) or transduced with lentivirus containing a RFP reporter. Cells were cultured in vitro for 3 days
to allow identification of transduced RFP+ cells
14. Add 500 μL SFEM II supplemented with 2 mM GlutaMAX to
each tube containing 3000 transduced cells for culture, and
centrifuge at 400 g 4 C for 5 min.
15. Dry pellet and resuspend cells in 200 μL of SFEM II supplemented with 2 mM GlutaMAX, 10 ng/mL rmSCF, and
10 ng/mL rhFlt3L, and place 100 μl per well in a 96-well
plate in a triple-mix incubator (5% O2, 10% CO2 in N2) for
72 h.
16. Collect cells, wash once with 3 mL of PBS 2% Se, and resuspend control cells in 100 μL of PBS 2% Se and transduced cells
in 0.125 μg/mL DAPI in PBS 2% Se for flow cytometric
analysis. Run the cell samples, and sequentially gate through
FSC-H versus FSC-A, SSC-A versus FSC-A, FSC-H versus
DAPI, and RFP versus SSC-A to determine the percentage of
RFP+ cells as a measure of transduction efficiency (Fig. 2).
3.5
HSC Transplant
1. Prepare recipients using ablation method approved by the
institution’s ethics’ committee. We use total body irradiation
given in a split dose of 5.5 Gy each, 5 h apart 24 h prior to
transplant using two opposing 137Cs sources Gammacell
(40, Atomic Energy of Canada) (see Note 10).
2. Place recipient animals under a heat lamp to dilate the tail vein.
HSC Tracking Post Transduction
139
3. Fill 1 mL syringe attached to a 27-gauge needle with wellmixed cell suspension.
4. Place recipient into mouse immobilization apparatus, and wipe
tail with 70% ethanol.
5. Inject 200 μL of cell suspension (from Subheading 3.4.2,
step 13) into recipient via the lateral tail vein (see Note 11).
6. Release mouse and house in appropriate box with chow and
water ad libitum.
3.6 Transplant
Analysis
1. After the desired time period (we analyze after 6, 12, and
20 weeks), collect 50 μL of peripheral blood using the ethically
approved method at your institution (e.g., by tail vein, retroorbital plexus of saphenous vein bleeding). Collect blood in
heparinized or EDTA-coated tubes, and perform a cell count.
Make sure blood is also collected from control mice.
2. Transfer blood individually into 14 mL tubes containing 5 mL
lysis buffer, keep cells for 5 min at room temperature, and
check color of buffer solution (see Note 12). If buffer is visually
hemoglobinized, add 5 mL PBS 2% Se, and centrifuge at
400 g for 5 min, 4 C. If red blood cells are still visible,
increase lysis time or repeat lysis step.
3. Decant supernatants and resuspend in 3 mL PBS 2% Se. Transfer the cell suspension into 5 mL polystyrene tubes, filtering
through the cell strainer cap.
4. Wash cells again and store on ice until antibody staining.
5. Prepare antibody cocktail including T-cell, B-cell, and myeloid
markers. Allow 50 μL per sample (see Note 13).
6. Centrifuge cell samples at 400 g for 5 min, 4 C, and decant
supernatant.
7. Resuspend samples in 50 μL of antibody cocktail or single
antibodies for compensation. Ensure cell pellets are completely
disrupted and cell suspensions well mixed.
8. Incubate cells for 20 min on ice and light protected.
9. Centrifuge cell samples at 400 g for 5 min, 4 C, decant
supernatant, and resuspend cells in PBS 2% Se at a final volume
recommended for the flow cytometer used (see Note 14).
10. Run samples on flow cytometer, and sequentially gate through
FSC-H versus FSC-A, SSC-A versus FSC-A, FSC-H versus
DAPI, RFP versus SSC-A, and B220/Gr1/Mac1 vs
CD3/B220 to determine % RFP+ and to assess multi-lineage
reconstitution (Fig. 3).
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Fig. 3 Six-week peripheral blood analysis of recipients transplanted with mock-transduced (no lentivirus) or
lentiviral-transduced HSC. Progeny arising from transduced HSC is identified as RFP+ cells, and multi-lineage
reconstitution is assessed by analyzing CD3+ (T-cells), B220+ (B-cells), or Gr1+Mac1+ (myeloid) cells
4
Notes
1. This method is for 10 donor animals. Volumes can be altered
for more or less donors. We use the same method for a range of
transgenic mice.
2. This osmolarity is appropriate for murine cells and results in
better cell recovery.
3. The use of this limited antibody cocktail results in the removal
of approximately 70% of marrow cells. To gain higher purity,
additional antibodies directed against T-cell markers like CD3,
CD4, CD5, and CD8 can also be added to the cocktail.
4. Other conjugates can be used.
5. Each antibody needs to be individually titred to determine the
optimal working concentration for lineage depletion. It is a
good practice to centrifuge the antibody cocktail briefly in a
microfuge before use; the supernatant is then used, eliminating
non-specific background staining by any protein aggregates
formed during storage of the antibodies.
6. The Dynabead/cell number ratio of 1:3 was established
in-house based on the lowest number of Dynabeads required
without significant loss of depletion efficiency.
HSC Tracking Post Transduction
141
7. In order to reduce the incidence of nozzle clogs during sorting,
sort as soon as possible after labelling, and filter the sample
immediately prior to the sort.
8. For compensation tubes, it is valid to use the antibody conjugate used for the analysis of the samples as single antibody
controls. If CD45 is used, it is generally expressed at high levels
and can result in overcompensation. However, in our experience using CD45 antibodies to compensate for this sample
setup is suitable.
9. MOI equals the number of virus particles (TU) divided by the
number of cells, so an MOI of 50 is 50 times the number of
viral particles compared to cells.
10. When using irradiation as the method of preparative ablation,
an irradiation dose response should initially be performed to
select the optimal dose that removes the majority of recipient
HSC. We do this using a high-proliferative potential colonyforming assay (HPP-CFC) [10].
11. To avoid back flushing of the cell suspension, wait 20 s before
removing the needle after injecting cells. Only blood should be
visible at the injection site after removing the needle.
12. It is important to minimize the lysis time to avoid loss of
nucleated cells. However, a high number of non-lysed red
blood cells can result in turbulences in the flow chamber and
a consequential loss of scatter signal and a reduced quality flow
profile when samples are analyzed using flow cytometry. If in
doubt, optimize lysis time in your system with test blood
before lysing the samples.
13. By using the same antibody conjugated to two different fluorochromes (in this case B220-BV510 and B220-AF647) in the
same tube, T-cells, B-cells, and myeloid cells can easily be
distinguished in one dot plot. Alternate fluorochromes can be
used, keeping the reporter color in mind.
14. Depending on the cell number per tube, we generally resuspend
blood samples in 120 μL of buffer, and run the samples on a BD
LSRII flow cytometer. This results in approximately 1 min
running time per tube with a BD LSRII on setting “high.”
We would not recommend a flow rate higher than 10,000
events per second to ensure an optimal flow cytometry profile.
References
1. Schofield R (1978) The relationship between
the spleen colony-forming cell and the haemopoietic stem cell. Blood Cells 4:7–25
2. Gong JK (1978) Endosteal marrow: a rich
source of hematopoietic stem cells. Science
199:1443–1445
3. Mason KD, Carpinelli MR, Fletcher JI, Collinge JE, Hilton AA et al (2007) Programmed
anuclear cell death delimits platelet life span.
Cell 128:1173–1186
4. Nilsson SK, Johnston HM, Coverdale JA
(2001) Spatial localization of transplanted
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hemopoietic stem cells: inferences for the localization of stem cell niches. Blood
97:2293–2299
5. Domingues MJ, Cao HM, Heazlewood SY,
Cao B, Nilsson SK (2017) Niche extracellular
matrix components and their influence on
HSC. J Cell Biochem 118:1984–1993
6. Boulais PE, Frenette PS (2015) Making sense
of hematopoietic stem cell niches. Blood
125:2621–2629
7. Wilson A, Trumpp A (2006) Bone-marrow
haematopoietic-stem-cell niches. Nat Rev
Immunol 6:93–106
8. Samlowski WE, Daynes RA (1985) Bonemarrow engraftment efficiency is enhanced by
competitive-inhibition of the hepatic asialoglycoprotein receptor. Proc Natl Acad Sci U S A
82:2508–2512
9. Larcombe MR, Manent J, Chen J, Mishra K,
Liu X, Nefzger CM (2019) Production of high
titer lentiviral particles for stable genetic modification of mammalian cells. Methods Mol Biol
1940:47–61
10. Bertoncello I, Williams B (2001) Analysis of
hematopoietic phenotypes in knockout mouse
models. Methods Mol Biol 158:181–203
Chapter 10
Genetic Manipulation and Selection of Mouse Mesenchymal
Stem Cells for Delivery of Therapeutic Factors In Vivo
Donald S. Sakaguchi
Abstract
Bone marrow-derived mesenchymal stem cells (MSCs) hold great potential as an ex vivo cellular system for
delivery of therapeutic proteins to the diseased or damaged central nervous system (CNS). This adult stem
cell population has considerable translational potential for autologous transplantation due to lack of ethical
concerns, accessibility, multipotent nature, and plasticity. Here we describe a methodology and outline a
strategy using lentiviral vectors for producing lines of MSCs hypersecreting neurotrophic growth factors
(e.g., brain-derived neurotrophic factor (BDNF) and/or glial cell line-derived neurotrophic factor
(GDNF)) together with a reporter protein such as green fluorescent protein (GFP) that may be used for
in vitro and in vivo neuroprotection bioassays. This approach provides exciting opportunities for basic
research and proof-of-concept studies.
Key words Transplantation, Retinal transplant, Mesenchymal stem cells, Adult stem cells, Neurotrophic factors, Neuroprotection
1
Introduction
A serious problem with developing useful therapies for treatment of
neurodegenerative diseases or injuries to the brain is in implementing effective methods that prevent further degeneration and also
facilitate recovery of function. Stem cell transplantation offers a
novel and extremely exciting therapeutic approach. In this chapter,
a strategy is highlighted employing bone marrow-derived mesenchymal stem cells (MSCs) which hold great potential as an ex vivo
system for delivery of therapeutic proteins to the diseased or damaged CNS (Fig. 1). Mesenchymal stem cells are multipotent and
have the ability to differentiate into adipocytes, chondrocytes,
myoblasts, and osteocytes [1, 2]. They can be routinely isolated
from large bones in rodents and are easily maintained in vitro.
Significantly, MSCs can be engineered to produce exogenous therapeutic and growth factors targeted for long-term delivery for
neuroprotective agents to the injured CNS, including the retina
Ivan Bertoncello (ed.), Mouse Cell Culture: Methods and Protocols, Methods in Molecular Biology, vol. 1940,
https://doi.org/10.1007/978-1-4939-9086-3_10, © Springer Science+Business Media, LLC, part of Springer Nature 2019
143
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Donald S. Sakaguchi
Fig. 1 Genetic manipulation and selection of mouse bone marrow-derived mesenchymal stem cells (MSCs) for
delivery of neurotrophic factors in vivo. (a) Bone marrow MSCs are routinely isolated from the long bones of
rodents. (b) The MSCs are isolated based on their selective adherence to plastic tissue culture surfaces. (c)
Lentiviral vectors can mediate the efficient delivery, integration, as well as stable expression of transgenes in
dividing and nondividing cells, either in vitro or in vivo. (d) An important step in this strategy is to perform a
series of in vitro assays for transgene expression including ELISAs to determine quantities of the secreted
neurotrophic factors and functional bioassays to determine specific biological activity of the secreted
neurotrophic factors. To examine whether biologically active BDNF and GDNF are produced following lentiviral
vector-mediated transduction, conditioned medium from lentiviral-transduced MSCs should be used in in vitro
bioassay/s [3, 9–12]. In general, neurite outgrowth assays from embryonic or neonatal mouse or rat dorsal
root ganglia or neural-like cell lines (RGC-5 or PC-12) can be effective bioassay systems. (e) When generating
multiple lines of engineered MSCs, it is important to determine cell health and viability. Implementation of a
high-content screening system may be very useful for continued experiments. (f) It is only after the different
lines of engineered MSCs have been characterized in vitro and deemed as good candidates for transplant
studies, should in vivo experiments be initiated. The mammalian retina possesses several advantages and has
served as a unique CNS compartment for transplant studies. Although located in a peripheral location,
embryologically the retina is a part of the CNS and as such has become an important site for studies of
cellular transplantation. (g) After cell transplantation, animals can be submitted for noninvasive assays to
determine visual function (e.g., electroretinography (ERG) recordings, computerized pupillometry. (h) To
assess transplanted cell survival, eye tissue should be collected at the termination of the experiments and
presented for immunohistological analysis. Abbreviations: MSCs mesenchymal stem cells; LV lentiviral; GFP
green fluorescent protein; BDNF brain-derived neurotrophic factor; GDNF glial cell line-derived neurotrophic
factor; ELISA enzyme-linked immunosorbent assay; ERG electroretinogram
Genetic Manipulation and Selection of MSCs
145
[3–5]. From a clinical standpoint, MSCs can be isolated from the
patient and thus serve as an autologous cellular therapy. In addition, because MSCs express intermediate to low levels of MHC
Class I or II antigens, they are suitable for use in allogeneic transplantation procedures [6, 7]. Also, unlike pluripotent stem cells
(e.g., embryonic stem cells or induced pluripotent stem cells),
MSCs do not form teratomas following transplantation. Furthermore, since the MSCs would be isolated from adults, they evoke
little to no moral and ethical objections. Taken together, these
advantages support the notion that MSCs are excellent candidates
for genetic engineering and cellular transplants into damaged CNS
environments [8].
Here it is demonstrated that, as a strategy for stem cell-based
therapy, MSCs can be engineered using lentiviral vectors to overexpress neurotrophic factors (e.g., brain-derived neurotrophic factor (BDNF) and/or glial cell line-derived neurotrophic factor
(GDNF)) together with a reporter protein such as green fluorescent protein (GFP) [3, 9–12]. Lentiviral vectors can mediate the
efficient delivery, integration, as well as stable expression of transgenes in dividing and nondividing cells, either in vitro or in vivo.
Replication-deficient recombinant lentiviruses are widely used in
research laboratories and have become important tools for gene
delivery into a broad range of mammalian cells including MSCs,
neural stem cells, neurons, lymphocytes, and macrophages.
Although the modified lentivirus is still capable of infecting cells,
the required genes for producing new viral particles are lacking. As
such, this approach provides exciting opportunities for basic
research and proof-of-concept studies and the possible genetic
treatment of human diseases. These findings show MSCs, infected
with lentiviral vectors encoding BDNF or GDNF, noticeably
increase the release of neurotrophic factors in vitro. Furthermore,
these factors are bioactive and capable of stimulating neurite outgrowth and have provided neuroprotection using in vitro and
in vivo bioassays [3, 9–12].
2
Materials
All cell culture reagents should be prepared using aseptic technique
to ensure sterility.
Prepare and store all cell culture reagents at 4 C (unless
indicated otherwise by the vendor). Follow all waste disposal regulations mandated at your institution by the Environmental Health
and Safety office and your Institutional Biosafety Committee
(or comparable units) when disposing of waste materials.
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Donald S. Sakaguchi
2.1 Mouse MSC
Culturing
1. Mouse bone marrow-derived MSCs can be purchased from a
number of vendors or isolated from the bone marrow of adult
mice as per standard protocols for MSC isolation from rodents
(see Note 1).
2. MSC media: Iscove’s Modified Dulbecco’s Medium (IMDM)
containing 10% hybridoma-qualified fetal bovine serum (FBS)
10% equine serum 2 mM L-glutamine 10,000 U/mL penicillin
and 10,000 μg/mL streptomycin. Stored at 4 C and warmed
to 37 C in a water bath prior to use,
3. Cell detachment solution: 0.05% trypsin and 0.1% EDTA solution (e.g., from Invitrogen/Gibco).
4. Phosphate-buffered saline (PBS): 1 (e.g., from Invitrogen/
Gibco).
5. 0.4% trypan blue: in solution of 0.85% NaCl in tissue culture
water.
6. Ethanol: 70%.
7. Cell freezing medium (1): 65% MSC cell culture medium,
30% mixture of 50:50 mixture of FBS and equine serum, 5%
DMSO (tissue culture grade).
8. Earle’s balanced salt solution (EBSS): 1 (e.g., from Invitrogen/Gibco).
2.2 Engineering
Mouse MSCs Ex Vivo
Using Lentiviral
Vectors
1. MSC media for lentiviral transductions: IMDM, supplemented
with 2% FBS and 12 μg/mL Sequa-brene.
2. Lentiviral constructs are produced by packaging a gene of
interest in a non-replicative retroviral skeleton [13, 14]. Lentiviral vectors may be purchased or designed by a vector core
facility for one’s specific needs.
3. Standard enzyme-linked immunosorbent assay (ELISA) kits
can be purchased and should be used to quantify the amount
of neurotrophic factor/s released from the engineered MSCs.
Follow the vendor protocols accompanying each ELISA kit for
specific neurotrophic factors.
3
Methods
3.1 Culturing
of Mesenchymal Stem
Cells from Frozen Vials
1. Using aseptic techniques in a biosafety cabinet, prepare a
T-75 cm2 tissue culture flask by adding 25 mL of MSC
media. Place the flask into an incubator (high humidity, 5%
CO2 at 37 C) for 20 min in order to equilibrate culture media.
2. Remove cryovial of frozen MSCs from dry ice or liquid nitrogen storage, and spray with 70% ethanol. Place the vial into a
holder and rapidly thaw in a 37 C water bath, until about 95%
thawed. Gently swirl during thawing procedure.
Genetic Manipulation and Selection of MSCs
147
3. Remove the vial from water bath holder, wipe down with 70%
ethanol, and transfer the vial to the biosafety cabinet. Remove
the T-75 flask with MSC media from the incubator, spray with
70% ethanol, and place into the biosafety cabinet.
4. Transfer the contents of the cryovial into the T-75 flask containing the equilibrated MSC media. Rinse the vial several
times with MSC media from the flask to ensure removal of all
MSCs from the cryovial (see Note 2). Gently rock the flask back
and forth to disperse the cells (do not swirl in a circular
motion). Using a permanent fine-tip marker, label the flask
with all pertinent information obtained from the cryovial, and
place into the incubator (high humidity, 5% CO2 at 37 C), and
culture overnight.
5. On the following morning, aspirate the medium containing
nonadherent dead cells, and discard. Rinse the flask with
10 mL of pre-warmed (37 C) PBS. Aspirate and discard the
PBS rinse. Pipet 3 mL of pre-warmed (room temperature)
trypsin/EDTA cell detachment solution into the flask, gently
rock back and forth to cover the bottom of the flask, and
incubate for 2–3 min at 37 C (see Note 3).
6. Monitor MSC detachment on an inverted microscope. As soon
as the majority of MSCs have detached, add 7 mL of
pre-warmed MSC media to deactivate the trypsin/EDTA solution. Gently pipet the MSC media across the bottom of the
flask 3–4 times to detach remaining MSCs.
7. Transfer the MSC suspension to a 15 mL conical centrifuge
tube. Wash the bottom of the flask with an additional 3 mL of
pre-warmed MSC media, and transfer to the 15 mL conical
tube. Pellet the MSC suspension by centrifuging at 450 g for
10 min (centrifugation at room temperature).
8. Carefully pipet off the supernatant and discard. Using a 5 mL
pipette, add 1 mL of pre-warmed MSC media to the pellet, and
gently triturate to break up and disperse the pellet to a single
cell suspension.
9. Perform a trypan blue viable cell count using a hemocytometer.
10. Plate the MSCs into a T-75 flask containing 12 mL of
pre-warmed and equilibrated MSC media at an initial density
of about 60 cells/cm2, and place into the incubator (high
humidity, 5% CO2 at 37 C).
11. Every 3–4 days, pipet or aspirate off one-half the volume of
media, and add back an equal volume of fresh pre-warmed
MSC media. Monitor the MSCs on an inverted microscope.
Cells should maintain a spindle-shape and not become too
cuboidal and/or flat (Fig. 2).
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Donald S. Sakaguchi
Fig. 2 Mouse mesenchymal stem cells (MSCs) growing in a tissue culture flask.
Cells should maintain a spindle morphology. Scale bar ¼ 100 μm
12. Once the MSCs have reached about 70% confluence (approximately 10–14 days), they should be passaged or frozen down
(see Subheading 3.2 below). For passaging, repeat steps 5–11.
3.2 Freezing MSCs
for Storage
1. Once the MSCs have reached about 70% confluence (approximately 10–14 days), they can be frozen down for long-term
storage. Harvest and collect cells as described in Subheading
3.1, steps 5–7.
2. Carefully pipet off the supernatant and discard. Using a 5 mL
pipette, add 3 mL of cell freezing medium to the pellet, and
gently triturate to break up and disperse the pellet to a single
cell suspension.
3. Aliquot 1 mL of MSC cell suspension in cell freezing medium
to each of three, pre-labeled cryovials (see Note 4). The cryovials containing the MSCs should be placed into a cryofreezing container, and placed into a 80 C freezer overnight
to achieve optimal freezing rate of about 1 C/min.
4. The following day, the cryovials of frozen MSCs should be
transferred from the 80 C freezer to the vapor phase of a
liquid nitrogen freezing unit for long-term storage.
3.3 Lentiviral
Transductions
of MSCs: Determining
and Optimizing
the Multiplicity
of Infection (MOI) (See
Notes 5–11)
1. Harvest MSCs as described in Subheadings 3.1, steps 5–9, and
plate into seven wells of a 24-well tissue culture plate (e.g.,
1000 cells per well).
2. After 24 h of growth, replace the culture media in the seven
wells (of the 24-well plate) with 500 μL per well of MSC media
for lentiviral transductions (IMDM, containing 2% FBS and
12 μg/mL Sequa-brene—prepare about 10 mL of this media).
3. On ice, thaw the aliquot of lentiviral particles that only harbor
GFP (green fluorescent protein—LV-GFP vector), as they can
be easily identified and quantified based on fluorescence. Add
Genetic Manipulation and Selection of MSCs
149
1:1 volume of the MSC media for lentiviral transductions to
the lentiviral aliquot (typically the aliquots of lentivirus are
10–20 μL, based on MOI numbers). For easier handling, the
virus can be diluted with medium and pipetted in higher quantities into the wells. The quantity of the required diluted lentivirus will depend on the level of dilution. MOI dilutions of
1, 2, 5, 10, 15, and 30 are recommended (typically, the MOI
for lentiviral particles ranges from 1 to 30). The seventh well of
MSCs serves as a non-transduced control.
4. After 8 h of exposure of the MSCs to the lentiviral particles, the
media should be changed to fresh MSC growth media.
Engineered MSCs should be maintained as previously
described for 72 h.
5. Acquire images of the cells using an inverted fluorescence
microscope after 24, 48, and 72 h. The rate of GFP-lentiviraltransduced MSCs for each MOI and at each time point should
be determined. The lowest MOI at which all MSCs show the
GFP transgene expression should be used for further experiments (see Note 11).
3.4 Lentiviral
Transductions of MSCs
for Production
of Neurotrophic
Factors
1. Harvest MSCs as described in Subheading 3.1, steps 5–9, and
plate into six-well tissue culture plates at a density of approximately 1500 cells per well, and let MSCs attach for 24 h in the
incubator.
2. After cell attachment period, growth medium is replaced with
MSC media for lentiviral transductions (IMDM, containing 2%
FBS and 12 μg/mL Sequa-brene; ~1.5 mL/well).
3. Prepare lentiviral vectors encoding neurotrophic factors (see
Subheading 3.3, step 3): (1) BDNF (LV-BDNF), (2) GDNF
(LV-GDNF), and (3) GFP (LV-GFP) are added individually or
simultaneously to MSCs at a multiplicity of infection (MOI)
based upon prescreening test (Subheading 3.3, steps 1–5). A
population of control MSCs should be engineered with only
the LV-GFP vector at an MOI to match the viral titer of the
BDNF/GFP (and GDNF/GFP) engineered MSCs.
4. Viral particles are removed by rinsing each well after 8 h of
exposure, and media should be changed to fresh MSC growth
media. Engineered MSCs are subsequently maintained as previously described (see Subheading 3.1, steps 11 and 12). Verification of GFP expression should be determined by imaging
culture wells on an inverted fluorescence microscope.
5. As control and engineered MSCs approach 70–80% confluence, they are expanded to a larger vessel (T-25 cm2 tissue
culture flask) for continued optimal growth and determination
of MSC-derived neurotrophic factor production, secretion,
and bioactivity. Media should be collected and replaced with
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Donald S. Sakaguchi
fresh growth media and cells grown for an additional 24 h.
After the 24 h of additional growth, the MSC conditioned
growth media should be collected and used for ELISA for
determination of neurotrophic factor secretion and/or bioassays or frozen at 20 C for subsequent ELISA/bioassays (see
Notes 12 and 13).
6. When generating multiple lines of engineered MSCs, it is
important to determine cell health and viability. Implementation of a high-content screening system may be very useful for
continued experiments (see Note 14).
7. With continued propagation, engineered MSCs should be frozen down to maintain frozen stocks of the MSCs for future
experiments (see Subheading 3.2).
3.5 Transplantation
of Engineered
Mesenchymal Stem
Cells for Intraocular
Delivery
of Neurotrophic
Factors In Vivo
All animal studies should be conducted in accordance with the
Association for Research in Vision and Ophthalmology (ARVO)
Statement for the Use of Animals in Ophthalmic and Vision
Research, and procedures must be approved by your Institutional
Animal Care and Use Committee (IACUC) (see Note 15).
1. Prepare the cell microinjection apparatus in advance of harvesting cells (see Note 16). The cell injection apparatus consists of a
10 or 20 μL Hamilton syringe held in a microsyringe driver
connected via a fluid-filled (Earle’s balanced salt solution)
polyethylene tube to a beveled glass microinjection pipette
(see Note 17).
2. Harvest MSCs as described in Subheading 3.1, steps 5–7.
Following centrifugation, carefully pipet off the supernatant,
and discard. Using a P-1000 pipette, add 200 μL of Earle’s
balanced salt solution (EBSS) to the pellet, and gently triturate
to break up and disperse the pellet to a single cell suspension.
3. Perform a trypan blue viable cell count using a hemocytometer.
Dilute the cell suspension with EBSS to a density of approximately 50,000 cells/μL, and place on wet ice.
4. Mice (4–6 months of age) are anesthetized with isoflurane
inhalation: 3% at 500 mL/min. Gas flow for induction and
then 1.5–2% isoflurane for maintenance of anesthesia. The
animals are determined to be anesthetized when they no longer
perform the righting reflex (when overturned in the anesthesia
induction chamber, they do not attempt to right themselves)
and are nonresponsive to pain (gentle pinch with forceps on
their hindlimb).
5. While anesthetized and in preparation for the intraocular cell
transplants, the animals are placed on their side on a warm
heating pad.
Genetic Manipulation and Selection of MSCs
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6. Using a P 20 or P 100 micropipetter, pipette 10–15 μL droplet
of cell suspension onto the center of the open lid of a 35 mm
tissue culture plate. The beveled microinjection needle is held
in the droplet of cell suspension and ~5–10 μL of the suspension drawn up into the microinjection needle by applying
negative pressure through the microsyringe driver.
7. Animals receive intraocular injections of stem cells through the
dorsolateral aspect of their right eye while avoiding the lens
during the penetration. 2–4 μL of cell suspension in Earle’s
balanced salt solution (~50,000 cells/μL) is slowly injected
into the vitreal chamber of the eyes (volume of cell suspension
to be injected depends on the age of the animal). The microinjection needle should be held within the eye for approximately
20 s before carefully withdrawing the needle.
8. To prevent potential infection, a small amount of antibiotic
ointment (neomycin and polymyxin B and bacitracin (e.g.,
from Bausch & Lomb Pharmaceuticals) may be applied topically to the eye injection site after the procedure.
9. Animals should be allowed to recover from the anesthesia
before returning to their cage.
10. Animals should be removed from the study if they display
abnormal or unusual behaviors (see Note 18).
11. After cell transplantation, animals can be submitted for computerized pupillometry and/or electroretinography recording
or other noninvasive assays to determine visual function. To
assess transplanted cell survival eye tissue should be collected at
the termination of the experiments and presented for immunohistological analysis.
12. Transplant recipient eyes and control, fellow eye tissues should
be collected after the animals are euthanized in an induction
chamber using CO2 (see Note 19). Once the animals have been
euthanized, the eyes should be quickly collected, cleared of
excess tissues, rinsed, and fixed for histological analysis. A
scalpel should be used to pierce the cornea to permit fixative
penetration into the eyeball.
4
Notes
1. Bone marrow-derived MSCs from different strains (as well as
different genera and species) may differ in their media requirements for optimal growth [15]. As such, it may be necessary to
identify and define optimal growth conditions. The aim of this
protocol is to provide a framework methodology to conduct
experiments using MSCs with goals of genetic modification
and cell transplantation for a cell-based delivery system for
neurotrophic factors.
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Donald S. Sakaguchi
2. Thawed MSCs are very fragile at this stage and should be
transferred directly into the culturing vessel without trying to
wash out the DMSO contained within the freezing media.
Using an excess volume of MSC media at this stage helps to
dilute out the DMSO.
3. Trypsin/EDTA step should be closely monitored since the
MSCs are still quite fragile. Use an inverted microscope to
monitor cell detachment.
4. It is important that the MSCs do not sit at room temperature in
the cell freezing medium for more than about 20 min. Cryovials of MSCs should be transferred to 80 C using a cryofreezing unit.
5. All cell culture work and lentiviral transductions should follow
the guidelines put forth in the National Institutes of Health
(NIH) Guidelines for Research Involving Recombinant DNA
Molecules. Handling the instrumentation and conducting laboratory procedures should adhere to the BSL-2 of the Biosafety
in Microbiological and Biomedical Laboratories (BMBL) 5th
edition
(https://www.cdc.gov/biosafety/publications/
bmbl5/index.htm) guidelines.
6. The aim of this approach is to engineer MSCs to express GFP
(green fluorescent protein) and also to overexpress BDNF
(brain-derived neurotrophic factor) or GDNF (glial cell linederived neurotrophic factor) using lentiviral vectors. The engineered MSCs would be a potent candidate for stem cell-based
therapies of neurodegenerative diseases. The lentiviral vectors
used are replication incompetent. These viruses have key elements of their genome supplied in trans during the viral production process in order to eliminate recombination events
that would lead to an active virus. In addition, this virus lacks
the cellular machinery needed to package the virus, as well as an
envelope protein to package the virus within. In general, lentiviral vectors display high transduction efficiency and offer maximal biosafety, without sacrificing transduction efficiency.
7. When using lentiviral vectors, be sure to avoid repeated thawing and freezing cycles, as this can lead to a decrease in viral
titer. Additionally, when thawing, perform on ice, and try and
use vectors immediately. Long-term storage and freezing
should occur either on dry ice or at a temperature of 80 C.
To maintain the quality of the virus, vial contents should be
aliquoted on first use.
8. All culturing work of engineered cells and the engineering of
these cells should be performed in a BSL-2 certified biosafety
cabinet. All culture supplies and materials should be autoclaved
prior to disposal. Culturing surfaces should be cleaned with
bleach. Handling of cultures and cells should be performed
Genetic Manipulation and Selection of MSCs
153
under BSL-2 level. All supplies and instruments should be
autoclaved after procedures and/or prior disposal. Bench tops
and other potentially contaminated areas should be cleaned
with bleach.
9. Waste disposal procedures:
(a) Non-sharp waste: All cultures, stocks, and cell culture
materials must be disinfected with 10% bleach and autoclaved prior to being disposed.
(b) Sharps waste: All needles, syringes, razors, scalpels, Pasteur pipettes, and pipette tips must be disposed of in a
puncture-resistant sharps container. Sharps containers
should not be filled beyond 2/3 of their capacity.
(c) Decontamination procedures: All materials that have
come into contact with lentiviral vectors should be disinfected using a freshly prepared 10% bleach solution before
disposal. Additionally, all work surfaces must be disinfected with 10% bleach once work is completed. (Note:
A 15 min contact time is required for decontamination.)
10. When a cell type is being transduced with a lentivirus for the
first time, it is recommended to set up an initial experiment
with different MOIs using a lentivirus encoding a fluorescent
reporter protein such as green fluorescent protein (GFP). The
MOI is used to describe the number of viral particles needed to
infect a cell. The MOI can differ considerably for different cell
types, and therefore it is recommended to initially determine
the optimal MOI for each cell type under study. Due to the
random nature of integration of lentiviral vectors into the host
genome, varying levels of expression may be observed within
different infected colonies. Testing of multiple colonies and
conditions provides a reasonably straightforward and simple
method to determine optimal degree of expression.
11. High quantities of the virus (higher MOIs) may compromise
cell health and as such, consider selecting a lower MOI, so as to
avoid cytotoxic artifacts.
12. Analysis of transgene expression and secretion of neurotrophic
factors: Standard enzyme-linked immunosorbent assay
(ELISA) kits should be used to quantify the amount of neurotrophic factor/s released from the engineered MSCs. Follow
the vendor protocols accompanying each ELISA kit for specific
neurotrophic factors and compare to known amounts of
authentic BDNF and GDNF, respectively. Long-term secretion of BDNF and GDNF from the lentiviral-transduced MSCs
should be monitored by collecting and concentrating media
from cultures at 1–3 days and at multiple weeks post-viral
transduction (e.g., 1, 2, and 4 weeks post-viral transduction).
Medium from non-transduced MSCs and control MSCs transduced with the LV-GFP only vector should also be tested.
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Donald S. Sakaguchi
13. In vitro analysis of transgene expression: An important step is
to determine if the secreted neurotrophic factors possess specific biological activity. To examine whether biologically active
BDNF and GDNF are produced following lentiviral vectormediated transduction, conditioned medium (CM) from
LV-transduced MSCs (for 6 days) should be used in in vitro
bioassay/s [3, 9–12]. In general, neurite outgrowth assays
from embryonic or neonatal mouse or rat dorsal root ganglia
or neural-like cell lines (RGC-5 or PC-12) can be effective
bioassay systems [3, 9–12].
14. An essential step in developing cell-based therapeutic trophic
factor delivery systems is to determine the normal health of the
engineered MSCs. As such, implementation of image-based
high-content screening may be very useful when generating
multiple lines of engineered MSCs. This technology permits
automated image acquisition and analysis and is well-suited for
stem cell research applications but is beyond the scope of this
chapter, and the reader is referred to the literature [10].
15. All animals should be handled in accordance with the Guide for
the Care and Use of Laboratory Animals (referred to as the
Guide) [16] and follow all guidelines appropriate to your institution and any other applicable regulations. Furthermore, the
Guide recommends that the number of animals should be the
minimum number required to obtain statistically valid results.
A power analysis is strongly encouraged to justify group sizes
when appropriate.
16. When assembling the cell microinjection system, try and maintain sterility by wiping all parts with Kimwipes moistened with
70% ethanol. Clean tubing by flushing with sterile EBSS.
17. A conventional horizontal or vertical pipette puller is generally
used to make the microinjection pipettes using glass microinjection pipettes. Though sterile forceps may be used to break
the tip of the injection micropipette so the opening is about
~10 μm wide, beveling the tip is the recommended and preferred method so as to create a microinjection needle with a
sharp tip yet has an opening of ~10 μm wide. A larger opening
is required to draw up and to inject the cells into the eyes.
18. Following transplants, animals should be assessed and monitored for normal behavior. Parameters to be monitored should
include normal activity, overall responsiveness, appearance,
skin color, and eating and drinking habits. Animals should be
removed from the project if they do not eat or drink and display
excessive weight loss and dehydration, display labored breathing, or have impaired movements. In general, based on prior
studies, the recipients have tolerated cell transplants very well
Genetic Manipulation and Selection of MSCs
155
and have not displayed excessive weight loss or other behavioral anomalies.
19. All Euthanasia Guidelines for Research and Teaching should be
followed. The National Research Council Guide for the Care
and Use of Laboratory Animals states that methods of euthanasia must “induce rapid unconsciousness and death without
pain or distress.” Institutions should observe the above-stated
method definition and adhere to the euthanasia guidelines
specified in the American Veterinary Medical Association
Guidelines on Euthanasia (AVMA).
References
1. Prockop DJ (1997) Marrow stromal cells as
stem cells for nonhematopoietic tissues. Science 276:71–74
2. Ohishi M, Schipani E (2010) Bone marrow
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3. Harper MM, Grozdanic SD, Blits B, Kuehn
MH, Zamzow D et al (2011) Transplantation
of BDNF- secreting mesenchymal stem cells
provides neuroprotection in chronically hypertensive rat eyes. Invest Ophthalmol Vis Sci
52:4506–4515
4. Levkovitch-Verbin H, Sadan O, Vander S,
Rosner M, Barhum Y et al (2010) Intravitreal
injections of neurotrophic factors secreting
mesenchymal stem cells are neuroprotective in
rat eyes following optic nerve transection.
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5. Sasaki M, Radtke C, Tan AM, Zhao P, Hamada
H et al (2009) BDNF-hypersecreting human
mesenchymal stem cells promote functional
recovery, axonal sprouting, and protection of
corticospinal neurons after spinal cord injury. J
Neurosci 29:14932–14941
6. Aggarwal S, Pittenger MF (2005) Human
mesenchymal stem cells modulate allogeneic
immune cell responses. Blood 105:1815–1822
7. Le Blanc K, Tammik C, Rosendahl K,
Zetterberg E, Ringden O (2003) HLA expression and immunologic properties of differentiated and undifferentiated mesenchymal stem
cells. Exp Hematol 31:890–896
8. Sandquist EJ, Uz M, Sharma AD, Patel BB,
Mallapragada SK, Sakaguchi DS (2016) Stem
cells, bioengineering and 3-D scaffolds for nervous system repair and regeneration. In: Zhang
LG, Kaplan D (eds) Neural engineering: from
advanced biomaterials to 3D fabrication techniques. Springer, New York, pp 25–82
9. Harper MM, Adamson L, Blits B, Bunge MB,
Grozdanic SD, Sakaguchi DS (2009) Brainderived neurotrophic factor released from engineered mesenchymal stem cells attenuates glutamate - and hydrogen peroxide-mediated
death of staurosporine-differentiated RGC-5
cells. Exp Eye Res 89:538–548
10. Sharma AD, Brodskiy PA, Petersen EM,
Dagdeviren M, Ye EA et al (2015) High
throughput characterization of adult stem cells
engineered for delivery of therapeutic factors
for neuroprotective strategies. J Vis Exp (95):
e52242.
11. Ye E-A, Chawla SS, Khan MZ, Sakaguchi DS
(2016) Bone marrow-derived mesenchymal
stem cells (MSCs) stimulate neurite outgrowth
from differentiating adult hippocampal progenitor cells. Stem Cell Biol Res 3:3
12. Bierlein del la Rosa M, Sharma AD, Mallapragada SK, Sakaguchi DS (2017) Transdifferentiation of BDNF-hypersecreting MSCs
significantly enhances BDNF secretion and
Schwann cell marker proteins. J Biosci Bioeng
124:572–582
13. Delenda C (2004) Lentiviral vectors: optimization of packaging, transduction and gene
expression. J Gene Med 6:S125–S138
14. Chang LG, Gay EE (2001) The molecular
genetics of lentiviral vectors - current and future
perspectives. Curr Gene Ther 1:237–251
15. Peister A, Mellad JA, Larson BL, Hall BM,
Gibson LF, Prockop DJ (2004) Adult stem
cells from bone marrow (SCs) isolated from
different strains of inbred mice vary in surface
epitopes, rates of proliferation, and differentiation potential. Blood 103:1662–1668
16. National Research Council (2011) Guide for the
care and use of laboratory animals, 8th edn. The
National Academies Press, Washington DC
Chapter 11
Isolation and Culture of Primary Mouse Middle
Ear Epithelial Cells
Apoorva Mulay, Khondoker Akram, Lynne Bingle, and Colin D. Bingle
Abstract
Epithelial abnormalities underpin the development of the middle ear disease, otitis media (OM). Until now,
a well-characterized in vitro model of the middle ear (ME) epithelium that replicates the complex cellular
composition of the middle ear has not been available. This chapter describes the development of a novel
in vitro model of mouse middle ear epithelial cells (mMECs), cultured at the air-liquid interface (ALI). This
system enables recapitulation of the characteristics of the native murine ME epithelium. We demonstrate
that mMECs undergo differentiation into the varied cell populations seen within the native middle ear.
Overall, our mMEC culture system can help better understand the cell biology of the middle ear and
improve our understanding of the pathophysiology of OM. The model also has the potential to serve as a
platform for validation of treatments designed to reverse aspects of epithelial remodeling underpinning OM
development.
Key words Middle ear epithelial cells (mMECs), Air-liquid interface (ALI), In vitro, Otitis media,
Middle ear
1
Introduction
The upper airways consist of the trachea, nasopharynx (NP), Eustachian tube (ET), middle ear, and mastoid cells surrounding the
middle ear. The middle ear epithelium is a continuation of the
upper airways through the ET [1]. Therefore, the physiology and
immune defenses of the upper respiratory tract (URT) are very
similar to that of the middle ear. The middle ear epithelium provides the first line of defense against invading pathogens and acts as
a physical barrier. It is composed of ciliated cells, secretory cells,
nonsecretory cells, and basal cells. Secretory cells are responsible for
the production of high molecular glycoproteins called mucins
which increase the viscosity of epithelial secretions and are important in trapping pathogens, various antimicrobial proteins such as
lactotransferrin, lysozyme, defensins and surfactants [2, 3], and
other putative multifunctional host defense proteins such as
Ivan Bertoncello (ed.), Mouse Cell Culture: Methods and Protocols, Methods in Molecular Biology, vol. 1940,
https://doi.org/10.1007/978-1-4939-9086-3_11, © Springer Science+Business Media, LLC, part of Springer Nature 2019
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BPIFA1 [4]. Epithelial cells also express receptors, known as pattern recognition receptors (PRRs) such as Toll-like receptors
(TLRs), retinoic acid-inducible gene I-like receptors (RLRs), and
nucleotide-binding oligomerization domain (NODs) [5], which
recognize the invariant features of microbes known as pathogenassociated molecular markers (PAMPs) characteristics such as flagella, lipopolysaccharide (LPS) of Gram-negative bacteria, and
lipoteichoic acid of Gram-positive organisms [6]. Upon PAMP
recognition, most PRRs trigger the release of chemokines and
pro-inflammatory cytokines in order to mount an inflammatory
response and clear the pathogens [7, 8]. Thus the epithelium,
along with its secretions, is involved in maintaining homeostasis
and sterility within the middle ear cavity and epithelial remodelling
characterised by mucociliary metaplasia and infiltration of the middle ear space with inflammatory cells, is a common feature of
inflammatory middle ear diseases such as OM [9].
In most animals, the middle ear is a relatively inaccessible organ
lined by a thin mucociliary epithelium, and sampling of the mucosa
is a terminal procedure. Human middle ear tissue can be acquired
only during surgical procedures, and this limits the amount of
sample available for study of OM. Therefore, in vitro culture of
middle ear epithelial cells is vital for studying the basic cell biology
of the middle ear during homeostatic conditions and during disease
and for developing therapeutic interventions to treat middle ear
diseases. Culturing of middle ear cells in vitro enables maximization
of the available material and allows the effect of modifying culture
conditions to be studied more easily. It also enables studies of hostpathogen interactions. Previously, attempts have been made to
culture middle ear epithelial cells from a number of organisms
including rats [10–12], mice [13], chinchillas [14, 15], gerbils
[16–18], rabbits [19], and humans [20–22]. These studies have
included organ and explant cultures, primary cell cultures, and
development of middle ear cell lines. However, there remains a
lack of a robust in vitro middle ear epithelial model that differentiates into the different epithelial cell types of the middle ear and is
free of fibroblast contamination. This has greatly restricted the
ability to identify the function of different cell types and their
products within the middle ear and limits our understanding of
the pathophysiology of OM development.
In recent years, pulmonary research has been revolutionized by
use of an air-liquid interface (ALI) culture system for the propagation of tracheobronchial epithelial (TBE cells). The exposure of
apical cell surfaces to air and the supply of nutrients from the basal
compartment mimic the in situ arrangement and promote maximal
differentiation. ALI cultures of TBE cells have been generated from
several species [23–26]. More recently, this system has been applied
to the culture of murine nasal epithelial cells, recapitulating the
characteristics of the respiratory epithelium of the nasal passages
Isolation and Culture of Middle Ear Epithelium
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[27]. Studies employing ALI cultures of TBE and nasal epithelial
cells have enhanced our knowledge of airway epithelial biology
tremendously, by shedding light on various aspects such as cellular
differentiation and secretion, [28, 29], cell-cell communication
[30] inflammatory signaling [31], pathogenesis of infections
[32–34], drug transport [35, 36], effects of environmental and
occupational pollutants [37, 38], and genetic disorders such as
cystic fibrosis [39] and primary ciliary dyskinesia [40].
The mouse is an excellent model of OM owing to significant
functional and anatomical similarity between the ears of humans
and mice [41] and the high percentage of genetic conservation
between humans and mice [42]. Several disease-susceptibility
genes, thought to be involved in OM, have been disrupted in
mice, in an attempt to study their role in the pathogenesis of the
disease giving rise to the development of a number of mouse
models of OM [43]. We report here the development of a novel
in vitro primary model of the mouse middle ear epithelium using
ALI culture to systematically characterize the different cell types
present in the middle ear. This culture system can be utilized to
study host-pathogen interactions within the middle ear and thus
has the potential to allow investigation of the mechanisms of OM
pathogenesis.
2
Materials
2.1
Mice
All animal procedures in this study are carried out in accordance to
the procedures enlisted under the Home Office project and personal licenses. Mice are housed in individually ventilated cages in
specific pathogen-free (SPF) conditions, fed on an irradiated diet
and water containing 25 ppm chlorine, maintained in a 12-h light/
dark cycle at 21 C (2 C) and 55% (10%) humidity and
inspected daily. We typically use C57Bl/6 mice between 8 and
12 weeks and pool cells from six mice (12 middle ears) for a single
batch of cultured cells.
2.2
Media
Two closely related media are used for culturing mMECs: the
mMEC plus medium is used for proliferation of cells, and
mMEC-SF (serum-free) medium is used for differentiation of
cells. The base media and all stock additives can be purchased
commercially.
2.2.1 Stock Additives for
Media Preparation
First all the stock solutions can be prepared as follows:
1. Bovine serum albumin (BSA; 100 mg/mL): Dissolve 2.5 g of
BSA in 25 mL sterile HBSS to form a stock solution of
100 mg/mL. Filter through a 0.2 μm filter, and store in
5 mL aliquots at 20 C.
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2. Hank’s balanced salt solution (HBSS)/BSA: Dissolve BSA
(Fraction V) in HBSS at 1 mg/mL concentration. Filter and
store at 4 C.
3. Retinoic acid: First, prepare retinoic acid solution A (RA-A;
5 mM) by dissolving one vial of 100 mg retinoic acid in
66.6 mL 100% ethanol under dim light. Retinoic acid is light
sensitive. Filter the solution, and store in 1 mL aliquots at
80 C for up to 12 months. Prepare retinoic acid stock
solution B (RA-B; 50 mM) by adding 1 mL of RA-A to 9 mL
of 100% ethanol. Store 1 mL aliquots at 80 C, protected
from light. RA-B can be stored up to 1 week at 20 C while
using to supplement media.
4. Insulin: Dissolve 100 mg insulin in 50 mL of 4 mM HCl to
form a stock solution of concentration 2 mg/mL. Filter and
store in 1250 μL aliquots at 20 C.
5. Transferrin: Dissolve 100 mg transferrin in 20 mL of
HBSS/BSA solution to form a stock solution of concentration
5 mg/mL. Filter and store in 250 μL aliquots at 20 C.
6. Mouse epidermal growth factor (EGF): Prepare a 5 μg/mL
stock solution by dissolving 100 μg EGF in 20 mL of HBSS/
BSA solution. Filter and store in 1250 μL and 250 μL aliquots
at 20 C.
7. Cholera toxin (CT): Prepare a 100 μg/mL stock solution by
dissolving 0.5 mg cholera toxin in 5 mL of HBSS/BSA solution. Filter and store in 250 μL and 62.5 μL aliquots at 20 C.
8. Bovine pituitary extract (BPE): Store in aliquots containing
7.5 mg total protein at 80 C.
2.2.2 Preparation of
mMEC Plus Media
First prepare 500 mL of mMEC basic media by adding 10 mL
solution of penicillin (100 μg/mL) and streptomycin (100 μg/
mL) in the total volume of DMEM/F-12 HAMs media (Life Technology). To prepare 50 mL mMEC plus media, add 250 μL of the
insulin stock, 50 μL of transferrin stock, 50 μL of the CT stock,
250 μL of the EGF stock, 1.5 mg of the BPE stock, 2.5 mL of FBS to
46.7 mL of mMEC basic media. mMEC basic media can be stored at
4 C for up to 1 week. Add 5 μL of RA-B just before using the media.
2.2.3 Preparation of
mMEC-SF Media
To prepare 50 mL mMEC-SF media, add 125 μL of the insulin
stock, 50 μL of transferrin stock, 12.5 μL of the CT stock, 50 μL of
the EGF stock, 1.5 mg of the BPE stock, 500 μL of BSA to
49.1 mL of mMEC basic media. mMEC-SF can be stored at 4 C
for up to 1 week. Add 5 μL of RA-B just before using the media.
2.3 Other Equipment
and Reagents
1. Anesthetic: pentobarbital (50 mg/mL).
2. Dissection instruments including blunt and fine dissection scissors, fine forceps, and storkbill forceps from Surgipath, Leica
Biosystems.
Isolation and Culture of Middle Ear Epithelium
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3. Dissecting microscope.
4. 15 mL and 50 mL falcon tubes.
5. 60 mm surface treated tissue culture dishes.
6. 24-well format tissue culture dishes.
7. Transwell inserts: 0.4 μm pore sized transparent PET (polyethylene terephthalate) in a 24-well supported transwell format.
8. DNase solution: 0.5 mg/mL pancreatic DNase I and 1 mg/
mL BSA in mMEC basic media. First add 2 mL of BSA stock
(100 mg/mL) with 198 mL of mMEC basic media, and then
dissolve 100 mg of DNase in it to prepare 200 mL of solution.
Sterile filter and aliquot 5 mL/vial, and store at 20 C.
9. Pronase solution: Make fresh 0.15% (w/v) pronase solution
each time just before use by dissolving 0.015 g of pronase in
10 mL of mMEC basic media, and mix gently by rocking the
tube. Aliquot required amount in two 15 mL centrifuge tubes
before use. 5 mL of pronase solution is sufficient for middle
ears from six mice.
10. Collagen 1 solution for coating: Make 10 mL of 50 μg/mL
collagen 1 solution for coating the transwell membranes by
reconstituting 148 μL of stock rat tail collagen 1 solution
(3.37 mg/mL) in total 10 mL of HBSS, and store at 4 C.
11. mMEC basic 10% FBS media: Add 5 mL of FBS to 50 mL of
mMEC basic media.
12. Rho kinase inhibitor Y-27632 dihydrochloride (ROCKi).
3
Methods
3.1 Collagen Coating
of Transwell
Membranes
1. Add 150 μL of the prepared collagen solution on each upper
chamber of transwell, and incubate for 4 h in the incubator at
37 C, 5% CO2.
2. Aspirate the collagen solution, and allow the transwell air dry
(at least for 5 min). Then wash them with PBS three times to
remove extra collagen.
3. Allow the transwell to air dry, and then transwells are ready to
use. Collagen coated transwells are stable for 2 weeks.
3.2 Harvesting
Mouse Middle Ear
Cavities
Figure 1 gives a step-by-step illustration of the dissection protocol
to harvest mouse middle ear cavities or bullae.
1. Euthanize mice by terminal intraperitoneal injection of 100 μL
of anesthetic and exsanguinate them by cutting the inferior
vena cava.
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Fig. 1 Dissection and harvesting of the mouse middle ear cavity. Wild-type C57Bl/6J or C3HeH mice (a) were
decapitated (b). The head was skinned (c) and skull cap removed (d). Dorsal (e) and ventral (f) view of the
head after removing the brain, showing the bullae or middle ear cavities, MECs. Bisected head showing
the outer ear, OE (g). Under dissecting microscope, the bullae were separated from surrounding tissue (h). The
MEC is attached to the outer ear cavity (OEC) at the tympanic membrane (Tm) and tapers toward the opening
of the Eustachian tube (ET) near its posterior end (i). Removal of the OEC and the Tm reveals the cochlea of
the inner ear on the ventral side of the MEC (j). The cup-shaped MEC was detached from the inner ear (k).
Tissue was dissected along the dotted lines. Figure reproduced with permission from Disease Models and
Mechanisms [45]
2. Under direct visualization, decapitate mice, incise the skin at
the nape of the neck, cut the skin anteriorly, and peel it to
expose the bony surface of the skull and the nose.
3. Detach the mandible with a pair of blunt scissors.
4. Under a dissecting microscope, with the nares facing away,
gently open the skullcap with a pair of fine forceps, and remove
the brain. Removal of the brain exposes the anterior most part
of the skull base, which is attached to the posterior most part of
the nasal cavity.
5. Bisect the skull at the midline, and orient each half with the
opening of the ear facing upward. Any muscle, soft tissue, and
remnant hair surrounding the ear should be removed using fine
Isolation and Culture of Middle Ear Epithelium
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dissecting scissors and forceps, leaving the middle ear cavity
(bulla), still attached to the outer ear canal and the inner ear.
6. Further clean the bony shell of the bulla of any attached extraneous tissue.
7. The outer ear canal appears a shade lighter than the middle
ear cavity. Gently break it away from the bulla using
storkbill forceps. The tympanic membrane (Tm) and the ossicles usually detach from the bulla along with the outer ear
canal. Alternatively, physically remove them using fine storkbill
forceps.
8. Lastly, lift the cup-shaped bulla away from the inner ear, and
add it to a tube containing the pronase solution (see Note 1).
For each batch of cells, we harvest middle ear cavities from
approximately six mice, and pool them in a single tube.
3.3 Isolation of
Middle Ear Epithelial
Cells from the
Harvested Tissue
The protocol for isolation, culture and differentiation of
mouse middle ear epithelial cells (mMECs) was adapted from a
method for isolation of mouse tracheal epithelial cells (mTECs)
[25, 44].
1. After harvesting the middle ear cavities, subject them to overnight proteolysis at 4 C in the pronase solution (see Note 2).
2. The next day, neutralize the pronase by the addition of 10%
FBS, and gently agitate the samples by inverting the tube
approximately 25 times.
3. Transfer the samples to 2 mL of fresh mMEC basic 10% FBS
media, invert the tube again 25 times, and repeat this process a
third time. The combined proteolytic and mechanical actions
lead to dissociation of the epithelial cells from the tissue.
4. Combine the media from the three tubes in a fresh 15 mL
falcon tube, and centrifuge at 500 g for 10 min at 10 C.
Resuspend the cell pellet in 1 mL of media containing 1 mg/
mL bovine serum albumin (BSA) and 0.5 mg/mL DNase I
(Sigma-Aldrich).
5. Assess the cell viability and number using trypan blue staining
and a hemocytometer. This will give an indication of the total
number of live cells harvested.
6. Centrifuge the cells were centrifuged at 500 g for 5 min at
10 C, and resuspended the pellet in 5 mL of mMEC basic 10%
FBS media.
7. In order to remove contaminating fibroblasts from epithelial
cells, perform a differential adherence step by plating the cells
on 60 mm tissue culture dishes at 37 C in a 5% CO2 incubator
for 3–4 h. Fibroblasts attach to the plastic faster and the
non-adherent epithelial cells after 4 h can be collected,
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Apoorva Mulay et al.
centrifuged at 500 g for 5 min at 10 C, and resuspended in
1 mL of mMEC plus media: mMEC basic media (see Note 3).
8. Perform a second cell count to determine the total number of
live epithelial cells/mL of media. The average number of
mMECs isolated is 74,667 10,621/bulla (n ¼ 12 Wt batches).
9. Seed the cells at a density of 1 104 cells/collagen-coated
transwell membrane in the presence of 10 μM of Rho kinase
inhibitor Y-27632 dihydrochloride. We have found that addition of ROCKi enables the use of a seeding density that is
5 times lower than that required when ROCKi is not used.
Usually, 30–35 transwells can be seeded from each batch of six
mice (see Note 4).
3.4 Culture of Mouse
Middle Ear Epithelial
Cells
1. Culture the cells initially in submerged conditions in mMEC
plus (proliferation media) with 300 μL of media in the top
chamber and 700 μL in the bottom chamber.
2. Perform media changes every 48 h, until the cells become
completely confluent (see Note 5); whereafter media from the
top chamber should be removed, and media in the bottom
chamber should be replaced with 700 μL mMEC-SF differentiation media. This system of culture, with media in the bottom
culture and apical surfaces of cells exposed to air, is known as
ALI (air-liquid interface) culture, and it promotes maximal
differentiation of cells by mimicking the in vivo physiology
(see Note 6). We routinely culture cells at ALI for 14 days to
enable complete differentiation.
3. Change mMEC-SF media every 48 h. On weekends, media can
be changed on Friday afternoon and first thing on Monday
mornings (see Note 7).
4. Wash the apical surfaces of cells with 200 μL of sterile, warm
HBSS to clear any cellular secretions and mucous deposition
(see Note 8).
5. There are a number of ways to evaluate successful differentiation of mMECs using the ALI system. At the required time
points, cells can be lysed in 250 μL of TRIzol reagent (SigmaAldrich) for RNA extraction and transcriptional analysis. Transwell membranes can be fixed by incubating in 10% paraformaldehyde for 30–45 min for studying the localization of proteins
by immunofluorescent staining. Secreted proteins can be
assessed by collection of 48-h apical washes from the cells.
Routine methods for scanning and electron microscopy can
also be used to examine the morphology of the differentiated
cells. Typically, we collect samples for transcriptional or proteomic analysis at ALI Day 0 (submerged Day 10), Day 3, Day
7 and Day 14. Figure 2 gives a brief summary of the cell culture
system.
Isolation and Culture of Middle Ear Epithelium
165
Fig. 2 Primary culture of mouse middle ear epithelial cells (mMECS). Timeline for culture of mMECs is shown
above (a). Bullae were dissected and treated with pronase for dissociation of the middle ear epithelial cells,
and fibroblasts were excluded from culture by differential adherence to plastic. Epithelial cells were grown in
submerged culture till confluence, before ALI was induced. Samples for transcriptional and proteomic analysis
were collected at regular time points. Phase contrast images showing cells in culture under 10 magnification (b–i). In the proliferative submerged conditions, a small number of cells attached to form epithelial islands
3 days after seeding (b). The cells proliferated faster from day 5 (c) through day 7 (d) and formed a confluent
monolayer at day 9. This was termed as ALI Day 0 (e). Morphology of cells changed from ALI Day 3 (f), and
clusters of compactly arranged cells started forming at ALI Day 7 (g). ALI Day 14 cultures were composed of
flat polygonal and compactly clustered pseudostratified cells with active cilia. White arrowheads mark
elevated ciliated cells, and asterisk marks flatter polygonal cells (h). Fibroblasts cultured on plastic plates
through differential adhesion method. (i) Figure reproduced with permission from Disease Models and
Mechanisms [45]
Our recently published report [45] describes the assessment
of differentiation of mMECs using the ALI system and their use as a
model system to study infection by otopathogen, nontypeable
Haemophilus influenzae.
166
4
Apoorva Mulay et al.
Notes
1. Care should be taken to clean the bulla cavity of any extraneous
attached tissue, and complete removal of the tympanic membrane should be ensured to avoid contamination of mMECs by
unwanted cell types and to reduce the number of contaminating fibroblasts.
2. We have observed that it is best to prepare the pronase solution
fresh each time.
3. Fibroblasts attached to the petri dish during the differential
adherence step can be expanded using mMEC basic+10% FBS
media. We passage the fibroblasts at least twice in T25 flasks to
obtain a pure line. These fibroblasts can be used as a negative
control for epithelial markers.
4. Primary middle ear epithelial cells are mortal, and the seeding
density greatly influences their growth characteristics. Moreover, attachment and growth of cells from different batches of
mice may vary. Therefore typically, generous seeding densities
of mMECS on transwell membranes are required to obtain
confluent and consistent cultures that successfully differentiate.
For determination of optimal seeding densities, we plated cells
on both tissue culture plastic (2.5 104 cells/well) and collagen coated 0.4 μm pore sized transwell membranes at various
initial densities. We found that a minimum seeding density of
5 104 without ROCKi and/or 1 104 cells with ROCKi is
required to establish a confluent monolayer.
5. We have observed that the rate of cell growth is slower just after
seeding and the cells grow more rapidly after day 5 in
submerged culture. Normally cells take 9–10 days to reach
complete confluence.
6. Formation of a confluent monolayer during the initial
submerged culture is a prerequisite for establishing an ALI. If
the cells are not completely confluent, media from the basal
chamber can seep into the apical chamber and disrupt the
formation of an air-liquid interface. This leads to poor differentiation of cells. It is advisable to measure transepithelial
resistance (TER) to verify the formation of a tight monolayer.
7. mMEC plus and MMEC-SF media can be stored for up to
1 week at 4 C.
8. Care should be taken to warm media during media changes and
HBSS before washing apical washes to 37 C.
Isolation and Culture of Middle Ear Epithelium
167
Acknowledgments
This work was supported by a University of Sheffield PhD Studentship (supervised by LB and CDB) and funds from MRC Harwell.
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Chapter 12
Isolation and Propagation of Lacrimal Gland Putative
Epithelial Progenitor Cells
Helen P. Makarenkova and Robyn Meech
Abstract
We present a protocol for isolation of putative epithelial progenitor cells from mouse lacrimal gland (LG) by
fluorescence-activated cell sorting (FACS). Isolated LG epithelial progenitor cells can be cultured as 3D
reaggregates within extracellular matrix gel or plated as a monolayer. 3D cultures could be maintained for
several days and then dissociated with trypsin and plated as monolayer cultures, processed for analysis (e.g.,
mRNA/protein expression) and/or used for transplantations. Our goal is to provide researchers with a
method that can be used as is or modified if isolation of other LG epithelial cell types is required.
Key words Lacrimal gland, Epithelial progenitor cells, FACS, Isolation, 3D cultures
1
Introduction
Aqueous deficiency dry eye (ADDE) is characterized by a lack of
tear secretion from the lacrimal glands (LGs). ADDE affects
millions of Americans causing a debilitating loss of visual acuity,
ocular surface irritation, and adverse lifestyle changes. Currently
there is no cure and no satisfactory treatment for ADDE. One of
the new arising treatments for different pathologies, including
ocular diseases, is LG stem/progenitor cell transplantation
[1–8]. Our recent research shows that among c-kit-positive epithelial cell populations sorted from mouse LGs, the c-kit+dim/
EpCAM+/Sca1/CD34/CD45- cells are a putative epithelial
(acinar and ductal) cell progenitor (EPCP) population [4]. Isolated
EPCPs express pluripotency factors and markers of the epithelial
cell lineage Runx1 and EpCAM and form branches when grown in
reaggregated 3D cultures. When transplanted into injured or diseased LG (e.g., in the thrombospondin-1 null (TSP-1/) mouse
model of Sjogren’s syndrome), they have been shown to restore the
functional epithelial components of the LG [4]. Isolation and
further analysis of LG stem/progenitor cell function would open
new therapeutic possibilities to treat ADDE.
Ivan Bertoncello (ed.), Mouse Cell Culture: Methods and Protocols, Methods in Molecular Biology, vol. 1940,
https://doi.org/10.1007/978-1-4939-9086-3_12, © Springer Science+Business Media, LLC, part of Springer Nature 2019
169
170
2
Helen P. Makarenkova and Robyn Meech
Materials
1. Laboratory mice (four mice per sample), C57BL/6 strain,
4–6 weeks of age. From 4 mice (8 LGs), around
700,000–1,000,000 cells can be isolated. If more cells are
required, then the number of mice should be increased; however do not pool LG from more than four mice without adjusting the reagent volumes proportionally or cell yield/purity will
be compromised (see Note 1).
2. Isoflurane inhalation anesthetic.
3. Ethanol.
4. Eagle’s minimum
glutamine.
essential
medium
(MEM),
without
5. EpiLife™ Medium, with 60 μM calcium.
6. Fitton-Jackson Modification (BGJb) Medium.
7. AlbuMAX™ I Lipid-Rich BSA.
8. Human Corneal Growth Supplement (HCGS).
9. Transferrin human.
10. Insulin-Transferrin-Selenium (ITS-G) (100X).
11. Recombinant human EGF protein, carrier-free (CF).
12. GlutaMAX™ Supplement.
13. Antibiotic-antimycotic.
14. Collagenase type I.
15. HyClone™ Fetal Bovine Serum.
16. Trypsin from porcine pancreas, lyophilized powder.
17. Pancreatin.
18. Trypsin inhibitor.
19. MgCl2 1 M in H2O.
20. DNase I recombinant, RNase-free.
21. HBSS—Hank’s Balanced Salt Solution—with calcium, magnesium, no phenol red.
22. Red blood cell lysis buffer.
23. Dispase II.
24. Phosphate-buffered saline (PBS) (pH 7.4).
25. Ethylenediaminetetraacetic acid (EDTA).
26. Goat serum.
27. Accutase.
28. Glycerol 99%.
29. Hepes (1 M sterile).
Isolation of Lacrimal Gland Epithelial Progenitors
171
30. NaCl powder.
31. Trizma base.
32. Hydrochloric acid (to adjust Tris buffer pH).
33. CaCl2 (1 M sterile).
34. Corning® Matrigel® Basement Membrane Matrix.
2.1
Antibodies
1. PE Rat Anti-Mouse Ly-6A/E (Sca1), clone E13-161.7.
2. FITC Rat anti-Mouse CD34, clone RAM34.
3. APC-eFluor 780 CD117 (c-kit) Monoclonal Antibody (2B8).
4. APC Anti-Mouse CD326 (EpCAM) Monoclonal Antibody
(G8.8).
5. Alexa Fluor 700 CD45 Rat Anti-Mouse Antibody clone
30-F11.
6. FxCycle Violet stain (40 ,6-Diamidine-20 -phenylindole dihydrochloride (DAPI), 0.5 mg/mL).
2.2 Media and Stock
Solutions
1. Trypsin/pancreatin solution: To prepare 10 mL of pancreatintrypsin in Tyrode’s solution, add trypsin, 0.225 g (final cc is
2.25%), and pancreatin, 0.075 g (final concentration will be
0.75%). Sterile filter (0.22 μm) (change filter if it becomes
blocked), and dispense into 2 mL aliquots stored at 20 C
(see Note 2).
2. Tyrode solution (Ca, Mg free), pH 7.2: NaCl 8 g/L. KCl
0.2 g/L. NaH2PO4 + H2O 0.05 g/L (MW 137,99). Glucose
1 g/L (D(+) glucose, MW 180.16, H2O free). NaHCO3 1 g/
L. Sterile filter Tyrode’s solution and store at 4 C.
3. Stock collagenase type I solution: Weigh out 60 mg collagenase
powder, and dissolve in 10 mL HBSS 1 buffer with Ca and
Mg for a stock concentration of 6 mg/mL, 750 units/mL (see
Note 3). Prepare 500 μL aliquots and store at 70 C for up to
6 months or 4 C for 1 day (see Note 4).
4. Stock dispase type II solution. Weigh out 240 mg dispase
powder, and dissolve in 1 mL of 50 mM Hepes/150 mM
NaCl for a 50 stock solution at final concentration of
120 units/mL (see Note 3). Prepare 80 μL aliquots and store
at 70 C for up to 6 months or 4 C for 1 day (see Note 4).
5. Stock DNase type I solution: Weigh out 5 mg DNase I powder,
and dissolve it in 5 mL solution containing 50% glycerol,
20 mM Tris buffer (pH 7.5), and 1 mM MgCl2 for a stock
concentration of 1 mg/mL, 2000 units/mL (see Note 3).
Prepare by 25–50 μL aliquots and store at 70 C for up to
6 months or 4 C for 1 day (see Note 4).
6. Media I: MEM low glucose supplemented with GlutaMAX
(1) and stored at 4 C.
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Helen P. Makarenkova and Robyn Meech
7. Blocking media: HBSS supplemented with 10% FBS and 1 mM
EDTA
8. FACS buffer: 1 PBS supplemented with 2.5% goat serum and
1 mM EDTA and 25 mM Hepes. (486.5 mL 1 PBS 12.5 mL
goat serum (2.5% final) and 1 mL 0.5 M EDTA (1 mM final))
0.2 μm sterile filtered and stored at 4 C up to 3 weeks.
9. EpiLife or BGJb (Fitton-Jackson Modification), supplemented
with Human Corneal Growth Supplement (HCGS), 5 ng/mL
bFGF (FGF2), 10 ng/mL EGF (see Note 5), and 10% FBS (for
better results use low-endotoxin embryonic stem cell tested). If
serum-free medium is preferable, use AlbuMAX I (Lipid-Rich
Bovine Serum Albumin, Invitrogen, Cat. No.: 11020) (0.1%
final concentration) instead of serum (see Note 6).
2.3 Equipment
and Consumables
1. Isoflurane vaporizer, supply gas (oxygen), flowmeter and
induction chamber.
2. Stereo microscope for animal surgery, dissection.
3. TC-treated culture dish (10 cm).
4. Polypropylene centrifuge tubes, sterile (50 mL).
5. Polypropylene centrifuge tubes, sterile (15 mL).
6. Cell strainers (70 μm).
7. Syringes with 20G 100 needles.
8. Sterile 5 mL FACS round-bottom tubes.
9. Tools for lacrimal gland dissection and mincing: Razor blades
and/or small scissors with bended ends, forceps.
10. Tissue culture laminar flow hood.
11. Shaking water bath.
12. Standard
temperature-controlled
centrifuge.
table-top
low-speed
13. Flow cytometer (FACS cell sorter equipped with appropriate
for cell separation lasers) (see Note 7).
14. Hausser™ Bright-Line™ Phase Hemacytometer.
15. VWR® and VWR Signature™ Non-Bevel Pipet Tips
(Low-Binding Tips, Ultrafine Point).
16. Corning® cell strainer size 70 μm.
17. Sterile Eppendorf 2 mL round-bottom safe-lock microcentrifuge tubes.
2.4
Software
1. FlowJo flow cytometry software (Tree Star) or equivalent.
Isolation of Lacrimal Gland Epithelial Progenitors
173
Fig. 1 Isolation of mouse LG. (a) Sterilize mouse skin between the eye and ear with 70% ethanol; remove skin
covering lacrimal gland. (b) In adult mice the LG is located close to the ear and partially overlays the parotid
gland. (c) To dissect, pull LG anterior part gently using tweezers, and simultaneously use scissor tip to
dislodge the LG from the parotid gland and surrounding connective tissue. (d) When LG has been freed from
surrounding tissues, cut the posterior end with small curved scissors, and place into the Petri dish with HBSS
or other dissecting medium
3
Methods
3.1 Cell Isolation
Procedure (See Fig. 1)
1. Warm media in 37 C water bath.
2. Anesthetize mouse by isoflurane inhalation and sacrifice by
cervical dislocation.
3. Wash mouse skin between the eye and ear with 70% ethanol
and dry with clean cloth.
4. Carefully remove skin between the eye and ear covering lacrimal gland (Fig. 1A, B).
5. On a surgical bench or using the stereo microscope, harvest the
lacrimal glands; place glands in a 3.5 cm dish with 2 mL cold
PBS on ice (one mouse at a time). Note: To dissect a LG, pull
gently by the LG anterior part using tweezers, and at the same
time, use the sharp tip of small scissors to dislodge the LG from
the parotid gland and surrounding connective tissue (blunt
dissection) (Fig. 1C, D). Avoid cutting the LG free with scissors as the LG and parotid salivary glands are located very close
to each other and need to be separated by blunt dissection
before LG dissection.
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Helen P. Makarenkova and Robyn Meech
6. The LG is covered by a connective tissue capsule/envelope.
Remove LG capsule with two fine forceps under the
microscope.
7. Optional step: Place whole LGs in trypsin-pancreatin solution
for 10–20 min on wet ice (see Note 8). After this treatment
wash away trypsin-pancreatin solution with the dissection
medium (MEM, supplemented with GlutaMAX, FBS or trypsin inhibitor).
8. Important: In order to prevent the cells from sticking to the
bottom of the dish or incubation tubes, pretreat dishes and
tubes with a 1–4% BSA solution (see Note 9).
9. Transfer all LG into a 3.5 cm dish with 0.5 mL pre-warmed
media I, and mince into very small pieces using scissors (see
Notes 10 and 11).
10. Once all LG are minced, transfer all pieces and media from
Petri dish into a 2 mL centrifuge tube.
11. Add 500 μL collagenase stock solution and 12 μL of 1 M
calcium chloride (final concentration 6 mM), DNase I
2.5 μg/mL (5 units/mL), and 80 μL of dispase II stock
solution, and adjust volume with medium I to 2 mL (see
Note 12).
12. Mix and place tube on a shaking water bath, warmed to 37 C,
at 100–120 rpm for 90 min. Alternatively, if a shaking water
bath is not available, place tube in a 37 C incubator, and mix
contents manually by inverting the tube every 15–20 min.
However, the latter is likely to reduce cell yield due to suboptimal digestion.
13. Each 30 min triturate by slowly pipetting gland pieces up and
down 10–20 times using 1000 μL pipette tips with decreasing
bore size (cutting a 1000 μL pipette tip with sterile blades to
make the bore size larger).
14. After each 30 min incubation/trituration, take a 10 μL aliquot
and examine under the microscope. If the majority of cells are
still in clusters or attached to LG pieces, continue digestion/
trituration until a mainly single cell suspension is obtained (see
Note 13).
15. Optional: At the end of 90-min digestion, pass sample 5–10
times through a 10 mL syringe with 20 G needle to further
release cells into suspension and disrupt clumps. No visible
lacrimal gland pieces should remain in solution once digestion
is completed (see Note 10).
16. Transfer cell suspension into a 15 mL tube, and add up to 5 mL
of blocking medium, containing 100 μg/mL (or 200 units/
mL) DNase I, 5 mM MgCl2, and 10% FBS in HBSS. Slowly
invert the tube up and down 2–3 times to mix.
Isolation of Lacrimal Gland Epithelial Progenitors
175
17. Incubate cells for 15–30 min in the blocking media at room
temperature.
18. Pass cell suspension through a 70-μm cell strainer atop a sterile
50 mL centrifuge tube; wash the strainer with 1 mL of blocking media. Repeat this filtration step at least one more time to
remove any cell clumps.
19. Centrifuge samples at 1200–1500 rpm (300–400 g) for
5 min at RT.
20. Aspirate supernatant and resuspend the cells in 2 mL of cold
HBSS containing 5 mM MgCl2. Transfer cell suspension into a
2 mL microcentrifuge tube.
21. Centrifuge the sample at 1000–1500 rpm for 3 min at RT. Optional: Repeat the steps 20 and 21 to wash cells prior to the
next step (see Note 14).
22. Aspirate supernatant and resuspend cells in 1 mL of Accutase
solution (see Note 15).
23. Incubate the sample at 37 C, at 100–120 rpm in shaking water
bath for 2–3 min.
24. Resuspend cells in 1 mL of blocking medium.
25. Transfer cell suspension into 50 mL tube and add 20–25 mL of
blocking medium (HBSS containing 10% FBS, 1 mM EDTA).
26. Centrifuge sample at 1000–1500 rpm for 5 min. Optional:
Wash cells with blocking medium one more time to remove
any residual Accutase.
27. Discard the supernatant and resuspend cells in 2 mL of FACS
buffer containing DNAse I (at 8U/ml final concentration,
should be added prior to use) and transfer cells into 2 mL
Eppendorf tubes. Incubate cells at RT for 15–20 min.
28. Meanwhile, check number of cells under the microscope.
Count cells using hemacytometer, calculate number of cells
per 1 mL solution.
29. Adjust cell concentration to 250,000–500,000 cells/mL in
FACS buffer; if necessary concentrate by centrifuging the sample at 1500 rpm for 3 min at RT and resuspending cells in a
smaller volume of FACS buffer to achieve the minimum cell
concentration.
30. Aliquot cells into 400 μL volumes (100,000–200,000 cells/
400 μL) in 2 mL Eppendorf tubes, and stain with appropriate
antibodies (see step 31) for 20–40 min at 4 C in the dark. Do
not vortex the samples as it can damage your cells, mix by
pipetting.
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Helen P. Makarenkova and Robyn Meech
31. Single
color
controls:
For
each
sample
use
100,000–200,000 cells/400 μL FACS buffer with the following antibodies/dyes.
(a) No antibody (unstained control).
(b) 0.5 μL FxCycle Violet stain (DAPI).
(c) 4 μL FITC Rat anti-Mouse CD34.
(d) 1 μL Alexa Fluor 700 CD45 Rat Anti-Mouse Antibody.
(e) 2 μL APC-eFluor 780 CD117 (c-kit).
(f) 1 μL PE Rat Anti-Mouse Ly-6A/E (Sca1).
(g) 5 μL APC Rat Anti-Mouse CD326 (EpCAM).
32. Add all of the desired antibodies together to the samples that
will be FACS-sorted (called here ‘after sort samples’). If more
than 200,000 cells are used, a proportional adjustment of
antibody concentration is required (see Note 16).
33. Dilute sixfold with chilled FACS buffer (i.e., add 2 mL buffer
to 400 μL single color controls and 2–10 mL buffer to sort
samples depending on their volume).
34. Centrifuge samples at 1200 rpm for 5 min at 4 C.
35. Optional: Wash cells with FACS buffer one more time.
36. Aspirate supernatant, resuspend sort samples in 1–2 mL FACS
buffer or single color controls in 500 μL FACS buffer containing DNAse I (8U/mL), transfer to 5 mL FACS tubes, and add
2 μL FxCycle Violet stain (DAPI) to each sample (see Note
17).
37. The main population of live cells is determined by forward and
side scatter area gating, as well as dead cell exclusion via DAPI,
propidium iodide (PI), or 7-aminoactinomycin (7AAD), and
should be low on side scatter and low to medium on forward
scatter. Doublet exclusion is done via determining forward
scatter area vs. width and also side scatter area vs. width. Positive staining for fluorescent markers in stained samples is compared to unstained controls. The appropriate cell population
for sorting is gated based on the CD34, CD45, CD117,
Ly-6A/E, and CD326 marker profile (see Notes 18 and 19).
3.2 Reaggregated
Progenitor Cell
Cultures Using FACSIsolated EPCP
1. Prepare the culture media and warm it in a 37 C water bath.
2. Place 3–5 mL of medium per well in 6-well plate and fill the
empty wells and the inter-well space with sterile PBS. Keep
plates in the tissue culture incubator.
3. Centrifuge isolated EPCP in 1.5 mL Eppendorf tubes at
1200–1500 rpm (300–400 g) for 5 min at 4 C.
4. Carefully remove the supernatant, make sure that pellet is not
disturbed.
Isolation of Lacrimal Gland Epithelial Progenitors
177
Fig. 2 Preparation of progenitor cell reaggregated 3D cultures. FACS-isolated EPCP is counted, and approximately 1 105–2 105 cells are centrifuged and then resuspended in 20–50 μL of growth medium and
drawn into a 100–200 μL pipette tip. Pipette tip is carefully sealed with sterilized parafilm. Cells are
centrifuged at 1500 g for 10 min to form a plug or reaggregate which is inoculated (using tungsten needle
to help push cells into gel) into a 15 μL drop of Laminin I gel or Matrigel sitting on a polycarbonate filter. The
filters are then placed, gel-side up, on top of culture medium. These floating cultures could be maintained for
several days and later used for analysis or preparation of monolayer cultures
5. Gently resuspend pellet in the growth medium (use 20–50 μL
of medium per pellet) (Fig. 2).
6. Pipet cell suspension into each sterile 200 μL non-beveled tip
and seal tip with ethanol-sterilized parafilm (make sure the tip is
well embedded into the parafilm to avoid leaks).
7. Transfer tip into 15 mL tube and seal the tube.
8. Centrifuge at 1500 rpm (400 G), 5 min, at RT.
9. When reaggregates are ready for inoculation, place the Sterlitech hydrophilic polycarbonate membrane filter (Sterlitech
Corporation; catalog number, PCT00513100) into the
empty 3.5 mm culture dish.
10. Place a 15 μL drop of 3D Culture Matrix Laminin I gel diluted
1:1 with culture desired media (3 mg/mL final concentration)
or Corning® Matrigel® Basement Membrane Matrix, diluted
with desired culture medium 1:2–1:3 in the middle of the filter
(see Note 20).
11. Take tip out or tube and carefully remove parafilm.
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Helen P. Makarenkova and Robyn Meech
12. Inoculate reaggregate into a 15 μL drop of matrix gel, and then
place culture into a tissue culture incubator (37 C, 5%CO2) to
polymerize the gel.
13. Carefully place the Sterlitech filter with the gel-embedded
reaggregate on the top of the culture medium gel-side up.
14. Remove any bubbles underneath the filter carefully making
sure that the upper part of membrane is not immersed in
medium.
15. Culture for 3–7 days or until ready for analysis (Fig. 2).
4
Notes
1. All protocols were approved by the Scripps Research Animal
Care and Use Committee.
2. Pancreatin does not dissolve very well. All undissolved material
should be removed by filtering.
3. Units per milligram vary by lot and calculations need to be
adjusted accordingly.
4. Do not freeze/thaw more than one time.
5. Corneal supplement also contains a small amount of hydrocortisone. Additional EGF (corneal supplement contains only
1 ng/mL of EGF) and bFGF improve cell survival and
proliferation.
6. Lipid-Rich AlbuMAX I helps to stabilize cellular membrane
(especially important for epithelial cells).
7. Color combinations can be adjusted to match the laser combinations available.
8. The treatment with trypsin-pancreatin assists the penetration
of collagenase at the next step but is not essential. For LGs
isolated from 3 to 6 mice (6–12 glands), 2 mL trypsin/pancreatin solution is enough.
9. BSA solution could be reused several times.
10. During the dissection and mincing steps, look for and
discard large pieces of white fat surrounding the LG, these
will not be efficiently digested and may clog the syringe and
needle at step 15.
11. Scissors or razor-based mincing into small pieces prior to enzymatic digestion is a critical step in the protocol; mincing into
large pieces will result in an incomplete digestion of the tissue.
12. DNAse is included to digest DNA that has leaked into the
dissociation medium as a result of cell damage and will cause
the medium to become viscous and trap released cells.
Isolation of Lacrimal Gland Epithelial Progenitors
179
13. Cell suspensions following tissue digestion should be kept on
ice or at 4 C and labeled and sorted as quickly as possible
following sample preparation.
14. Washing will prevent any inhibition of Accutase by residual
serum.
15. Accutase is a marine-origin enzyme with proteolytic and collagenolytic activity that performs exceptionally well in detaching/dissociating cells for later analysis of cell surface markers.
16. It is not recommended to stain more than 1,000,000 cells/
400 μL. If a larger number of cells will be sorted, perform
staining in 15 mL tubes, and calculate the volume of antibodies
required based on volume of sample and number of cells.
17. FxCycle Violet stain (DAPI) can be used at 300–900 μg/mL
before increased autofluorescence is observed. Alternatively,
propidium iodide can be used to discriminate between live
and dead cells, but fluorescence compensation is more difficult
with this color scheme.
18. Cellular yield from one adult mouse, C57BL/6 strain, 1–2 months of age should be approximately 150,000–200,000
events (live cells) by FACS analysis.
19. The FACS gating and analysis for putative epithelial cell progenitors isolation has been described in detail in Gromova
et al., 2017 [4]. Briefly, we have used CD34 (hematopoietic
and mesenchymal progenitor cell antigen, vascular endothelial
cells) and CD45 (hematopoietic cells and various lymphocytes)
to exclude mesenchymal, hematopoietic, and endothelial cells.
Sca-1 and c-kit are expressed in multiple stem cell types,
whereas EpCAM labels epithelial cells. In the LG, two c-kitpositive cell populations could be detected: c-kitbright (i.e., high
level of expression) and c-kitdim (i.e., low level of expression)
[4]. All cells in the c-kitbright populations are EpCAMneg but
CD45pos. Thus c-kitbright cells most likely represent hematopoietic progenitors, lymphocytic and mast cells (note: granular
mast cells could be detected by immunostaining with c-kit
antibody). c-kitdim cells can be further parsed into three cell
populations: c-kitdim/EpCAMpos/CD45neg (22%, or 2.5–3.0%
of total cells), c-kitdim/EpCAMneg/CD45pos (70% or 8% of
total cells), and very small c-kitdim/EpCAMneg/CD45neg
(<1% of total cells) [4]. Sca-1 and CD34 were not detected
in the epithelial c-kitdim/EpCAMpos/CD45neg cell population.
Thus, the c-kitdim/EpCAMpos/CD45neg/Sca-1neg/CD34neg
cells are the putative epithelial progenitor cell (EPCP) population. Once isolated, EPCP could be maintained in monolayer
cultures or in reaggregated cultures or used for
transplantation.
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Helen P. Makarenkova and Robyn Meech
20. It is important to keep Laminin I and/or Matrigel on ice to
avoid gelation. Gel dilutions also need to be tested for their
ability to support cell growth prior to use for EPCP.
Acknowledgments
This protocol was adapted and modified from previous work published by both Helen Makarenkova and Robyn Meech and their
respective research groups [4]. This work was supported by
National Institutes of Health/National Eye Institute (NIH/NEI;
Bethesda, MD, USA) Grants 1R01EY026202 (to HPM).
References
1. Joe AW, Gregory-Evans K (2010) Mesenchymal
stem cells and potential applications in treating
ocular disease. Curr Eye Res 35:941–952
2. Sivan PP, Syed S, Mok PL, Higuchi A, Murugan
K et al (2016) Stem cell therapy for treatment of
ocular disorders. Stem Cells Int 2016:8304879
3. Aluri HS, Samizadeh M, Edman MC, Hawley
DR, Armaos HL et al (2017) Delivery of bone
marrow-derived mesenchymal stem cells
improves tear production in a mouse model of
Sjogren’s
syndrome.
Stem
Cells
Int
2017:3134543
4. Gromova A, Voronov DA, Yoshida M,
Thotakura S, Meech R et al (2017) Lacrimal
gland repair using progenitor cells. Stem Cells
Transl Med 6:88–98
5. Tiwari S, Ali MJ, Vemuganti GK (2014) Human
lacrimal gland regeneration: perspectives and
review of literature. Saudi J Ophthalmol
28:12–18
6. Bittencourt MK, Barros MA, Martins JF, Vasconcellos JP, Morais BP et al (2016) Allogeneic
mesenchymal stem cell transplantation in dogs
with keratoconjunctivitis sicca. Cell Med
8:63–77
7. Villatoro AJ, Fernandez V, Claros S, Rico-Llanos
GA, Becerra J et al (2015) Use of adiposederived mesenchymal stem cells in keratoconjunctivitis sicca in a canine model. Biomed Res
Int 2015:527926
8. Ackermann P, Hetz S, Dieckow J, Schicht M,
Richter A et al (2015) Isolation and investigation of presumptive murine lacrimal gland stem
cells. Invest Ophthalmol Vis Sci 56:4350–4363
Chapter 13
Organotypic Culture of Adult Mouse Retina
Brigitte Müller
Abstract
Retinal explant culture systems have the potential to mimic the functional dynamics of the organ beyond
those of the dissociated cells, thus making this technique a very powerful intermediate model system
between in vitro cell cultures and in vivo animal models. The different retinal layers made of highly
specialized cell types remain intact, while glia cell reactions and/or intercellular interactions can be
evaluated under well-defined conditions in the lab.
In retinal disorders neurodegeneration of mature retinal cells takes place. Therefore, we investigated the
adult murine neuroretina in organ culture to test its suitability for use in preclinical therapeutic applications.
Here we describe a method for the organ culture of adult murine retina (>12 weeks) used to establish
survival, cellular changes and early degeneration patterns of neuronal and glial cells. After enucleation of the
eyeball and careful dissection of the retina from the sclera and retinal pigment epithelium, the detached
retina is cultured with photoreceptor facing down on a supporting track-etched polycarbonate membrane
in a 6-well culture plate maintained in a humidified atmosphere of 5% CO2 and 95% air at 37 C. After 1, 2,
3, 4, 6, 8, or 10 days retinal explants can be harvested and immediately processed for RNA isolation or fixed
in paraformaldehyde for histological analysis.
Key words Retinal explant, Organotypic culture, Animal models, Photoreceptors, Apoptosis, Gliosis,
Retinal detachment, Retinal degeneration
1
Introduction
Retinal explant culture systems have the potential to mimic the
functional dynamics of the organ beyond those of the dissociated
cells, thus making this technique a very powerful intermediate
model system between in vitro cell cultures and in vivo animal
models. The different retinal layers made of highly specialized cell
types remain intact, while glia cell reactions and/or intercellular
interactions can be evaluated under well-defined conditions in the
laboratory.
The organotypic culture of the neonatal mouse retina has been
very useful for improving the knowledge of both normal retinal
development and retinal degeneration and especially to define the
role of various factors in photoreceptor degeneration and retinal
Ivan Bertoncello (ed.), Mouse Cell Culture: Methods and Protocols, Methods in Molecular Biology, vol. 1940,
https://doi.org/10.1007/978-1-4939-9086-3_13, © Springer Science+Business Media, LLC, part of Springer Nature 2019
181
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Brigitte Müller
cell fate determination and development [1–4]. It is now a widely
used tool, with broad applications in the field of ophthalmology.
Many alterations observed during in vitro retina culturing [1, 3–6]
resemble some characteristics of experimental retinal detachment
and diabetic retinopathy in vivo, respectively [7, 8]. Others studied
the relationship between retinal development, maturation, degeneration, and gene transfer in culture [9–12]. Furthermore, many
studies evaluated the therapeutic effect and potential toxicity of
substances [13, 14].
Several methods have been described for culturing retinal
explants from different species [3, 5, 15–17]. The method of
Caffé and colleagues [3], in which the neonatal mouse retina is
placed with the photoreceptor layer facing downward on rafts made
of nitrocellulose filters and polyamide gauze grids, has been used in
variations in several studies [9, 18].
Neonatal retinal organotypic cultures differ from adult ones
since the immature retinal neurons go through phases of differentiation and pruning, under the control of various growth factors
during the first 3 weeks postnatally [1, 19–24]. Additionally, neonatal organotypic retinal cultures have the fundamental problem
that outer and inner segments of photoreceptors do not develop
correctly [1]. The first study of adult rat and murine retinal explants
in culture reported reasonable viability of mouse retinas for at least
4 days in culture [25]. In this study, murine retinal explant cultures
were investigated after particle-mediated acute gene transfer. In a
more recent murine study, it was verified that organotypic cultures
from developing retinas show a higher rate of cell viability and
better preservation of the normal cytoarchitecture in comparison
to those obtained from adult retinas [26]. Since in retinal disorders
neurodegeneration takes place in mature retinal cells, our current
interest was to keep the adult murine neuroretina in organ culture
as long as possible to assess its viability and rate its possible use in
preclinical therapeutic applications [27]. For that we isolated the
retina from the sclera and retinal pigment epithelium (RPE) and
transferred it on the culture insert the photoreceptor layer facing
the supporting membrane. Even with different approaches for isolating the retina, it was not possible to keep the RPE attached to the
retina. We assume that an intact RPE would have helped to reduce
the loss of photoreceptors and therefore have kept the retinal
explants in culture viable for longer than 10 days [27–29]. Nevertheless, even without RPE the adult retinal explant showed GFP
signaling in some photoreceptors as well as in Müller cells after
6 days in culture due to transduction with AAV vectors at the day of
retinal dissection (Fig. 7 in [30]).
In conclusion, the adult murine organotypic retina culture can
be used for gene transfer and gene editing as an intermediate step
between cell culture and animal experiments even though the
viability of the retina decreases continuously during culture.
Organotypic Retina Culture
183
Fig. 1 (a) from left to right: micro scissors, spring type, for cutting open the eyecup; fine straight tweezers
(DUMONT No. 5, extra fine tip); glass transfer pipette with a pipette sucker at the wide end. (b) from left to
right: Curved forceps (Jeweler #7) for grabbing the eyeball underneath before enucleation; Petri dish (30 mm
diameter) for retina dissection; curved scissors (iris scissors, 11 cm) for cutting the optic nerve during
enucleation. (c) Inserts with track-etched polycarbonate membrane (TEPC), pore size 0.4 μm and 30 mm in
diameter, in a six-well plate used for cultivation of retinal explants. (d) Mouse eyeball in a Petri dish held by
fine tweezers from the left and pierced with an injection needle (30 Gauge, yellow). (e) Magnified setup of (d).
Injection needle piercing the eyeball (coming from the top). (f) Eyeball held down at the connecting tissue.
Micro scissor (coming from the top) is cutting open the eyeball at the level of the ora serrata. (g) Eyecup
consisting of the retina and sclera with the anterior part (lens and cornea) removed. (h) The sclera and retina
after placing four cuts through the complete eyecup at 3, 6, 9, and 12 o’clock position to get the shamrock
shape. The retina is halfway peeled off the sclera. (i) The sclera (left) and retina (right) after complete
separation. (j) Isolated retina after transfer onto the TEPC membrane of the insert. Ganglion cell side is facing
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Brigitte Müller
Further work needs to be done to inhibit apoptosis and promote
the survival of photoreceptor cells by varying and supplementing
culture conditions with various neurotrophic factors [31]. Additionally, efforts can be made to co-culture murine RPE cells
obtained by primary cell culture.
2
Materials
2.1 Dissecting
Instruments,
Containers, and
Culture Inserts
1. Fine straight forceps (DUMONT No. 5, extra fine tip) and
micro scissors, spring type, for cutting open the eyecup
(Fig. 1a).
2. Curved forceps (Jeweler #7) and curved scissors (iris scissors,
11 cm) for enucleation (Fig. 1b, see Note 1).
3. Injection needle: 30 gauge (yellow 0.3 13 mm).
4. Transfer pipette: Cut off the narrow part of a glass Pasteur
pipette and melt the edges in the flame of a Bunsen burner to
get rid of any sharp edges. Use a pipette sucker at the wide end
(Fig. 1a).
5. 200 μL and 1000 μL laboratory pipettes with their respective
sterile filter tips.
6. Petri dishes 30 mm in diameter were used for dissection of the
retina (Fig. 1b).
7. Inserts with track-etched polycarbonate membrane (TEPC),
pore size 0.4 μm and 30 mm in diameter, were used for cultivation of retinal explants (Fig. 1c, k).
8. Six-well plates (see Note 2).
9. Petri dish (100 mm in diameter) for swaps soaked with ethanol.
10. 70% ethanol in a lab spray bottle.
11. Plenty of sterile swaps.
12. pH test strips.
13. Surgical scalpel (blade no 10).
2.2 Buffers and
Culture Media
1. 10 Hanks’ Balanced Saline Solution (HBSS) without Phenol
Red (commercial): Add 5 mL HBSS (10) to 45 mL double
distilled water (deionized water) treated by autoclave to get 1
HBSS. Adjust pH with NaOH (see Notes 3 and 4).
ä
Fig. 1 (continued) up. Blunt ends of the tweezers flatten the curled retinal parts. (k) Culture insert with retinal
explant (arrow) sitting in a well of the six-well plate. Basic culture medium (of pink color) was added to the
well. (l) Retinal explant after 6 days in culture, ready to be harvested. Dark spots represent pigment granules
attaching to the vitreous body on top of the ganglion cell layer
Organotypic Retina Culture
185
2. To formulate basic culture medium, use plain DMEM (Dulbecco’s Modified Eagle Medium): Add 0.5 mL penicillin/
streptomycin (100 μg/mL streptomycin and 100 units/mL
penicillin) and 0.5 mL 200 mM L-glutamine to 49 mL
DMEM (plain). Saturate with 5% CO2 and 95% air in the
incubator overnight (see Note 5).
3. Complete culture medium: Add 50% of basic DMEM, 25%
fetal bovine serum (FBS), and 25% 1 HBSS, pH 7.4.
2.3 Specialist
Equipment
1. Dissection microscope
2. Incubator with humidified atmosphere of 5% CO2 and 95% air
at 37 C
3. Laminar flow
3
Methods
It is not necessary to do the retina dissection under a laminar flow if
all work space is disinfected and a surgical mask is worn while
handling retinal explants. Disinfect hands and arms. Wear gloves
and change gloves after animal handling. Use a different part of the
work bench for enucleation and retina dissection to minimize
contamination via animal hair, fur, etc.
3.1
Post Enucleation
1. Disinfect both eyeballs by quickly rolling them over a swab
soaked with 70% ethanol, which lies in a Petri dish (100 mm
diameter).
2. Transfer into a small Petri dish (30 mm diameter) filled with
warm 1 HBSS (see Note 6). Change once to wash off tissue
debris, fur, etc.
3. Transfer into a fresh Petri dish filled with warm basic CO2saturated basic culture medium (see Note 7). If you want to
keep left and right eye separately, use individual dishes per eye.
3.2
Retina Dissection
Use new injection needle with every animal. Clean all dissection
tools after dissecting both the eyes of an animal using a sterile swab
with 70% ethanol.
1. Transfer the eyeball into the top of a sterile Petri dish filled with
a large drop (1000 μL) of warm basic CO2-saturated culture
medium (see Notes 8 and 9).
2. Grab the eyeball with the fine tweezers (Fig. 1a) at the connecting tissue at the back of the eyeball to hold it down. Use an
injection needle (30 gauge, yellow) to pierce a small hole into
the eyeball at the ora serrata (Fig. 1d, e).
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Brigitte Müller
3. Keep holding down the eyeball at the connecting tissue (see
Note 10), insert one bracket of the micro scissors (Fig. 1a) into
the little hole, and cut open the eyeball at the level of the ora
serrata (Fig. 1f) (see Notes 11 and 12).
4. Carefully remove anterior part of the eye.
5. Grab the remaining eyecup (Fig. 1g) at the back with fine
tweezers at the connecting tissue or optic nerve, and place
four cuts through the complete eyecup, i.e., cut the sclera and
retina at 3, 6, 9, and 12 o’clock position to get the shamrock
shape (Fig. 1h, i) (see Note 13).
6. Separate the retina from the pigment epithelium and the sclera
by clipping two corners of the sclera with the fine tweezers
(Fig. 1a) and tearing them carefully apart toward the optic
nerve head. The retina will peel off the back of the eyecup.
The remaining connection with the sclera is at the optic verve
head (Fig. 1h).
7. Use the micro scissors to carefully cut the optic nerve head.
8. After that, tear scleral corners to isolate the retina from the
sclera completely (Fig. 1i).
3.3 Transfer of
Retinal Explants onto
the TEPC Membrane
1. Use 200 μL pipette to add a 200 μL drop of warm basic culture
medium onto the TEPC membrane insert sitting in a six-well
plate (Fig. 1c, k) (see Note 14).
2. Use glass transfer pipette (Fig. 1a) to transfer isolated retina
onto the membrane photoreceptor side down (see Note 15).
3. Carefully remove all basic culture medium around the retinal
explant with the 200 μL pipette to allow flattening of the retinal
tissue. Avoid touching the membrane.
4. Carefully flatten the retina on the membrane by using the blunt
ends of the tweezers in a flat angle. Eventually, the retina lies
flat on the membrane (Fig. 1j) (see Note 16).
5. Add 1000 μL warm basic culture medium into the six well and
transfer the six-well plate into the incubator (humidified atmosphere of 5% CO2 and 95% air at 37 C) (Figs. 1k and 2).
Culture medium is only applied into the space between the six
well and the culture insert (see Fig. 2).
6. Repeat all steps in Subheadings 3.1 and 3.2 for the remaining
eyecups.
7. Exchange basic culture medium to complete culture medium
after the retinal explants on the TEPC membrane have been
2–3 h in the incubator.
Organotypic Retina Culture
187
Fig. 2 Schematic of shamrock shaped retinal explant laying flat on the TEPC
membrane of the insert. The insert is sitting in a well which is filled with 1000 μL
culture medium
3.4 Culturing and
Harvesting of Retinal
Explants
1. Cultivate retinal explants for 1, 2, 3, 4, 6, 8, or 10 days in the
incubator (humidified atmosphere of 5% CO2 and 95% air at
37 C).
2. Exchange complete culture medium every second day by
removing the used medium with the 1000 μL pipette and
replacing it with 1100 μL fresh warm complete medium (see
Note 17).
3. For harvesting retinal explants have warm 1 HBSS, transfer
pipette and forceps ready.
4. Transfer insert with retinal explant into an empty well of the
six-well plate. Add 1000 μL warm 1 HBSS into the well.
5. Carefully add 1000 μL warm 1 HBSS into the insert to
separate the explant from the membrane (see Note 18)
(Fig. 1l).
6. Retinal explants intended for histological analysis should be
immersed immediately in 4% paraformaldehyde in phosphatebuffered saline (PBS) at room temperature for 45–70 min.
7. Retinal explants intended for isolation of total mRNA should
be transferred into RLT buffer including 2-mercaptoethanol
(10 μL/1 mL RLT buffer), homogenized, and quickly frozen
in liquid nitrogen (see Notes 19 and 20).
4
Notes
1. All dissecting instruments must be clean and disinfected by the
autoclave.
2. All used plastic ware should be sterile.
3. All media should be prepared under the laminar flow to keep
them sterile. Hence, they can be used at several occasions after
opening them once. Dividing the culture media in 15 mL
Falcon tubes prevents pH changes due to oxygen. Store all
media in the fridge.
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Brigitte Müller
4. After diluting the 10 HBSS to 1 HBSS with sterile deionized water, the pH should be checked and adjusted to 7.4 by
adding 1 M NaOH dropwise. Mix by agitating the Falcon tube.
Use pH test strips.
5. Basic medium containing DMEM supplemented with 2 mM Lglutamine, 5.75 mg/mL glucose, and antibiotics (100 μg/mL
streptomycin and 100 units/mL penicillin) should be put in
the incubator for at least 12 h (overnight) to saturate with 5%
CO2 and 95% air. During retina dissection basic medium
should be exchanged regularly as soon as the color turns pink.
6. Keep all 1 HBSS and basic DMEM at 37 C during retina
dissection.
7. Basic culture medium is lacking fetal bovine serum. It should
be used for dissection of the retina and placing the retinal
explant on the membrane of the insert. This is important to
allow the retina to attach to the membrane. Complete medium
prevents proper attachment of the retina onto the membrane.
Therefore, complete medium containing 25% fetal bovine
serum should be added to retinal explants on the TEPC membrane the first time after they have been 2–3 h in the incubator.
8. Use the top part of the 30 mm Petri dish to open the eyeball
and to dissect the retina from the eyecup.
9. Use a dissection microscope for retina dissection.
10. After punctuation of the eyeball, keep holding the eyeball at
the connecting tissue and change instruments with the
other hand.
11. While cutting open the eyeball, keep turning it accordingly to
allow the micro scissors to cut easily without denting or
squeezing the eyeball.
12. Make sure to cut the eyeball open at the level posterior the ora
serrata (toward the posterior part of the eye) to separate the
retinal tissue easily from the pigment epithelium afterward.
13. For a shamrock-like shape of the retina and sclera, perform four
cuts halfway between ora serrata and optic nerve head in
length. This is important to flatten the retina on the membrane
later.
14. Cultivation of retinal explants on hydrophobic polycarbonate
membranes gave the best result. Avoid polyethylene membranes, since the retinal tissue is hard to remove from the
membrane after the cultivation period. In our experience, the
polyethylene has negative consequences for the viability and
tissue preservation.
Organotypic Retina Culture
189
15. The photoreceptor layer of the retinal explants should be facing
the supporting track-etched polycarbonate membrane
(TEPC). The membrane has a pore size of 0.4 μm and is
30 mm in diameter to fit six-well plates. If the ganglion cell
layer is facing the membrane after the initial transfer suck the
retinal explant back into the transfer pipette and try again by
putting it back onto the membrane.
16. Carefully uncurl the retina by using the blunt ends of the
tweezers in a flat angle stroking the retinal surface. Avoid
piercing or squeezing the retinal tissue.
17. Medium change during culture period of retinal explants
should be performed under sterile conditions under a laminar
flow. Culture medium should be brought to 37 C before use.
Culture medium is only applied into the space between the six
well and the culture insert (see Fig. 2). Avoid putting complete
medium directly on top of the retinal explant. Avoid filling the
six well with more than 1100 μL medium. Otherwise, the
retinal explant will detach from the membrane and float,
which has negative consequences on the viability and tissue
preservation.
18. While harvesting the retinal explants, it might not be possible
to separate the tissue from the TEPC membrane by gently
washing with warm 1 HBSS. If so, cut the TEPC membrane
off the plastic insert part using a scalpel. Transfer membrane
with retinal explant on top to a Petri dish (100 mm in diameter) filled with warm 1 HBSS. Hold on to the edge of the
membrane with forceps and carefully slide the scalpel between
the tissue and the membrane. If the retinal explant tissue is
used for histological purposes, transfer the membrane with
retinal explant on top into the fixative directly, before separating the tissue. The physical separation can be done after the
fixation.
19. Three retinal explants (10–15 mg) of one culture period were
pooled per mRNA isolation.
20. RNeasy® Micro Kit (#74004, QIAGEN, Hilden, Germany)
was used to isolate mRNA according to manufacturer’s
instructions from freshly harvested retinal explants.
Acknowledgments
Funded by ERC starting grant 311244. I thank Franziska Wagner
and Wilhelm Reihnhard for excellent assistance with taking the
photographs during retinal explant dissection.
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Brigitte Müller
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1. Ogilvie JM, Speck JD, Lett JM et al (1999) A
reliable method for organ culture of neonatal
mouse retina with long-term survival. J Neurosci Methods 87:57–65
2. Caffé AR, Söderpalm A, Holmqvist I et al
(2001) A combination of CNTF and BDNF
rescues rd photoreceptors but changes rod differentiation in the presence of RPE in retinal
explants. Invest Ophthalmol Vis Sci
42:275–282
3. Caffé AR, Visser H, Jansen HG et al (1989)
Histotypic differentiation of neonatal mouse
retina in organ culture. Curr Eye Res
8:1083–1092
4. Caffé AR, Söderpalm A, van Veen T (1993)
Photoreceptor-specific protein expression of
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28. Strauss O (2005) The retinal pigment epithelium in visual function. Physiol Rev 85:845–881
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Chapter 14
Langendorff-Free Isolation and Propagation of Adult
Mouse Cardiomyocytes
Matthew Ackers-Johnson and Roger S. Foo
Abstract
Isolation of healthy, intact cardiomyocytes from the adult mouse heart for cardiac research is challenging.
Traditional protocols depend upon specialized Langendorff apparatus, which requires extensive training
and presents significant technical and logistical barriers. Described here is a much simplified technique,
introducing optimized dissociation buffers to the heart by direct needle injection into the left ventricle,
allowing deep myocardial perfusion and the isolation of high yields of viable, rod-shaped cardiomyocytes,
using only standard surgical and laboratory equipment. This method also permits the concurrent isolation
of cardiac non-myocyte cellular populations.
Key words Cardiomyocyte, Cardiac myocyte, Cardiac fibroblast, Cell isolation, Langendorff, Cell
culture, Mouse heart
1
Introduction
The isolation of high-quality, viable myocytes from myocardial
tissue is an essential prerequisite for molecular and cellular investigation of cardiac function and pathology. Cardiomyocytes in the
intact adult myocardium exist in close association with neighboring
cells and extracellular matrix and are highly sensitive to mechanical
perturbations, enzymatic damage, hypoxia, nutrient bioavailability,
pH, and ionic fluctuations. Simple cutting or mincing of heart
tissue with subsequent enzymatic digestion produces poor yields
of healthy adult myocytes, particularly in rodents, wherein physiologically high intracellular sodium predisposes cardiomyocytes to
calcium overload [1]. Traditional protocols for isolation of cardiomyocytes from adult rodent hearts rely on retrograde aortic perfusion using specialized Langendorff apparatus, which enables deep
myocardial infiltration of enzymatic dissociation buffers via the
coronary vasculature [2, 3]. However, this poses considerable logistical and technical barriers to researchers and demands extensive
training investment.
Ivan Bertoncello (ed.), Mouse Cell Culture: Methods and Protocols, Methods in Molecular Biology, vol. 1940,
https://doi.org/10.1007/978-1-4939-9086-3_14, © Springer Science+Business Media, LLC, part of Springer Nature 2019
193
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Matthew Ackers-Johnson and Roger S. Foo
Fig. 1 Schematic overview of injection method. The emerging aorta is clamped,
and dissociation buffers are injected into the base of the left ventricle. Buffers
are forced through the coronary circulation (arrows), enabling deep perfusion of
myocardial tissue. RA right atrium; LA left atrium; RV right ventricle; LV left
ventricle
Detailed here is a simplified alternative technique for the isolation of high yields of healthy, calcium-tolerant cardiomyocytes from
the adult mouse heart, using only standard surgical tools and
equipment [4, 5]. The procedure utilizes a series of optimized
dissociation buffers, which are introduced ex vivo by direct intraventricular injection. Deep myocardial perfusion via the coronary
vasculature is induced by clamping of the emerging aorta (Fig. 1).
Isolated cardiomyocytes are then purified by sequential gravity
settling steps, with the option of gradual calcium reintroduction
to produce calcium-tolerant cells for functional studies or culture.
Isolated cardiomyocytes can be harvested immediately; applied
directly to functional calcium, electrophysiological, or imaging
studies; or cultured for extended periods to allow in vitro manipulations such as adenoviral gene transfer. Furthermore, this technique permits the concurrent isolation, culture, and co-culture of
non-myocyte resident cardiac populations, from the same regions,
in the same adult mouse heart.
2
Materials
Surgical instruments, skin forceps (RS-5248), blunt-end scissors
(RS-5965), curved-end forceps (RS-5137), sharp-end scissors
(RS-5840), Reynolds full-curved hemostatic forceps (RS-7211),
and straight-end forceps (RS-5070), were purchased from Roboz,
USA.
Simplified Isolation of Adult Mouse Cardiomyocytes
2.1 Cardiomyocyte
Isolation
195
1. EDTA buffer: 130 mM NaCl, 5 mM KCl, 0.5 mM NaH2PO4,
10 mM HEPES, 10 mM glucose, 10 mM 2,3-butanedione
monoxime (BDM), 10 mM taurine, 5 mM EDTA. Dissolve
directly in 1 L ultrapure 18.2 MΩ.cm H2O and adjust to
pH 7.8 using NaOH. Sterile filter, store at 4 C for up to
2 weeks, and keep sterile.
2. Perfusion buffer: 130 mM NaCl, 5 mM KCl, 0.5 mM
NaH2PO4, 10 mM HEPES, 10 mM glucose, 10 mM BDM,
10 mM taurine, 1 mM MgCl2. Dissolve directly in 1 L ultrapure 18.2 MΩ cm H2O and adjust to pH 7.8 using NaOH.
Sterile filter, store at 4 C for up to 2 weeks, and keep sterile.
3. Collagenase buffer: 0.5 mg/mL collagenase 2, 0.5 mg/mL
collagenase 4, 0.05 mg/mL protease XIV (Sigma P5147).
Dissolve in perfusion buffer; make fresh immediately before
isolation (see Note 1).
4. Stop buffer: Stop buffer is made with perfusion buffer containing 5% sterile fetal bovine serum (FBS). Make fresh on day of
isolation.
5. Tyrode’s solution: 137 mM NaCl, 4 mM KCl, 10 mM HEPES,
1 mM MgCl2, 0.33 mM NaH2PO4, 1.2 mM CaCl2, 5.5 mM
glucose. Dissolve directly in 1 L ultrapure 18.2 MΩ cm H2O
and adjust to pH 7.4 using NaOH. Sterile filter, store at 4 C
for up to 2 weeks, and keep sterile.
2.2 Cardiomyocyte
Culture
1. Laminin solution: Dilute murine laminin (Thermo Scientific,
23017-15) in sterile phosphate-buffered saline (PBS) to final
concentration 5 μg/mL; mix well. Keep sterile; use
immediately.
2. Plating media: M199 medium with Earle’s salts and
l-glutamine, supplemented with 5% FBS, 10 mM BDM (see
Note 2), and 100 units/mL penicillin with 100 μg/mL streptomycin (P/S; optional). Keep sterile; store at 4 C for up to
2 weeks.
3. Culture media: M199 medium with Earle’s salts and
l-glutamine, supplemented with 10 mM BDM (see Note 2),
P/S (optional), 0.1% w/v bovine serum albumin (BSA) (see
Note 3), 1 insulin-transferrin-selenium (ITS) supplement,
1 chemically defined (CD) lipid supplement (Thermo Scientific 11905-031) (see Note 4). Keep sterile, protect from light,
and store at 4 C for up to 2 weeks.
4. Calcium reintroduction buffers: These are three buffers with
increasing calcium concentrations, made by mixing culture
media and perfusion buffer in increasing proportions: buffer
1 (1:3), buffer 2 (1:1), and buffer 3 (3:1). If performing
immediate calcium or electrophysiological experiments rather
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Matthew Ackers-Johnson and Roger S. Foo
than culturing cells, use Tyrode’s solution instead of culture
media.
5. Fibroblast growth media: DMEM/F12 1:1 media (l-glutamine
included) supplemented with 10% FBS and P/S.
3
Methods
Important: National and institutional guidelines and regulations
must be consulted and adhered to before commencement of all
animal work. All buffers and procedures are at room temperature
unless otherwise specified. See Table 1 for troubleshooting.
3.1 Pre-coating
of Culture Surfaces
Only if cardiomyocytes are to be subsequently plated and/or
cultured. Tissue culture surfaces are pre-coated with laminin solution for at least 1 h at 37 C or overnight at 4 C (see Note 5).
3.2 Preparation
of Buffers and Media
Media and buffers are prepared as detailed in Subheading 2. Enzymatic digestion can be carried out at room temperature but is more
efficient at 37 C, in which case collagenase buffer is warmed
immediately before use in a clean water bath or equivalent. Isolation of one heart requires roughly 30 mL EDTA buffer, 20 mL
perfusion buffer, up to 60 mL collagenase buffer (or less if recycling; see Note 6), and 10 mL stop buffer.
3.3 Preparation
of Equipment
and Surgical Area
1. Surgical area and instruments (1 skin forceps, 1 blunt-end
scissors, 1 round-end forceps, 1 sharp-end scissors, 1
Reynolds forceps (hemostatic clamp) or equivalent, 1 sharpend forceps) are sterilized with 70% ethanol.
2. EDTA, perfusion, and collagenase buffers are aliquoted into
2, 1, and 5 10 mL sterile syringes, respectively, and sterile
27 G hypodermic needles are attached. 10 mL syringes are
selected largely due to ease of handling; other sizes may be
used if preferred. For 37 C digestion, collagenase syringes are
kept warm in a clean water bath or equivalent.
3. 60 mm sterile petri dishes are prepared containing 1 10 mL
EDTA buffer, 1 10 mL perfusion buffer, 1 10 mL collagenase buffer, and 1 3 mL collagenase buffers. Isoflurane anesthetic system apparatus is set up, with connections to a
ventilation chamber and a nose-cone ventilator, which is positioned centrally on the surgery area (see Note 7).
4. Mice are anesthetized in the chamber with 100% O2 at 0.5 L/
min flow rate, containing isoflurane (atomizer dial at 4%, scale
1–5%). Once unconscious, mice are transferred to the surgery
area, with anesthesia maintained using the nose cone.
Simplified Isolation of Adult Mouse Cardiomyocytes
3.4 Surgical
Procedure
197
1. Full anesthesia is confirmed by reduced breathing rate and lack
of toe-pinch reflex response.
2. EDTA buffer and perfusion buffer syringes are prepared by
removal of needle caps. Ensure that no bubbles exist in the
syringes or needles.
3. The mouse chest is wiped generously with 70% ethanol and
opened using skin forceps and blunt-end scissors just below the
diaphragm, which is then opened to expose the heart (Fig. 2a).
4. Using the round-end forceps, the left lung may be moved aside
to reveal the descending aorta and inferior vena cava. Both are
cut using the sharp-end scissors, at which point 7 mL EDTA
buffer is injected steadily within around 1 minute into the right
ventricle (RV), which can be identified by its darker color. To
flush out as much blood as possible, the needle should enter at
Table 1
Troubleshooting
Problem
Possible cause
Poor digestion, heart does
not soften
Choice of euthanasia other
than oxygen-isoflurane
anesthesia technique
Use oxygen-isoflurane. If this is
unavailable, heparin pre-administration
may be necessary. Try to reduce time
between animal death and flushing of
the heart with EDTA buffer
Old/degraded enzymes
Purchase/prepare new enzymes
New enzyme batch with low Optimize enzyme concentration
activity
Bubbles in syringe
Ensure removal of bubbles before
injection. Bring buffers to correct
temperature before filling syringes
Incomplete clearance of
Increase volume and time for EDTA
blood from heart
buffer injection to RV if necessary
Old/fibrotic heart
Increase digestion time, use 37 C if not
already, increase enzyme concentration
Complete digestion, heart
softens, but low yield of
viable rod-shaped cells
Old/contaminated buffers/
reagents
Impure water
Incorrect buffer preparation
Overdigestion
Good yield, but cells die
while in stop buffer
Old/contaminated FBS
Impure water
Solution
Prepare new buffers, filter sterilize.
Purchase new reagents, particularly
BDM or taurine
Use only ultrapure
> ¼ 18.2 MΩ cm H2O
Check preparation, remake buffers.
Calibrate pH meter and check pH
Reduce digestion time/enzyme
concentration
Use new FBS. Try new batch if still
unsuccessful
Use only ultrapure 18.2 MΩ cm H2O
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Matthew Ackers-Johnson and Roger S. Foo
Fig. 2 Schematic illustrations of in situ heart flushing, removal, and ex vivo
injection. (a) Chest cavity of anesthetized mouse is opened to below the
diaphragm, which is then cut through to expose the heart. The descending
aorta and inferior vena cava are cut (1), and the heart is immediately flushed
with EDTA buffer by injection into the right ventricle (2). Reynolds hemostatic
forceps then reach around the heart to clamp the emerging aorta (3), and hold
the heart while it is removed by cutting around the forceps (4). (b) The excised
heart, still held by the clamp, is transferred to 60 mm dishes for subsequent
injection and digestion steps
the base of the RV, penetrating no more than a few mm, and
the angle of entry may be carefully varied during injection.
3.5
Removal of Heart
1. The emerging aorta is then clamped. Any hemostatic clamp will
suffice, but full-curved-ended Reynolds forceps are preferred.
These can easily reach around the heart and clamp the
emerging aorta in situ, which does not require high precision,
and inclusion of additional emerging vessels does not matter,
although clamping of atrial appendages should be avoided.
2. The heart is removed by simply cutting around the outside of
the forceps and transferred, still held by the clamped forceps, to
the 60 mm dish containing EDTA buffer, where it should be
almost completely submerged (Fig. 2b).
3.6
Heart Digestion
1. Locate the left ventricle (LV), which is the larger of the ventricles and forms a pointed apex at the base of the heart. Using
the second EDTA syringe, insert the needle into the base of the
LV wall, 2 or 3 mm above the apex, penetrating no more than a
few mm into the LV chamber, and inject the EDTA buffer
Simplified Isolation of Adult Mouse Cardiomyocytes
199
starting at a flow rate of around 1 mL per 2 or 3 min (see Note
8).
2. After 6 min or application of all 10 mL EDTA buffer, whichever is first, the needle is removed, and the heart is transferred,
still held by the clamped forceps, to the dish of perfusion
buffer. 3 mL perfusion buffer is then similarly injected into
the LV, if possible via the same perforation left by the previous
injection. Inexperienced users may find a magnification lens
beneficial for identification of the original injection point.
3. After 2 min or application of all 3 mL perfusion buffer, whichever is first, the heart is transferred to the dish containing
10 mL collagenase buffer, and the LV is injected sequentially
with the five syringes of collagenase buffer (see Notes 6 and 9).
4. The clamp is removed, and scissors may be used to separate the
heart into its constituent chambers, or other specific regions, as
desired. The selected region is then transferred to the final
3 mL dish of collagenase buffer (multiple dishes can be used
here in order to isolate cells from multiple regions).
5. Tissue is gently teased apart into roughly 1 mm 1 mm sized
pieces using the round and sharp-end forceps, which requires
very little force following a successful digestion.
6. Dissociation is completed by gentle trituration for 2 min using
a 1 mL pipette, with a wide-bore tip (purchased or homemade
using sterile scissors) to reduce shear stress.
7. To stop the enzymatic digestion, 5 mL of stop solution is added
to the cell-tissue suspension, which may be gently pipetted for a
further 2 min, and inspected under a microscope (see Note 10).
8. Cell suspension is then transferred to a 50 mL centrifuge tube,
which should be stored on its side at room temperature to
reduce clumping and hypoxic damage. Cells may be stored
with little loss of viability for up to 2 h, in which time further
isolations may be performed. However, such delays may not be
suitable for sensitive applications.
3.7 Collection
of Cardiomyocytes by
Gravity Settling
If cells are to be cultured, subsequent steps are best undertaken in a
laminar flow cabinet, to maintain sterility.
1. Cell suspension is passed through a 100 μm pore-size strainer,
in order to remove undigested tissue debris. The filter is
washed through with a further 5 mL stop buffer.
2. Total volume of cell suspension is now typically around 15 mL.
This can be divided into two 15 mL centrifuge tubes, and cells
are then allowed to settle by gravity for 20 min. Most myocytes
will settle to a pellet, while most non-myocytes and cellular/
extracellular debris remain in suspension (see Note 11).
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Matthew Ackers-Johnson and Roger S. Foo
3. Supernatant is removed. If cells are to be harvested immediately without further in vitro experiments, myocyte fractions
are purified simply by three further rounds of sequential gravity
settling for 10 min in 4 mL fresh perfusion buffer, retaining the
myocyte-containing pellet each time.
3.8 Calcium
Reintroduction
and Culture of Cells
Where myocytes are to be returned to physiological extracellular
calcium levels and/or plated, it is important to do so in gradual
increments, in order to avoid spontaneous contraction and achieve
healthy populations of calcium-tolerant cells, which may then be
subjected to a wide range of experimental applications [4]. This can
be easily incorporated into the gravity settling steps.
1. Similar to Subheading 3.7, step 3, myocyte pellets are instead
resuspended sequentially in three calcium reintroduction buffers, containing increasing proportions of either Tyrode’s solution (for immediate calcium handling or electrophysiology
experiments) or culture media (for plating and/or culturing
of cells); see Subheading 2.2, item 3.
2. If required, the supernatant fractions, which contain
non-myocyte cell populations as well as rounded myocytes
and some viable myocytes, may be collected and combined
from each round of gravity settling. Plating and fibroblast
media can be warmed and equilibrated in a 37 C, 5% CO2,
humidified tissue culture incubator during this process.
3. For plating of cardiomyocytes, laminin solution is aspirated
from the prepared culture surfaces (see Subheading 3.1),
which are then washed once with PBS.
4. The final cardiomyocyte pellet is resuspended in
pre-equilibrated plating medium, and cells are plated at
application-specific densities: Typically around 25,000 cells/
mL, or 5000 cells/cm2, but this may be substantially lowered
for imaging studies.
5. Cardiomyocytes are transferred to the tissue culture incubator
and shaken gently in a side-to-side (not swirling) motion to
ensure even distribution. Adhesion of rod-shaped myocytes
occurs rapidly, within 20 min for most cells. Culture medium
may be pre-equilibrated in the incubator during this time.
6. Cells in the combined supernatant fraction may be collected by
centrifugation at 300 g for 5 min, resuspended in
pre-equilibrated fibroblast growth media, plated on tissue culture surfaces (area ~20 cm2 per LV), and transferred to the
culture incubator (see Note 12).
7. After 1 h, plated cardiomyocytes are gently washed once with
pre-equilibrated culture media and then incubated in culture
media, for the required experimental duration. Rounded
Simplified Isolation of Adult Mouse Cardiomyocytes
201
Fig. 3 Representative example of adult mouse cardiomyocytes after poor isolation procedure (a), containing
many rounded, hypercontracted, and dying cells, and good quality procedure (b), showing a majority of
healthy, rod-shaped cells. Healthy cardiomyocytes were plated and visualized at 40 (c) and 400 (d)
magnification, whereby characteristic angular morphology and sarcomeric striations are clearly visible. Scale
bars are 100 μm
myocytes do not adhere strongly and are removed by this
process (see Note 13).
8. Culture and fibroblast media are changed after 24 h and every
48 h in culture thereafter.
A successful isolation procedure yields up to one million cardiomyocytes per left ventricle, with 80% viable, healthy, rod-shaped
cells [4]. Poor isolations yield high numbers of dying, round,
hypercontracted cells, and troubleshooting is required; see Table 1
and Fig. 3a–d.
4
Notes
1. 100 collagenase and 1000 protease XIV (¼50 mg/mL)
stocks may alternatively be prepared in ultrapure
18.2 MΩ cm H2O, filter-sterilized, and stored in aliquots at
80 C for at least 4 months. These can be added to perfusion
buffer to produce collagenase buffer immediately before the
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Matthew Ackers-Johnson and Roger S. Foo
isolation. We use collagenases 2 (LS004176) and
4 (LS004188) from Worthington Biochemical, Lakewood,
USA, which exhibit high batch-to-batch reproducibility (collagenase 2, ~210 units/mg; collagenase 4, ~260 units/mg).
Collagenase 2 is a less pure extract with more basal clostripain
activity than collagenase 4, which can sometimes be advantageous, and in many cases, 2.5 mg/mL collagenase 2 alone is
sufficient at 37 C to attain good yields of myocytes. However,
using the described mixture as standard performs consistently.
2. 100 BDM (¼1 M) stocks can be prepared by dissolving
1.01 g BDM in 10 mL ultrapure 18.2 MΩ cm H2O, filtersterilized, and stored in aliquots at 20 C. Stock may require
incubation at 37 C to redissolve BDM before adding to media
as required. BDM is a myosin II ATPase inhibitor, used to
reduce myocyte contractions and improve the yield of isolated
cardiac myocytes. BDM must be removed from cultures before
conducting contractility, calcium handling, or electrophysiology experiments. It is normal to see a number of cardiomyocytes becoming hypercontracted and dying 1–2 h after BDM
removal. In culture media, blebbistatin can be used in place of
BDM, which may have fewer off-target effects, and improve
long-term survival and adenoviral transduction efficiency [6].
We find that 5 μM blebbistatin is optimal for this purpose.
3. 50 BSA (¼5% w/v) stocks can be prepared by dissolving 1 g
BSA in 20 mL PBS, filter-sterilized, and stored at 4 C for at
least 8 weeks, if kept sterile.
4. CD lipid mix is included to improve myocyte survival in longterm culture [4]. It is not typically required for short term
culture. The lipid mix is susceptible to oxidation and to light
damage and is therefore best stored in small aliquots containing
minimal air space, at 4 C, in the dark.
5. Laminin-coated surfaces are best prepared fresh but may be
sealed and stored at 4 C for up to 4 days. When using glass
surfaces, extra volume may be required for complete coverage.
Note that cells adhere less strongly to glass than plastic.
6. Following application of each syringe of collagenase buffer to
the heart, 10 mL will need to be removed from the dish, to
prevent overflow. To reduce consumption of enzyme, this
buffer may be collected and recycled for subsequent injection.
Care must be taken to prevent needle-prick injuries. However,
to prevent cellular cross-contamination, fresh collagenase
buffer is generally prepared for each heart.
7. Feedback from users suggests that the choice of euthanasia
technique is one of the most common causes of problems
encountered. Induction of rapid-onset anesthesia using
oxygen-isoflurane ventilation is strongly recommended. This
Simplified Isolation of Adult Mouse Cardiomyocytes
203
involves no injections and causes the mouse minimal stress.
Furthermore, circulation is intact, and blood is well oxygenated
up to the point of chest opening and introduction of EDTA
buffer. Injected anesthetics such as pentobarbital and ketamine
have a longer onset and significantly reduce respiration,
increasing the risk of ischemia and subsequent cardiomyocyte
calcium overload [1, 7]. Cervical dislocation carries the same
risk, in addition to likely blood coagulation and thus blockage
of coronary circulation in the time taken to open the chest and
inject EDTA buffer, particularly for inexperienced users, leading to poor myocardial perfusion of dissociation buffers and
low yields of healthy cardiomyocytes. If these techniques are
necessary, heparin administration is recommended 30 min
prior to euthanasia. It should be emphasized that euthanasia
by CO2 inhalation causes ischemia and is not appropriate for
myocyte isolation techniques.
8. The ideal flow rate when injecting buffers into the LV will vary
between hearts, but the best measure of adequate perfusion is
simply the minimum required to maintain full inflation of the
heart. Initially, very little force is required for the heart to
inflate, and flow rate may be only 1 mL per 2 or 3 min. As
digestion progresses and the heart softens, flow rate typically
reaches around 2 mL/min. A temptation is to over-apply,
which can cause buffer to force into and perforate the left atrial
appendage. This alone does not cause poor isolation results,
and the researcher may proceed as normal, although such
pressure is unnecessary. If desired, this protocol is compatible
with automated infusion pump setups [4].
9. The volume of collagenase buffer required for complete digestion varies between hearts. Small, young, healthy hearts can
digest in as little as 25 mL, while larger, older, or fibrotic hearts
may pass beyond 50 mL, necessitating the recycling of buffer
(see Note 6). Signs of complete digestion include a noticeable
reduction in resistance to injection pressure, loss of shape and
rigidity, holes and/or extensive pale and fluffy appearance at
the heart surface, and ejection of myocytes into the effluent
buffer, which are just visible to the naked eye. The point of
injection will often widen until significant buffer appears to be
flowing directly backwards, but the researcher may proceed as
necessary.
10. Myocytes may display contraction immediately after isolation
due to mechanical stimulation but should quickly acquiesce.
The presence of large numbers of rounded, hypercontracted
myocytes (Fig. 3a) indicates a poor isolation and requires
troubleshooting (see Table 1).
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Matthew Ackers-Johnson and Roger S. Foo
11. Sequential gravity settling is a method to obtain a highly pure
myocyte population and avoids damage caused by centrifugation. Viable rod-shaped myocytes tend also to settle faster than
round hypercontracted and dying myocytes, so enriching the
pellet for viable rod-shaped cells. Division of cell suspension
into two 15 mL centrifuge tubes rather than one 50 mL tube
aids the formation of a pellet due to the more steeply angled
base. Sterile polystyrene round-bottom tubes are also good
alternatives for this purpose.
12. Cardiac fibroblasts, and some other non-myocytes [4], adhere
to untreated tissue culture plastic surfaces within 1–2 h.
Remaining cardiomyocytes do not and can be washed off at
this stage to effectively purify the (mostly) cardiac fibroblast
population.
13. Cultured myocytes must be handled with great care. Avoid
shocks, vibrations, and rapid aspiration/introduction of
media. Always wash gently using warm culture media to reduce
ionic fluctuations, and change media one well at a time to avoid
prolonged exposure to air, particularly if culturing on glass
surfaces. When fixing cells with formaldehyde for imaging,
best results are obtained by adding 8% formaldehyde dissolved
in culture medium slowly to an equal volume of culture
medium already in the well and incubating for 15 min. Do
not swirl or shake.
References
1. Bers DM (2002) Cardiac Na/Ca exchange function in rabbit, mouse and man: what’s the difference? J Mol Cell Cardiol 34:369–373
2. Berry MN, Friend DS, Scheuer J (1970) Morphology and metabolism of intact muscle cells
isolated from adult rat heart. Circ Res
26:679–687
3. Powell T, Twist VW (1976) A rapid technique
for the isolation and purification of adult cardiac
muscle cells having respiratory control and a
tolerance to calcium. Biochem Biophys Res
Commun 72:327–333
4. Ackers-Johnson M, Li PY, Holmes AP, O’Brien
S-M, Pavlovic D, Foo RS (2016) A simplified,
Langendorff-free method for concomitant
isolation of viable cardiac myocytes and nonmyocytes from the adult mouse heart. Circ Res
119:909–920
5. Chen X, O’Connell TD, Xiang YK (2016) With
or without Langendorff: a new method for adult
myocyte isolation to be tested with time. Circ
Res 119:888–890
6. Kabaeva Z, Zhao M, Michele DE (2008) Blebbistatin extends culture life of adult mouse cardiac myocytes and allows efficient and stable
transgene expression. Am J Physiol Heart Circ
Physiol 294:H1667–H1674
7. O’Connell TD, Rodrigo MC, Simpson PC
(2007) Isolation and culture of adult mouse cardiac myocytes. Methods Mol Biol 357:271–296
Chapter 15
Isolation, Culture, and Characterization of Primary
Mouse Epidermal Keratinocytes
Ling-Juan Zhang
Abstract
Epidermis, the outermost layer of the skin, plays a critical role as both a physical and immunological barrier
protecting the internal tissues from external environmental insults, such as pathogenic bacteria, fungi,
viruses, UV irradiation, and water loss. Epidermal keratinocytes (KC), the predominant cell type in the skin
epidermis, are in the front line of skin defense. Here we describe methods to isolate and culture primary
epidermal KC from neonatal and adult mouse skin and describe in vitro assays to study and characterize KC
proliferation and differentiation and pro-inflammatory responses to viral products and UVB irradiation.
These methods will be useful for researchers in the field of epidermal biology to set up in vitro assays to
study the barrier and pro-inflammatory function of epidermal keratinocytes.
Key words Skin epidermis, Keratinocyte, Skin barrier, Keratinocyte proliferation, Keratinocyte differentiation, Pro-inflammatory response, dsRNA, UVB irradiation, TNF release
1
Introduction
Skin is the largest organ of the body, and epidermis is the outermost
layer of the skin. At the front line of defense, epidermis plays a
critical role in forming an intact barrier to protect the body from
dehydration and external insults, such as pathogenic bacteria, virus,
allergens, and UVB irradiation [1, 2]. This barrier function is
mainly provided by keratinocytes (KC), the predominant cell type
in the epidermis, and it is maintained by a tightly controlled balance
between proliferation and differentiation of KC [3, 4]. KC located
on the basal layer of the epidermis, including both epidermal stem
cells and transit-amplifying cells, are proliferative. As these basal
cells exit cell cycle, KC commit to terminal differentiation and
gradually move upward toward the surface of the skin [5, 6]. During
the differentiation process, KC undergo a series of biochemical and
morphological changes that result in the formation of distinct layers
of the epidermis. At the spinous layer, directly above the basal layer,
KC express early differentiation markers, such as keratin 10 (K10).
Ivan Bertoncello (ed.), Mouse Cell Culture: Methods and Protocols, Methods in Molecular Biology, vol. 1940,
https://doi.org/10.1007/978-1-4939-9086-3_15, © Springer Science+Business Media, LLC, part of Springer Nature 2019
205
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Ling-Juan Zhang
As KC migrate to the granular layer, these cells become
interconnected by tight junctions and express late differentiation
markers, such as filaggrin (FLG), loricrin (LOR), and involucrin
(INV). Eventually, when these cells reach the outer surface of the
epidermis, KC become terminally differentiated corneocytes, which
are enucleated and flattened and eventually sloughed into the environment as new cells replace them [1, 5, 6].
At the front line of host defense, epidermal KC are also an
important component of the skin’s innate immune system. In
response to PAMPs (pathogen-associated molecular patterns)
released by invading pathogens or DAMPs (damage-associated
molecular patterns) released by host cells during UVB irradiation
or wounding, KC produce a variety of pro-inflammatory cytokines
or chemokines, such as TNFα, IL6, IL8, CXCL10, and IFNβ [2, 7,
8]. These pro-inflammatory signals released from KC recruit or
activate myeloid and resident immune cells, mounting a rapid
host defense immune response leading to efficient pathogen clearance. However, uncontrolled inflammatory response may trigger
the development of auto-inflammatory skin diseases, such as psoriasis and rosacea [9, 10].
Here we describe methods to isolate epidermal KC from neonatal or adult mouse skin. This is an extended protocol modified from
our previous published protocol in Journal of Visualized Experiments
[10]. While neonatal KC are collected from both whole body skin
from neonates, adult KC are isolated from tail skin, which has thicker
epidermis and lower hair follicle density compared to body skin in
adult mice. Skin is first digested overnight with dispase, an enzyme to
dissociate the epidermis from dermis. The separated epidermal sheet
is then digested with a trypsin-like enzyme to release epidermal
KC. Isolated KC are seeded on culture dishes coated with extracellular matrix and cultured in low calcium medium supplemented with
defined growth supplements. Between days 2 and 5 after the initial
seeding, dead cells or differentiated cells are washed away by daily
medium changes; the remaining cells are proliferating and have
cobblestone morphology, a characteristic morphology of basal
KC. We also describe methods to characterize and study proliferation, growth factor starvation, and/or high calcium-induced differentiation as well as pro-inflammatory response of these primary KC
triggered upon exposure to viral product or UVB irradiation.
2
Materials
2.1
Animals
C57B/6 wild-type mice are bred and maintained in a specific
pathogen-free (SPF) environment according to animal facility regulations. Neonates are used within 2 days of birth (postnatal days
0~2), and adult mice are used between 6 and 15 weeks of age and
either female or male mice can be used.
Isolation and in Vitro Culture of Primary Mouse Keratinocytes
2.2 KC Isolation
and Culture
207
1. Sterile PBS, pH 7.4.
2. 10 cm Petri dish.
3. KC basal medium with 0.06 mM CaCl2.
4. Defined growth supplements include epidermal growth factor
(EGF), bovine transferrin, insulin-like growth factor1 (IGF1),
prostaglandin E2 (PGE2), bovine serum albumin (BSA), and
hydrocortisone (Life Technologies, Carlsbad, CA, Catalog
S0125).
5. Dispase.
6. Complete KC growth medium: Basal KC medium (0.06 mM
CaCl2) supplemented with defined KC growth supplement and
1 antibiotic-antimycotic.
7. Dispase digestion buffer: 4 mg/mL dispase in complete KC
growth medium.
8. Gelatin coating material.
9. Type 1 collagen coating material (either from bovine or rat tail
or recombinant human protein).
10. Trypsin-like enzyme/TrypLE (Life Technologies, Carlsbad,
CA, Catalog 12604-013).
11. 100 μm cell strainer.
2.3 Functional Cell
Assay
1. Culture dishes: 24-well clear flat bottom TC-treated cell culture plates are used for phase contrast imaging and/or
RTqPCR analyses. 96-well clear flat bottom TC-treated microplate is used for colorimetric cell counting assay. 8-well chamber slide is used for immunocytochemistry analysis.
2. Colorimetric cell counting kit: Colorimetric assays to measure
metabolic activity of living cells, such as Cell Counting Kit-8
(CCK-8) or MTS assay or MTT assay, can be used for
mouse KC.
3. 5-bromo-20 -deoxyuridine (BrdU) is dissolved as 20 mM stock
in DMSO in aliquots.
4. Rat anti-BrdU antibody.
5. High molecular weight (HMW) poly(I:C).
6. Corded handheld UV lamps.
7. 8-watt UV tubes.
8. Mouse TNF ELISA kit.
9. Light inverted microscope for cell culture.
10. Fluorescent microscope for cover slides.
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3
Ling-Juan Zhang
Methods
3.1 Overnight
Dispase Digestion
of Neonatal or
Adult Skin
1. Euthanize the postnatal day 0–day 2-old C57BL/6 wild-type
neonatal pups by decapitation using scissors, and follow steps
2 and 3 to set up dispase digestion. For adult mice, euthanize
adult mice according to animal facility regulations. Cut off the
tail from the base, and follow steps 4 and5 to set up dispase
digestion.
2. To peel neonatal skin off the body, first cut off limbs just above
the wrist and joints, and cut off the tail from the base leaving a
small hole. Insert sharp scissors through the hole from the tail
and cut the skin along the dorsal midline through the neck.
Next, use one forceps to gently lift a corner of skin off the neck
and the other forceps to grasp the exposed neck and body, and
then carefully peel the whole skin off the body and over the leg
stumps in one continuous motion (see Note 1).
3. Rinse the peeled neonatal skin in a 10 cm Petri dish with 15 mL
of sterile PBS, and then transfer the skin to a 2 mL tube
prefilled with 2 mL ice cold dispase digestion buffer (see Note
2). Digest the skin overnight at 4 C on a rotator in a refrigerator. Next day, proceed to Subheading 3.2.
4. To peel adult tail skin off the bone, first use a sharp blade to cut
through the tail base from the tail tip. Next, use one forceps to
gently lift a corner of the skin off the tail bone at the base and
the other forceps to grasp the exposed tail bone, and then
carefully peel tail skin off the bone with one continuous
motion. Cut each of the peeled skin into 2~3 pieces, each of
which is ~2 cm in length.
5. Rinse the peeled adult tail skin in a 10 cm Petri dish with 15 mL
of sterile PBS, and then transfer skin pieces from each tail to a
2 mL tube prefilled with 2 mL ice cold dispase digestion buffer
(see Note 3). Digest the skin overnight at 4 refrigerator on a
rotator. Next day, proceed to Subheading 3.2.
3.2 Isolation
of Keratinocytes from
Skin Epidermis
1. On the second day (within 12~18 h post dispase digestion),
carefully transfer the skins together with the dispase solution to
a Petri dish and then to a new Petri dish with 15 mL sterile PBS
to wash away excess dispase. Using two pairs of forceps, carefully transfer each skin piece to a dry Petri dish with epidermis
side down and dermis side up. Stretch the skin folds so that the
skin is fully extended on the Petri dish.
2. Before separating the epidermal sheet from the dermis, place a
drop of 500 μL TrypLE, a trypsin-like digestion solution
(room temperature) in a new Petri dish (see Note 4). To
remove the dermis, use one forceps to hold down a corner of
the epidermis, and use the other forceps to gently lift the
Isolation and in Vitro Culture of Primary Mouse Keratinocytes
209
dermis (pink, opaque, gooey) away from the epidermal sheet
(whitish, semitransparent; see Note 5). Dispose of the dermis as
biohazardous material.
3. Use two pairs of forceps to grasp the cross corner of the
separated epidermal sheet, and slowly transfer it onto the surface of the TrypLE digestion solution with the basal layer
downward (see Note 6).
4. Cover skin on Petri dish with lid, and incubate for 20 min at
room temperature on a horizontal shaker with gentle agitation.
Basal KC become loosely attached to the epidermal sheet or are
released from the epidermal sheet during this digestion
process.
5. To stop digestion, add 2 mL ice cold complete KC growth
medium per epidermis to the Petri dish. Using forceps vigorously rub the epidermal sheet, and the medium will become
turbid as KC are released into solution from the epidermal
sheet. Tilt the Petri dish to collect and transfer the cell suspension to a 50 mL centrifuge tube leaving the remaining epidermal sheet on the dish. Keep the collection tube on ice during
the procedure.
6. Repeat step 5 two more times, and combine the cell suspensions into the same 50 mL tube.
7. Pipet the cell suspension up and down gently a few times to
disperse cell clumps using a serological pipette, and then pass it
through a 100 μm filter to a new 50 mL centrifuge tube.
8. Centrifuge the filtered cells at 180 g for 5 min. Aspirate the
supernatant, and resuspend the cell pellet in 1 mL cold KC
growth medium, and determine the cell number using a hematocytometer (see Note 7).
3.3 Primary Mouse
KC Culture
1. Prior to cell seeding (see Note 8), culture dishes should be
coated with appropriate ECM materials (see Note 9) according
to the manufacturer’s instruction at 37 C for 30 min. Remove
the coating material completely immediately before adding the
cell suspension.
2. Seed the isolated neonatal KC at a density of 5 104/cm2, or
adult KC at a density of 10 104/cm2, in KC growth medium
in culture dishes coated with ECM material as described above.
3. Change the medium 24 h after the initial plating, and then
change medium daily to remove unattached cells or cells that
spontaneously differentiate and detach from culture dish.
Between day 2 and day 5 of initial plating, cells should reach
>70% confluency. Cells can then be used for experimentation.
Representative phase contrast images for adult mouse KC from
8 h to 4 days after the initial plating are shown in Fig. 1a.
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Ling-Juan Zhang
Fig. 1 In vitro assays to measure KC growth and proliferation. (a) Phase contrast
images at 10 magnification of primary adult mouse KC at 8 h, day 1, day 2, day
3, and day 4 post the initial seeding. Scale bar ¼ 100 μm. (b) The relative cell
number of primary adult mouse KC at indicated day after the initial seeding was
measured by the CCK-8 cell viability assay. All error bars indicate mean s.e.m.
*P < 0.05, **P < 0.01, ***P < 0.001 (ANOVA). (c) Subconfluent neonatal KC
were pulse-labeled with BrdU prior to immunocytochemistry analyses using
anti-BrdU antibody (red), and nuclei were counterstained with DAPI in blue.
Scale bar ¼ 100 μm
3.4 In Vitro Assays
to Study KC
Proliferation (See Note
10)
1. Measurement of cell proliferation by colorimetric cell counting
assay: Primary KC are seeded in 96-well flat bottom clear plate,
and KC growth medium is changed daily during the assay
(100 μL medium /well). To measure relative cell number,
10 μL of CCK-8 solution is added to each well and incubated
for 1 h in a cell culture incubator (5% CO2 at 37 C), and then
O.D. at 450 nm is measured by a spectrometer. Measurement
of relative cell number over a time course of 4 days by CCK-8
assay is shown in Fig. 1b.
2. Labeling of S-phase cells by BrdU incorporation: Primary KC are
grown on coverslips and incubated for 30 min with 10 μM
BrdU, followed by fixation in 4% PFA/PBS. Fixed cells are
treated with 0.2 M HCl for 30 min at room temperature
followed by neutralization with a borate buffer. Cells are then
permeabilized with 0.1% Triton X-100 and subjected to standard immunocytochemistry procedures using a rat anti-BrdU
antibody and a Cy3-conjugated anti-rat secondary antibody.
Nuclei are counterstained with DAPI. Representative image for
BrdU labeling of primary neonatal KC in the S-phase of cell
cycle is shown in Fig. 1c using fluorescence microscope.
Isolation and in Vitro Culture of Primary Mouse Keratinocytes
211
Fig. 2 Growth factor starvation and/or High calcium triggered KC differentiation.
(a) Primary neonatal KC were cultured in growth medium (first lane), KC basal
medium without growth factors (lanes 2 and 3), or growth medium with high
calcium for 24 or 48 h as indicated. The expression of early differentiation
marker K10 was measured by RTqPCR analysis. Fold induction of K10 compared
to control cells (lane 1) was shown, and Hprt was used as housekeeping gene in
the analysis. All error bars indicate mean s.e.m. ***P < 0.001,
****P < 0.0001 (ANOVA). (b) Phase contrast images at 10 magnification of
primary adult mouse KC treated with 0.2 mM CaCl2 at indicated time. Scale
bar ¼ 100 μm
3.5 In Vitro Assays
to Study KC
Differentiation (See
Note 11)
1. Early differentiation by growth factor starvation: Growth factor depletion alone is more efficient than high calcium to
induce the expression of KC early differentiation markers,
such as K10 (see Note 12). To starve the cells, remove growth
medium and replace with basal medium without added growth
supplements. K10 is induced (15-fold) as early as 24 h of GF
removal, and this induction further increases by 48 h (~
300-fold), whereas high calcium only leads to ~ten-fold of
K10 induction by 48 h (Fig. 2a).
2. To trigger terminal differentiation by high calcium, proliferating KC are first starved overnight in basal medium without
added growth supplements as described above (see Note 13).
Next day, add CaCl2 to 0.2 mM in culture medium. As shown
in Fig. 1a, b, within 8~12 h after high calcium switch, cells
become flattened, and the distinct intercellular space becomes
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Ling-Juan Zhang
less apparent; by 24 h the cell-cell adhesion with tight junction
becomes apparent, and the formation of corneocytes/cornified
envelop and vertical cell stratification is observed around
48~72 h post high calcium switch.
3. In vitro assays to study KC differentiation: Cells can be harvested at desired time points for RTqPCR and/or western blot
analyses to determine the mRNA or protein expression of KC
differentiation markers, such as early differentiation marker
K10 or late differentiation markers such as FLG, INV, and/or
LOR [3].
3.6 In Vitro Assays
to Study
Pro-inflammatory
Response of KC to Viral
Products
3.7 UVB IrradiationMediated Cell Death
and Secretion of TNFa
from KC
1. Culture the mouse KC in KC growth medium until cells reach
~80% confluency. Starve the cells for 6~16 h in basal medium
without added growth supplement (see Note 14).
2. Add 1 μg/mL poly(I:C), the synthetic viral dsRNA, directly to
the culture medium of the cells (see Note 15). Cells can be
harvested 4~24 h posttreatment for RTqPCR analysis of the
expression of pro-inflammatory cytokines, such as Ifnb1 as
shown in Fig. 3a, or ELISA to measure cytokine secretion to
conditioned medium.
1. Culture the mouse KC in KC growth medium until cells reach
desired confluency. Immediately prior to UVB irradiation,
remove growth medium and replace with sterile PBS warmed
to 37 C.
2. Treat cells with 25 mJ/cm2 UVB using handheld UVB lamps.
After UVB irradiation, change cells back to fresh growth
medium. Representative images of cells treated with UVB for
12 h and 24 h are shown in Fig. 3b.
3. To measure and quantify cell viability, cells grown in 96-well
flat bottom plate are first treated with UVB as described above,
and then treated cells are subjected to CCK-8 cell assay at
12 and 24 h post-UVB treatment as described in Subheading
3.4, step 2.
4. To measure TNFα secretion (see Note 16), cells grown in
24-well flat bottom plate are first treated with UVB as
described above; conditioned medium is then collected from
UVB-treated cells at desired time point. The amount of TNFα
in the conditioned medium is measured by the mouse TNFα
ELISA kit following the manufacturer’s instructions. As shown
in Fig. 3c, TNFα was abundantly secreted from UVB-treated
adult KC compared to untreated control cells.
Isolation and in Vitro Culture of Primary Mouse Keratinocytes
213
Fig. 3 Pro-inflammatory response of primary mouse KC to viral dsRNA or UVB
irradiation. (a) Primary adult KC were treated with 1 μg/mL poly(I:C) or vehicle
control for 4 h, and cells were subjected to RTqPCR analysis. Fold induction of
Ifnb1 in poly(I:C)-treated cells compared to control cells was shown, and Hprt
was used as housekeeping gene in the analysis. (b, c) Primary adult mouse KC
were grown to confluency and then exposed to 25 mJ/cm2 UVB irradiation. (b)
Phase contrast images at 10 magnification 12 h or 24 h after UVB irradiation
compared to untreated control cells. Scale bar, 200 μm. (c) Secretion of TNFα
was measured by ELISA in conditioned medium from control cells or cells treated
with UVB for 24 h. All error bars indicate mean s.e.m. ***P < 0.001 (ANOVA)
4
Notes
1. Peel the skin off the whole body slowly as one piece, and be
careful not to break skin into pieces as this will result in cell loss
during the dispase digestion step.
2. Make sure the skin is extended and not folded in the tube to
allow efficient digestion of the whole skin.
3. Tail skin pieces from mice with the same genetic background
can be combined and incubated in one 15 mL tube (up to five
tails per tube) filled with 15 mL dispase solution.
4. Compared to trypsin, the TrypLE digestion solution is gentler
on cells and can be inactivated by dilution alone without the
need for trypsin inhibitors, such as FBS. Each 10 cm Petri dish
should fit up to five drops of the TrypLE solution.
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Ling-Juan Zhang
5. Because the epidermal sheet is very fragile, the dermis should
be lifted away from the epidermis slowly to prevent tearing of
the epidermal sheet.
6. Use forceps to carefully stretch and unfold the epidermal sheet
so that it is fully extended and floats on the digestion solution.
7. KC spontaneously differentiate in suspension, so prior to cell
seeding, the cell suspension should be kept on ice and plated
onto ECM-coated culture dishes as soon as possible (preferably
within 1 h).
8. Coating of culture dishes with the appropriate extracellular
matrix (ECM) should be done immediately after the cell
count and as soon as possible prior to cell seeding.
9. A gelatin-based coating matrix works well for neonatal KC,
whereas a collagen-based coating matrix is preferable for adult
KC due to the decreased ability of adult cells to adhere compared to their neonatal counterparts. Either rat tail, bovine, or
recombinant human type 1 collagen can be used here.
10. Cell proliferation can be measured by either a colorimetric cell
counting assay using a dye that measures the metabolic activity
from living cells or 5-bromo-20 -deoxyuridine (BrdU) incorporation assay to measure BrdU incorporated into the newly
synthesized DNA during cell proliferation.
11. Calcium is considered the most physiological agent to trigger
epidermal KC differentiation in vitro and in vivo in a similar
manner. In vitro, low calcium (0.02 mM~0.1 mM) maintains
the proliferation of basal KC as a monolayer, whereas high
calcium (>0.2 mM) rapidly triggers a terminal differentiation
process converting KC from basal cell morphology to stratified
corneocyte morphology.
12. We show there that growth factor starvation is more efficient
than high calcium to induce genes that are associated with KC
early differentiation, such as K10 (Fig. 2a). While high calcium
weakly induces the expression of early differentiation genes, it
strongly induces the expression of KC late differentiation process, such as FLG, INV, and LOR [3]. These observations are
in line with the in vivo observation that calcium concentration
is actually low in both basal and the spinous layer (in which K10
is expressed) but rises in the granular layer (where late differentiation markers express) [11–13]. Together these evidences
suggest that calcium is unlikely the key factor that drives basal
cells to commit to early differentiation process. Instead cell
cycle arrest (which can be triggered by growth factor starvation
in vitro) is likely the key factor that drives basal cells to commit
to early differentiation stage.
Isolation and in Vitro Culture of Primary Mouse Keratinocytes
215
13. The growth factor removal step may enhance but is not
required for the high calcium triggered cellular changes associated with the late differentiation processes, including tight
junction formation and vertical cell stratification.
14. We always include a growth factor starvation step so that cells
are synchronized and more consistent results can be obtained.
Starvation can also lower basal inflammatory signal. Medium
change should be done at least 6 h prior to treatment as
medium change alone induces stress and inflammatory
response from the cells.
15. Poly (I:C) is diluted and added in a small volume (10 μL)
directly to culture well to minimize disturbance to the cells.
16. TNFα is an important pro-inflammatory cytokine that is
induced by UVB irradiation, and it drives KC apoptosis following UVB irradiation [14].
References
1. Fuchs E, Raghavan S (2002) Getting under the
skin of epidermal morphogenesis. Nat Rev
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2. Bernard JJ, Cowing-Zitron C, Nakatsuji T,
Muehleisen B, Muto J et al (2012) Ultraviolet
radiation damages self noncoding RNA and is
detected by TLR3. Nat Med 18:1286–1290
3. Zhang LJ, Bhattacharya S, Leid M, GanguliIndra G, Indra AK (2012) Ctip2 is a dynamic
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4. Sambandam SAT, Kasetti RB, Xue L, Dean
DC, Lu Q, Li Q (2015) 14-3-3sigma regulates
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Kilkenny AE, Roop DR (1988) Signal transduction for proliferation and differentiation in
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8. Zhang LJ, Sen GL, Ward NL, Johnston A,
Chun K et al (2016) Antimicrobial peptide
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serine protease activity and cathelicidin promotes skin inflammation in rosacea. Nat Med
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10. Li FW, Adase CA, Zhang LJ (2017) Isolation
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from neonatal and adult mouse skin. J Vis
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11. Menon GK, Grayson S, Elias PM (1985) Ionic
calcium reservoirs in mammalian epidermis:
ultrastructural localization by ion-capture cytochemistry. J Invest Dermatol 84:508–512
12. Elias PM, Nau P, Hanley K, Cullander C,
Crumrine D et al (1998) Formation of the
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Chapter 16
Isolation and Propagation of Mammary Epithelial Stem
and Progenitor Cells
Julie M. Sheridan and Jane E. Visvader
Abstract
Several methods of mammary gland dissociation have been described that utilize a combined strategy of
mechanical and enzymatic dissociation to isolate mammary epithelial cells (MECs) from intact tissue
(Smalley et al., J Mammary Gland Biol Neoplasia 17:91–97, 2012). Here we detail a robust method that
enables the isolation of all major stem and progenitor MEC populations, which has been successfully used
to study stem cell behavior when coupled with transplantation and in vitro assays (Shackleton et al., Nature
439:84–88, 2006; Bouras et al., Cell Stem Cell 3:429–441, 2008; Sheridan et al., BMC Cancer 15:221,
2015; Jamieson et al., Development 144:1065–1071, 2017). Furthermore, we outline two prominent
methods for culturing MECs for the purposes of ex vivo manipulation or study: 2D feeder layer cultures and
3D Matrigel colony assays. Importantly, all outlined methods retain stem and progenitor cell behaviors and
can be used in combination with downstream in vivo, in vitro, or in silico analyses.
Key words Mammary gland, Epithelial stem cell, Progenitor cell, Single cell suspension, Stem cell
culture
1
Introduction
A large body of evidence suggests that a stem cell-based mammary
epithelial differentiation hierarchy establishes the mammary gland
and maintains function through rounds of differentiation and
regression that accompany pregnancy and weaning [1]. Several
mammary epithelial stem cell (MaSC) and progenitor cell populations have been isolated including transplantable bipotent MaSCs
that demonstrate luminal and basal/myoepithelial differentiation
capacity as well as a range of more restricted cell types that contribute to limited luminal or basal cell subtypes [2–6]. The characterization of these populations has provided significant insights into
the processes that guide normal mammary development and
homeostasis and tumorigenesis [12]. Beyond phenotyping, the
isolation of MaSC and progenitor cell populations provides a valuable tool with which to study or manipulate mammary epithelial
Ivan Bertoncello (ed.), Mouse Cell Culture: Methods and Protocols, Methods in Molecular Biology, vol. 1940,
https://doi.org/10.1007/978-1-4939-9086-3_16, © Springer Science+Business Media, LLC, part of Springer Nature 2019
217
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Julie M. Sheridan and Jane E. Visvader
cell behavior ex vivo, and robust purification and culture protocols
are key to these endeavors [7–10].
Herein we describe a MEC isolation protocol that reliably
isolates and maintains the viability of MaSC and progenitor cell
populations [2, 11]. Additionally, we outline two methods of
MaSC and progenitor cell culture: (1) a tractable 2D feeder layer
system that readily expands MaSC and progenitor cells ex vivo,
providing a means to chemically or genetically manipulate cells
prior to downstream analyses [2], and (2) a Matrigel-based assay
that is permissive for the differentiation of different stem/progenitor cells into morphologically distinguishable colony types, a feature that makes this system suitable for studies of cell behavior or
function [2].
2
Materials
Where possible, materials are prepared using aseptic technique and
solutions are filter sterilized with 0.2 μm filters.
2.1 Materials
for the Dissociation
of Mammary Gland
Tissue to a Single Cell
Suspension
1. Sterile Dulbecco’s phosphate buffered saline solution, without
calcium and magnesium (DPBS).
2. Sterile wash buffer: DPBS with 2% fetal calf serum (FCS).
3. Sterile MEC medium supplemented with 1% FCS (1% MEC
medium): DMEM/Ham’s F12 containing GlutaMAX, 5 μg/
mL insulin, 500 ng/mL hydrocortisone, 10 ng/mL epidermal
growth factor, 20 ng/mL cholera toxin plus 1% FCS (Table 1).
4. 10 concentrated digestion buffer I: 150,000 U collagenase,
50,000 U hyaluronidase, 50 mL DPBS. Mix, filter sterilize, and
use immediately or freeze in single use aliquots at 20 C.
5. Digestion buffer II: Dissolve 40 mg EGTA and 10 mg polyvinyl alcohol in 90 mL DPBS on a low heat with stirring, allow to
cool, and then add 10 mL 2.5% trypsin. pH to 7.4, filter
sterilize, and freeze in single use aliquots at 20 C.
Table 1
Recommended volumes of dissociation reagents for virgin mammary glands
Number of virgin mice (BL6)
1–2
3–5
6–8
Number of virgin mice (FVB/N)
1
2–4
5–7
Digestion buffer I
5 mL
10 mL
20 mL
Digestion buffer II
0.5–1 mL
2 mL
3 mL
Digestion buffer III
1 mL
2 mL
5 mL
DNase I volume
100
200
400
Mammary Stem Cell Isolation and Culture
219
6. Digestion buffer II: Dissolve 250 mg dispase in 50 mL DPBS.
Filter sterilize, and use within 1 week of storage at 4 C or
freeze in single use aliquots at 20 C.
7. 1 mg/mL DNase I solution: Dissolve 10 mg DNase I in 10 mL
medium, filter sterilize, and use within 1 week of storage at
4 C or freeze in single use aliquots at 20 C.
8. Optional, 1.25 concentrated red blood cell lysis solution; 8 g
NH4Cl in 1 L deionized water, filter sterilize, and store at 4 C.
9. Orbital shaking incubator.
10. McIlwain tissue chopper with standard table fitted with razor
blade as per manufacturer’s instructions.
2.2 Materials
for the Purification
of MaSC
and Progenitor Cells
by Flow Cytometric
Sorting
1. Single cell suspension of mammary gland cells (as obtained
from Subheading 3.2).
2. Sterile DPBS.
3. Sterile wash buffer.
4. Sterile collection buffer: DPBS with 10% FCS.
5. Fluorescently conjugated antibodies, refer to Table 2.
6. Viability dye such as propidium iodide (PI) or 7-actinomycin D
(7-AAD).
2.3 Materials
for the Culture
and Expansion
of Primary MECs
on a Feeder Layer
1. MEC as obtained from Subheadings 3.2 or 3.3
2. Sterile MEC medium supplemented with 1% FCS.
3. Sterile MEC medium supplemented with 5% FCS (5% MEC
medium, modified version of 1% MEC medium made in Subheading 2.1).
4. Sterile collagen coated 6-well tissue culture plates (see Note 1).
5. NIH/3T3 cells, irradiated (i3T3) (see Note 2).
6. 37 C incubator maintained with 5% CO2 and 5% O2.
2.4 Materials
for the Culture of MEC
in a Matrigel Colony
Assay
1. MECs as obtained from Subheadings 3.2 or 3.3.
2. Growth factor reduced Matrigel, thawed on ice.
3. 1% MEC medium
4. Glass chamber slides (Ibidi).
5. 37 C incubator maintained with a low O2 gas phase (5% CO2,
5% O2, and 90% N2)
6. Pipette tips, precooled to 4 C.
7. Harvest only: Cell recovery solution (BD Bioscience) and wash
buffer.
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Julie M. Sheridan and Jane E. Visvader
Table 2
Anti-mouse antibody clones that facilitate the identification and isolation of indicated mammary
epithelial stem and progenitor cell populations
Cell population and marker criteria
Antibody clone
(conjugate)
References
Blocks non-specific (FcγIII and FcγII) staining
CD16/CD32
(2.4G2)
Lineage cocktail (to deplete hematopoietic cells, red blood cells, and
endothelial cells)
CD45
TER-199
CD31
Lin CD29lo CD24+ CD14+ luminal progenitor-enriched
Lin CD29lo CD24+ CD14 mature luminal cell-enriched
(pregnant/lactating)
CD14
[13]
Lin CD29hi CD24+ basal cells incl. MaSC; Lin CD29lo CD24+
luminal cells incl. Luminal progenitor
CD29
[2]
Lin CD29hi CD24+ basal cells incl. MaSC; Lin CD29lo CD24+
luminal cells incl. Luminal progenitor
CD24
[2]
Lin CD24+ CD29lo CD49b+ Sca-1 hormone-receptor luminal
progenitor; Lin CD24+ CD29lo CD49b+ Sca-1+ hormonereceptor+ luminal progenitor
CD49b
[14]
Lin CD29lo CD24+ CD61+ LP-enriched; Lin CD29lo CD24+
CD61 mature luminal cell-enriched (virgin)
CD61
[6]
Lin CD24+ CD29lo CD49b+ Sca-1 hormone-receptor luminal
progenitor; Lin CD24+ CD29lo CD49b+ Sca-1+ hormonereceptor+ luminal progenitor
SCA-1
[14]
Lin CD29hi CD24+ TSPAN8hi/lo MaSC subsets
TSPAN8
[9]
3
Methods
Where possible, manipulations are performed using aseptic technique and/or in a sterile environment such as a tissue culture hood.
3.1 Harvesting
Mouse Mammary
Gland Tissue
1. Wipe the ventral midline of the euthanased mouse with 70%
ethanol.
2. Make an incision along the ventral midline through the skin
from the pubis to the neck taking care to avoid damage to the
peritoneal membrane (Fig. 1).
3. At the base of the midline incision, make further oblique cuts
toward and half of the way along the hind legs (Fig. 1).
4. Using forceps to hold the skin to one side of the junction of the
midline and oblique cuts, gently peel it outward to separate the
skin from the internal membrane to reveal the fourth and fifth
Mammary Stem Cell Isolation and Culture
221
3
4 LN
5
Fig. 1 Diagram of mammary gland dissection. Incisions (dashed lines) are made
through the skin along the midline and hind legs. The skin is peeled away from
the body and pinned in position to reveal the third, fourth, and fifth mammary
glands (3, 4, and 5, respectively). The inguinal lymph node (LN) can be removed
prior to excision of the fourth gland to minimize hematopoietic cell
contamination of the harvested tissue
mammary glands. This is best achieved using a second pair of
forceps to apply counter-pressure to the internal membrane to
prevent it from being pulled in the same direction as the skin.
5. Repeat this motion on the opposite side and pin the flayed skin
to the dissection board (Fig. 1).
6. Grasp the skin to one side of the midline cut at the level of the
forelegs and peel the skin outward from the body to reveal the
second and third mammary glands on the underside of the skin.
7. Repeat this motion on the opposite side and pin the flayed skin
to the dissection board (Fig. 1).
8. Remove and discard the inguinal lymph node that is located at
the intersection of the three prominent vessels on the surface of
the fourth mammary gland. This is best achieved by pushing
the skin upward from the outside, thus raising the area of the
vessel junction. In this position, the relative firmness of the
lymph node and its characteristically gray-white, circular
appearance facilitate its identification and excision using forceps or scissors.
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Julie M. Sheridan and Jane E. Visvader
9. Remove the mammary gland tissue by grasping and raising the
outer edge of the attached gland away from the skin and
membrane. Cut and peel the outer edges of the gland away
from the serous membrane and repeat this process while
moving toward the dorsal midline of the animal, releasing the
whole gland in one piece.
10. Place the mammary glands into MEC medium on ice.
3.2 Dissociation
of Mammary Gland
Tissue to a Single Cell
Suspension
1. Prepare the materials outlined in Subheading 2.1.
3.2.1 Preparation
3. Thaw a suitable amount of digestion buffers II and III and
DNase I at room temperature (Table 1).
2. Thaw and then dilute a suitable volume (Table 1) of 10 stock
of digestion buffer I in MEC medium to yield a 1 solution
and warm to 37 C.
4. Pre-warm a shaking incubator to 37 C.
3.2.2 Mechanical
Disruption
1. Prepare the McIlwain tissue chopper as per manufacturer’s
instructions. Briefly, with the machine turned off, affix a clean
plastic disc to the cutting table using the spring-loaded clips
and position the blade so that it touches the plastic disc. Once
ready, position the blade in the starting position at the righthand side of the plastic circle (see Notes 3 and 4).
2. Drain the mammary glands of excess buffer/medium and place
on the plastic disc. Multiple runs may be necessary to chop all
of the tissue depending upon the number of glands collected
and the mass of tissue obtained (see Note 4).
3. Turn the chopper on and allow to run across the tissue until it
stops. Raise the blade, return it to the starting position, and
rotate the plastic disc one quarter turn to crosscut the sample.
This process should be repeated for a total of four cutting runs
or until the gland no longer presents as lumps when lifted with
forceps (see Notes 4 and 5).
4. Place chopped tissue into a 50 mL conical tube for further
processing.
3.2.3 Enzymatic
Digestion to a Single Cell
Suspension
1. With reference to Table 1, add an appropriate volume of
pre-warmed 1 digestion buffer I, seal the tube, and place in
a shaking incubator at 37 C for 30 min (see Note 5).
2. Triturate the sample ten times using a pipette and incubate for a
further 20 min at 37 C.
3. Triturate the sample a further ten times and check for the
absence of large fragments. If any remain, incubate for a further
10 min and then re-triturate until a relatively homogeneous
organoid solution has been achieved.
Mammary Stem Cell Isolation and Culture
223
4. Add 20–40 mL wash buffer to the sample and centrifuge for
5 min at 1200 rpm (200 RCF) to collect the organoids.
5. Pre-warm digestion buffers II and III and DNase I to 37 C.
6. Using a vacuum pump or pipette, remove the supernatant.
Care must be taken to remove the concentrated band of adipose flocculate that overlays the supernatant (see Note 6).
7. Add DNase I to the organoid pellet, tap to partially resuspend,
and leave at room temperature for 2 min.
8. Add a suitable volume (Table 1) (see Note 5) of pre-warmed
digestion buffer II to the organoids, pipette gently to fully
suspend, and incubate in a water bath at 37 C for 2–3 min.
9. Inactivate and dilute the trypsin by adding 30 mL wash buffer.
Centrifuge for 5 min at 1200 rpm and discard the supernatant
(see Note 6).
10. Add DNase I to the organoid pellet, tap to partially resuspend,
and wait for 2 min.
11. Add a suitable volume (Table 1) (see Note 5) of pre-warmed
digestion buffer III to the organoids, pipette gently to resuspend the organoids, and incubate in a water bath at 37 C for
5 min.
12. Gently triturate the suspension to check for the presence of a
single cell suspension; no clumps should remain, and the
“grainy” appearance of the cell/organoid suspension that was
evident before digestion buffer III should now be gone.
13. Add 30 mL wash buffer and centrifuge for 5 min at 1200 rpm
(200 RCF) to collect the cells. Remove the supernatant to yield
a pellet of single cells and proceed with assays or further
purification.
3.3 Purification
of Mammary Epithelial
Stem and Progenitor
Cell Populations
1. Prepare the materials outlined in Subheading 2.2.
2. Isolate mammary cells using the protocol outlined in Subheading 3.2.
3.3.1 Preparation
3.3.2 Red Blood Cell
Lysis
1. Resuspend cells in 100 μL DNase I and 900 μL wash buffer.
2. Slowly add 4 mL of room temperature red blood cell lysis
buffer while swirling tube to facilitate gentle mixing.
3. Incubate at room temperature for 3 min.
4. Add 20 mL wash buffer and pass through a 40, 70, or 100 μm
cell strainer to remove any clumps.
5. Pass another 20 mL wash buffer through the cell strainer to
ensure maximal cell recovery.
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Julie M. Sheridan and Jane E. Visvader
6. Take a small aliquot to determine the cell concentration using a
hemocytometer or similar. Together with the known volume,
use the cell concentration to calculate the total cell count.
7. Centrifuge for 5 min at 1200 rpm to collect the cells. Discard
the supernatant and place the tube of cells on ice. Form this
point on, use chilled solutions and keep the cells on ice.
3.3.3 Staining a Single
Cell Suspension
for the Purpose
of Mammary Stem
and Progenitor Cell
Identification
What follows is a brief overview of a basic staining protocol. It is by
no means exhaustive, and other considerations not covered in this
protocol include the type of sorter to be used, the aim of the sort
(yield or purity), and antibody-fluorochrome panel design.
1. Prepare the materials outlined in Subheading 2.2.
2. Resuspend the cells in cold wash buffer at a concentration of
2.5 107 cells per mL.
3. [Optional] Incubate the cells with an appropriate volume of
unconjugated anti-mouse CD16/CD32 antibody (Table 2) to
minimize non-antigen-specific binding of fluorescently conjugated antibodies to Fc receptors.
4. Distribute the cells to staining vessels as required. Where necessary, include cells for single color and fluorescence-minusone controls.
5. Add primary antibodies to the cell suspensions (Table 2). Incubate on ice for 25 min.
6. Add wash buffer and centrifuge for 5 min at 1200 rpm
(200 RCF) to collect the cells. Discard the supernatant.
7. [Optional] When necessary, steps 5 and 6 may be repeated
with secondary antibodies.
8. Prepare collection tubes for cells post-sort containing a small
amount of collection buffer. Vessel choice and volume depend
on several factors including expected cell yield and sort nozzle
(and thus droplet) size. For example, a small number of cells
(10,000) might be sorted through a 100 μm nozzle into an
Eppendorf tube. However, a larger number of cells will require
a 5 mL FACS tube or multiple Eppendorf tubes.
9. Resuspend the cell pellet in a suitable volume of wash buffer
containing a viability dye such as 2 μg/mL propidium iodide
(PI) or 5 μg/mL 7-actinomycin D (7-AAD) in preparation for
sorting.
10. Fluorescence-minus-one controls and population contours
should be used to inform appropriate gate placement. Gates
may be configured to exclude doublets and debris on the basis
of light scatter properties, and viable cells can be selected on
the basis of viability dye exclusion. Several mammary epithelial
stem and progenitor cell populations lie within the Lineage
(Lin ) CD29hi CD24+ fraction and suggested sort criteria for
Singlets
Cells
Viable
FSC-A
Luminal
FSC-A
Basal/
MaSC
Mature
Luminal
Modal
CD24
CD45/TER-119/CD31
FSC-A
225
7-AAD
SSC-A
SSC-W
Mammary Stem Cell Isolation and Culture
Luminal
Progenitor
Lineageneg
CD29
CD29
CD61
Fig. 2 Representative plots showing a flow cytometric gating strategy designed to distinguish single, viable
CD29hi CD24+ basal cells, Lin CD29lo CD24+ CD61+ luminal progenitors, and Lin CD29lo CD24+ CD61
mature luminal cells
specific mammary stem or progenitor cell populations are outlined in Table 2 and Fig. 2.
11. Cells may be sorted on any flow cytometer and each requires
optimization. Consideration should be given to both nozzle
size and sort pressures since the factors can greatly affect viability of large fragile cells such as MECs (see Note 7).
12. [Optional] Reanalysis may be performed on a small aliquot of
collected cells to confirm the identity and establish the purity of
sorted cells.
3.4 Ex Vivo
Propagation of MEC
on a Feeder Layer
1. Prepare materials as per Subheading 2.3 under aseptic
conditions.
3.4.1 Preparation
3.4.2 Plating Cells
for Expansion
1. All cell manipulations are performed using aseptic technique in
a sterile TC hood.
2. For each well of a 6-well plate to be plated, mix 2.5 mL 5%
MEC medium with 100,000 i3T3 (100 μL of thawed stock,
final density approximately 14,000/cm2) and purified MEC.
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Julie M. Sheridan and Jane E. Visvader
Fig. 3 Representative bright-field images of colonies generated from plating
purified mammary epithelial stem and progenitor cell-enriched populations. (Top
panel) 300 Lin CD29hi CD24+ basal/MaSCs or Lin CD29lo CD24+ CD61+
luminal cells were plated in 2D feeder layer-based cultures. After 6 days,
cultures were briefly fixed and stained with Giemsa to visualize colony morphology. Scale bars, 250 μm. (Bottom panel) 1000 Lin CD29hi CD24+ basal cells or
1500 Lin CD29lo CD24+ luminal cells were grown for 14 days in the 3D Matrigel
assay. Scale bars, 500 μm. Inset, magnified region showing typical solid (basal)
or acinar (luminal) colony types. Scale bars, 250 μm
As a guide, 10,000–20,000 basal/MaSC or luminal cells from
FVB/N or 20,000–50,000 basal/MaSC or luminal cells from
C57BL/6 will give a MEC-dominated plated after 6 days in
culture.
3. Place in a 37 C incubator maintained at 5% CO2 and 5% O2.
4. Once situated, to ensure that cells are evenly distributed across
the plate, gently slide the plate forward and backward five times
and then repeat using a side-to-side motion.
5. 24 h later, viable cells will have attached. Discard the medium
in the well and immediately replace with fresh 1% MEC.
6. Maintain the culture by changing the MEC medium every
3 days. Cells may be manipulated at any time during culture
using transduction, transfection, or application of medium
additives as described elsewhere.
Mammary Stem Cell Isolation and Culture
227
7. Culture until the colonies of epithelial cells have expanded and
dominate the plate (typically less than 1 week with purified
basal or luminal cells). Several different colony morphologies
will contribute to epithelial outgrowth. These are most obvious
when cells are plated at lower densities as shown in Fig. 3.
3.4.3 Harvesting
Expanded MECs
1. Remove medium and wash cells gently with DPBS.
2. Add 0.5 mL pre-warmed trypsin-EDTA to each well and incubate at 37 C for 10–15 min.
3. Using a microscope, check to see that cells are detaching from
the plate. Luminal cells will detach more easily.
4. Using a P1000 pipette, gently triturate cells to aid detachment
from the plate and re-incubate if necessary.
5. Repeat steps 3 and 4 until cells are in a single cell suspension.
6. Collect cells into a falcon tube and add several times the volume
of cold wash buffer (containing FCS) to quench the trypsin
activity.
7. Centrifuge for 5 min at 1200 rpm to collect the cells for
downstream analyses.
3.5 Culture of MECs
as Colonies in Matrigel
3.5.1 Preparation
3.5.2 Organoid Culture
Initiation and Maintenance
1. Prepare materials as in Subheading 2.4.
Due to the small volumes involved with Matrigel culture initiation, manipulations should be done in bulk with master mixes,
where possible. Always prepare extra to accommodate unavoidable
losses, and keep plates, tubes, and pipette tips cold to facilitate the
handling of Matrigel, which polymerizes quickly when warmed.
1. Calculate the cell number and Matrigel volume to be plated. As
a starting point, 1000 single cells can be plated in one 20 μL
drop of Matrigel with 1 drop per well of an 8-chamber slide.
2. Prepare materials as per Subheading 2.4, taking into account
the quantity of material to be plated.
3. Cells to be plated are centrifuged in an Eppendorf tube and
medium aspirated to leave a minimum volume (not greater
than 10–15 μL).
4. Resuspend cell pellet in Matrigel by gentle trituration with a
pre-chilled pipette tip, taking care not to introduce any bubbles
(see Note 8).
5. Pipette 20 μL drop into each chamber of the slide (see Note 8).
6. Transfer to 37 C incubator for 10–15 min to allow Matrigel to
polymerize.
7. Gently pipette 400 μL of pre-warmed 1% MEC medium into
each well.
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Julie M. Sheridan and Jane E. Visvader
8. Return to incubator and maintain with medium changes every
3–4 days intervals.
9. After a total of 12–14 days, mature colonies may be imaged or
harvested for further analysis. Characteristic colony morphologies will be identifiable following culture (Fig. 3).
10. Cell recovery solution used as per manufacturer’s instructions
can be used to isolate colonies for downstream analyses (see
Note 9).
4
Notes
1. The use of plates pre-coated with collagen is not necessary but
shortens the time to confluency.
2. For production: Grow sufficient numbers of NIH3T3, e.g.,
1–5 T225 flasks, and harvest when actively growing but nearing confluence (~85% confluent). Harvest cells using trypsin,
quench with NIH3T3 growth medium containing 10% FCS,
centrifuge and resuspend cells in NIH3T3 growth medium
containing 10% FCS, and place on ice. Irradiate the cells with
50 Gy and count the cell suspension using an automated cell
counter or hemocytometer to determine the total cell number.
Spin and resuspend the cells at a density of 1 106 cells/mL in
cold freezing medium such as 50% DMEM + 40% FCS + 10%
DMSO. Aliquots of 1 mL (1 106 i3T3) can be frozen using a
typical cell freezing protocol for long-term storage.
3. When cells will be cultured, ensure that the cutting station is
clean and the plastic discs are clean and sterilized prior to use.
4. Slice thickness and strike force for the McIlwain tissue chopper
should be determined empirically. The chopped gland should
have only small lumps and have a thick slurry-like consistency.
5. This protocol generalizes the requirements to dissociate glands
from virgin mice into single cells. Due to the increased mass of
pregnant or lactating glands, volumes in Table 1 should be
adjusted commensurate with size.
6. Large flocculates may be observed that consist of viable cells
held together by DNA released from damaged cells. Although
these will disperse when DNase I is added, care must be taken
to retain them since they can be easily lost during supernatant
aspiration.
7. We routinely use 70 or 100 μm nozzles and sort at low-tomedium sort pressures to achieve consistent viability and yields
8. Avoid the introduction of bubbles into the Matrigel by careful
pipetting. If bubbles are introduced, they will be difficult to
remove but, by preparing a little extra Matrigel/cell suspension, they may be excluded during future pipetting strokes.
Mammary Stem Cell Isolation and Culture
229
A single bubble that has been introduced while pipetting 20 μL
drops into the chambers can be occasionally removed by simply
drawing the Matrigel back up into the tip and re-pipetting it.
9. To mitigate the sticky nature of 3D colonies during harvesting,
care should be taken to avoid unnecessary manipulations and
minimize the surfaces that they contact. Additional safeguards
include using pipette tips pre-wet with FCS to resuspend the
organoids, since tapping can distribute organoids across the
Eppendorf tube walls, which will result in cell losses.
References
1. Visvader JE, Stingl J (2014) Mammary stem
cells and the differentiation hierarchy: current
status and perspectives. Genes & Dev
28:1143–1158.
2. Shackleton M, Vaillant F, Simpson KJ et al
(2006) Generation of a functional mammary
gland from a single stem cell. Nature
439:84–88.
3. Stingl J, Eirew P, Ricketson I et al (2006)
Purification and unique properties of mammary
epithelial
stem
cells.
Nature
439:993–997.
4. Sleeman KE, Kendrick H, Robertson D et al
(2007) Dissociation of estrogen receptor
expression and in vivo stem cell activity in the
mammary gland. J Cell Biol 176:19–26.
5. Fu NY, Rios AC, Pal B et al (2017) Identification of quiescent and spatially restricted mammary stem cells that are hormone responsive.
Nat Cell Biol 19:164–176.
6. Asselin-Labat M-L, Sutherland KD, Barker H
et al (2007) Gata-3 is an essential regulator of
mammary-gland morphogenesis and luminalcell differentiation. Nat Cell Biol 9:201–209.
7. Bouras T, Pal B, Vaillant F et al (2008) Notch
Signaling Regulates Mammary Stem Cell
Function and Luminal Cell-Fate Commitment.
Cell Stem Cell 3:429–441.
8. Sheridan JM, Ritchie ME, Best SA et al (2015)
A pooled shRNA screen for regulators of primary mammary stem and progenitor cells
identifies roles for Asap1 and Prox1. BMC
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9. Jamieson PR, Dekkers JF, Rios AC et al (2017)
Derivation of a robust mouse mammary organoid system for studying tissue dynamics.
Development 144:1065–1071.
10. Smalley MJ, Kendrick H, Sheridan JM et al
(2012) Isolation of Mouse Mammary Epithelial Subpopulations: A Comparison of Leading
Methods. J Mammary Gland Biol Neoplasia
17:91–97.
11. Pal B, Bouras T, Shi W et al (2013) Global
changes in the mammary epigenome are
induced by hormonal cues and coordinated by
Ezh2. Cell Rep 3:411–426.
12. Asselin-Labat M-L, Sutherland KD, Vaillant F
et al (2011) Gata-3 negatively regulates the
tumor-initiating capacity of mammary luminal
progenitor cells and targets the putative tumor
suppressor caspase-14. Mol Cell Biol
31:4609–4622.
13. Li W, Ferguson BJ, Khaled WT et al (2009)
PML depletion disrupts normal mammary
gland development and skews the composition
of the mammary luminal cell progenitor pool.
Proc Natl Acad Sci USA 106:4725–4730.
14. Barcellos-Hoff MH, Aggeler J, Ram TG, Bissell MJ (1989) Functional differentiation and
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Chapter 17
An Organoid Assay for Long-Term Maintenance
and Propagation of Mouse Prostate Luminal Epithelial
Progenitors and Cancer Cells
Yu Shu and Chee Wai Chua
Abstract
Historically, prostate luminal epithelial progenitors and cancer cells have been difficult to culture, thus
hampering the generation of representative models for the study of prostate homeostasis, epithelial lineage
hierarchy relationship and cancer drug efficacy assessment. Here, we describe a newly developed culture
methodology that can efficiently grow prostate luminal epithelial progenitors and cancer cells as organoids.
Notably, the organoid assay favors prostate luminal cell growth, thus minimizing basal cell dominance upon
the establishment and continuous propagation of prostate epithelial cells. Importantly, organoids cultured
under this condition have demonstrated preservation of androgen responsiveness and intact androgen
receptor signaling, providing a representative system to study castration resistance and androgen receptor
independence.
Key words Organoid culture, Prostate luminal progenitors, Prostate cancer, Androgen receptor,
Genetically engineered mouse, Castration resistance
1
Introduction
Recent advances in the three-dimensional culture system have
enabled the maintenance and propagation of various cell lineages
from different organ systems under defined in vitro conditions
[1–3]. In normal prostate, the epithelium consists of three major
cell lineages, including luminal cells, basal cells as well as an
extremely rare neuroendocrine cell population [4]. The majority
of prostate cancers in comparison are adenocarcinomas, which are
characterized by an exclusively luminal phenotype [4]. In the past
decades, there have been many attempts to establish culture conditions for prostate luminal and cancer cells, but most of these studies
have yielded very little promising results [5]. However, these culture conditions are neither optimized for prostate luminal and/or
cancer cell survival nor favor long-term propagation of the luminal
Ivan Bertoncello (ed.), Mouse Cell Culture: Methods and Protocols, Methods in Molecular Biology, vol. 1940,
https://doi.org/10.1007/978-1-4939-9086-3_17, © Springer Science+Business Media, LLC, part of Springer Nature 2019
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Yu Shu and Chee Wai Chua
population. With prolonged culture, basal cells will eventually outcompete luminal cells and become the dominant cell population in
culture. When using primary mouse or human prostate specimens
to isolate different prostate epithelial cell lineages, introduction of
tissue dissociation procedures may lead to changes in lineage
marker expression (unpublished observation). Consequently, it is
extremely difficult to ascertain the lineage origin of sorted cell
populations to be used for subsequent optimization of culture
conditions.
The use of genetically engineered mouse (GEM) models has
provided excellent opportunities to study prostate homeostasis,
regeneration, tumor initiation, and progression [6]. In particular,
lineage tracing using GEM models has identified the first prostate
luminal epithelial progenitor population, termed castrationresistant Nkx3.1-expressing cells (CARNs) that can generate basal
and luminal cell populations and are capable of serving as a cell of
origin for prostate cancer [7]. Using CARNs as a starting population, we have developed and optimized a novel organoid assay that
can maintain and propagate the prostate luminal progenitor population as well as other prostate epithelial cells in the long term
(Figs. 1, 2 and 3) [8, 9]. Notably, using lineage-marked prostate luminal and basal cells derived from GEM models, we have
convincingly demonstrated that the culture method favors luminal
cell growth. Moreover, under this culture condition, prostate luminal progenitors can generate basal progeny, implying a model that
recapitulates in vivo condition.
While organoids derived from basal cells are generally much
smaller in size, co-culture of basal and luminal cells can promote
organoid growth (Fig. 4). These results have indicated that our
organoid methodology minimizes the basal cell dominance issue in
prostate epithelial culture and can serve as a representative model
for the study of prostate homeostasis. More importantly, the organoid assay has facilitated the establishment of various tumor organoid lines from GEM models of prostate cancer (Fig. 5). Notably,
derived organoids demonstrate preservation of androgen responsiveness and intact androgen receptor signaling, which are highly
crucial and relevant for the study of prostate homeostasis as well as
cancer initiation and progression (Fig. 6). Lastly, it is worth noting
that another study has also established an ENR (EGF, Noggin,
R-spondin)-based organoid culture condition, which enables the
growth of both prostate basal and luminal epithelial cells [10].
Because this assay favors the growth of basal over luminal cells
[10], it remains unclear whether basal and luminal cells in the
derived organoids can be maintained equally after serial passaging.
Here, we describe a comprehensive protocol, which involves the
use of hepatocyte-based medium that is cost-effective and easy to
prepare for the isolation, maintenence and propagation of mouse
prostate luminal progenitors and cancer cells as organoids. We also
Organoid Culture for Prostate Luminal progenitors and Cancer Cells
233
present detailed protocols for the characterization of various organoid types as well as growth and gene expression assessment of
prostate epithelial organoids in response to androgen withdrawal.
2
Materials
We have included suppliers and catalogue numbers for certain
reagents because in our hands, we have found that similar reagents
from other suppliers do not perform as well.
2.1
Mouse Models
See Table 1 for primers used to genotype major GEM models listed
below.
1. To lineage-mark the prostate luminal progenitor population,
CARNs, castrate 8–12 weeks old Nkx3.1CreERT2/+; R26R-YFP/
+ mice and treat the mice with tamoxifen (Sigma #T5648)
(9 mg per 40 g body weight in corn oil) by daily oral gavage
for 4 consecutive days after a month of castration. Dissect and
analyze the mice after a month of tamoxifen induction (see
Note 1).
2. Use the following Cre- or inducible Cre-expressing mouse
models to isolate different prostate epithelial and cancer cells
(see Note 2):
(a) Nkx3.1Cre/+; R26R-YFP/+: Lineage-marks all prostate
epithelial cells during embryonic stage
(b) Nkx3.1CreERT2/+; R26R-YFP/+: Lineage-marks mainly
prostate luminal cells as well as a small prostate epithelial
population that co-expresses basal and luminal markers
upon tamoxifen induction at adult stage
(c) CK8-CreERT2; R26R-YFP/+ or CK18-CreERT2; R26RYFP/+: Lineage-marks prostate luminal cells upon tamoxifen induction at adult stage
(d) Various types of GEM model of prostate cancer that carry
R26R-YFP allele, such as Nkx3.1CreERT2/+; Pten flox/flox;
R26R-YFP/+ (NP), Nkx3.1CreERT2/+; Pten flox/flox;
R26R-YFP/+
(NPK),
and
KrasLSL-G12D/+;
Nkx3.1CreERT2/+; Pten flox/flox; p53 flox/flox; R26R-YFP/+
(NPP53): Lineage-marks and oncogenic transforms prostate epithelial cells upon tamoxifen induction at adult
stage
3. Prepare the following mice for isolation of unlabeled prostate
epithelial and cancer cells:
(a) Wild-type C56BL/6 mice—for isolation of prostate epithelial cells
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Yu Shu and Chee Wai Chua
Table 1
Primers used for mouse genotyping
Allele
Amplicon
size
Forward
0
Reverse
Nkx3.1CreERT2 500 bp
5 -CAG ATG GCG CGG CAA
CAC C-30
50 -GCG CGG TCT GGC AGT AAA
AAC-30
Nkx3.1 wildtype
500 bp
50 -CTC CGC TAC CCT AAG
CAT CC-30
50 -GAC ACT GTC ATA TTA CTT
GGA CC-30
Nkx3.1 null
232 bp
50 -TTC CAC ATA CAC TTC
ATT CTC AGT-30
50 -GCC AAC CTG CCT CAA TCA
CTA AGG-30
Nkx3.1 wildtype
707 bp
50 -GTC TTG GAG AAG AAC
TCA CCA TTG-30
50 -GCC AAC CTG CCT CAA TCA
CTA AGG-30
CreERT2
500 bp
50 -CAG ATG GCG CGG CAA
CAC C-30
50 -GCG CGG TCT GGC AGT AAA
AAC-30
R26R-YFP
320 bp
50 -AAA GTC GCT CTG AGT
TGT TAT-30
50 -AAG ACC GCG AAG AGT TTG
TC-30
R26R wild-type 600 bp
50 -AAA GTC GCT CTG AGT
TGT TAT-30
50 -GGA GCG GGA GAA ATG GAT
ATG-30
R26R-Tomato
196 bp
50 -CTG TTC CTG TAC GGC
ATG G-30
50 -GGC ATT AAA GCA GCG TAT
CC-30
R26R-CAGYFP
212 bp
50 -ACA TGG TCC TGC TGG
AGT TC-30
50 -GGC ATT AAA GCA GCG TAT
CC-30
R26R wild-type 297 bp
50 -AAG GGA GCT GCA GTG
GAG TA-30
50 -CCG AAA ATC TGT GGG AAG
TC-30
Pten flox
320 bp
50 -CAA GCA CTC TGC GAA
CTG AG-30
50 -AAG TTT TTG AAG GCA AGA
TGC-30
Pten wild-type
156 bp
50 -CAA GCA CTC TGC GAA
CTG AG-30
50 -AAG TTT TTG AAG GCA AGA
TGC-30
Pten null
320 bp
50 -TTG CAC AGT ATC CTT
TTG AAG-30
50 -ACG AGA CTA GTG AGA CGT
GC-30
Pten wild-type
240 bp
50 -TTG CAC AGT ATC CTT
TTG AAG-30
50 -GTC TCT GGT CCT TAC TTC
C-30
KrasLSL-G12D
550 bp
50 -AGC TAG CCA CCA TGG
CTT GAG TAA GTC TGC
A-30
50 -CCT TTA CAA GCG CAC GCA
GAC TGT AGA-30
Kras wild-type
500 bp
50 -GTC GAC AAG CTC ATG
CGG GTG-30
50 -CCT TTA CAA GCG CAC GCA
GAC TGT AGA-30
TRAMP
650 bp
50 -GCG CTG CTG ACT TTC
TAA ACA TAA G-30
50 -GAG CTC ACG TTA AGT TTT
GAT GTG T-30
p53 flox
370 bp
50 -CAC AAA AAC AGG TTA
AAC CCA G-30
50 -AGC ACA TAG GAG GCA GAG
AC-30
(continued)
Organoid Culture for Prostate Luminal progenitors and Cancer Cells
235
Table 1
(continued)
Allele
Amplicon
size
Forward
p53 wild-type
288 bp
50 -CAC AAA AAC AGG TTA
AAC CCA G-30
50 -AGC ACA TAG GAG GCA GAG
AC-30
Hi-Myc
177 bp
50 -AAA CAT GAT GAC TAC
CAA GCT TGG C-30
50 -ATG ATA GCA TCT TGT TCT
TAG TCT TTT TCT TAA TAG
GG-30
Reverse
(b) Various types of GEM model of prostate cancer that are
not lineage-marked, such as TRAMP, Nkx3.1/, Hi-Myc
and Nkx3.1+/; Pten+/ mice (see Note 3).
2.2 General
Equipment
and Consumables
1. CO2 euthanasia chamber.
2. Laminar flow hood or biological safety cabinet.
3. Dissecting microscope.
4. Micro-dissecting instruments.
5. P20, P200, and P1000 pipettes and pipette tips.
6. Water baths set at 37 C and 55 C.
7. CO2 incubator.
8. Centrifuges (for Eppendorf and Falcon tubes).
9. Orbital shaker.
10. Hemocytometer/automated cell counter.
11. BD FACSAria cell sorter (or similar).
12. Olympus IX51 inverted microscope with fluorescent lamp,
camera, and computer (or similar).
13. Tissue processor and embedder.
14. Leica microtomes, histology water bath, and slide warmer
(or similar).
15. Leica cryostats (or similar).
16. Histology microscope.
17. Leica SP5 confocal microscope (or similar).
18. Eppendorf Mastercycler Realplex2 (or similar).
19. Luminometer plate reader.
20. Food steamer.
21. DAKO pen.
22. Humidified chamber.
23. Insulated cryofreezing container.
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Yu Shu and Chee Wai Chua
24. Sterile petri dishes.
25. 96-well low attachment plates (Corning #3474).
26. 1.5 mL Eppendorf tubes.
27. 15 and 50 mL Falcon tubes.
28. Cell strainer 40 and 70 μm.
29. 1.8 mL cryotubes.
30. Parafilm.
31. Cryomold for biopsy specimen 10 10 5 mm, 100/pcs.
32. Greiner 96-well CELLSTAR plate (or similar opaque-walled
96-well plate).
2.3 Prostate
Dissection and
Dissociation,
and Preparation
of Single Cell
Suspension
1. Phosphate-buffered saline (PBS) for dissection.
2. 10 collagenase/hyaluronidase solution (STEMCELL Technologies #07912).
3. Dulbecco’s Modified Eagle Medium: Nutrient Mixture F12
supplemented with 5% fetal bovine serum (FBS).
4. 0.25% trypsin-EDTA (STEMCELL Technologies #07901).
5. Hanks’ balanced salt solution modified (HBSS) (STEMCELL
Technologies #37150) supplemented with 2% FBS.
6. Dispase 5 U/mL (STEMCELL Technologies #07913).
7. DNase I solution 1 mg/mL (STEMCELL Technologies
#07900).
8. 0.4% trypan blue solution.
9. Reconstitute ROCK inhibitor Y-27632 (STEMCELL Technologies #72302) in PBS to make 5 mM stock solution (see
Note 4).
2.4 FluorescenceActivated Cell Sorting
(FACS)
1. APC anti-mouse EpCAM antibody (BioLegend #118214).
2. PerCP-Efluor710 anti-mouse/human E-cadherin antibody
(eBiosciences #46-3249-82).
3. Prepare 0.5 mg/mL DAPI solution by dissolving DAPI powder in Milli-Q water (see Note 5).
4. Prepare collecting medium consisting of HBSS supplemented
with 2% FBS and 10 μM ROCK inhibitor Y-27632.
2.5 Prostate
Epithelial Organoid
Culture
1. Hepatocyte Culture Media Kit (Corning #355056) (see
Note 6).
2. Prepare heat-inactivated charcoal-stripped FBS by heating the
reagent in 55 C water bath for 60 min (see Note 7).
3. 100 Glutamax.
4. Matrigel (Corning #354234) (see Note 8).
Organoid Culture for Prostate Luminal progenitors and Cancer Cells
237
5. 5 mM stock solution of ROCK inhibitor Y-27632.
6. 105 M dihydrotestosterone (DHT, Sigma #A-8380) in absolute ethanol (see Note 9).
7. 100 antibiotic-antimycotic.
2.6 Organoid
Passage
and Preparation
of Frozen Organoid
Stocks
1. Cold PBS.
2. 0.25% trypsin-EDTA.
3. HBSS supplemented with 2% FBS.
4. Organoid media with complete supplements (see Subheading
3.3).
5. Dispase 5 U/mL.
6. DNase I solution 1 mg/mL.
7. FBS.
8. Dimethyl sulfoxide (DMSO).
2.7 Histology
and Immunostaining
of Organoids
1. 10% neutral buffered formalin.
2. 4% paraformaldehyde (PFA) solution in PBS.
3. 30% sucrose in PBS.
4. Tissue-Tek OCT compound.
5. Collagen I, high concentration, rat tail, 100 mg (Corning
#354249).
6. Prepare setting solution for collagen I by combining 100 mL
10 RPMI (with phenol red), 7.5 mL 1 M NaOH, and
42.5 mL Milli-Q water (see Note 10).
7. Citrus Clearing Solvent (or xylene).
8. 70%, 95%, and 100% ethanol. Prepare 70% and 95% ethanol by
diluting 100% ethanol with Milli-Q water.
9. Hematoxylin and eosin stains.
10. 0.5% acid alcohol.
11. Scott water.
12. ClearMount mounting medium.
13. Prepare 3% hydrogen peroxide solution by diluting 30% hydrogen peroxide solution with Milli-Q water.
14. Antigen unmasking solution, citrate acid based.
15. Normal goat serum.
16. Prepare 1 PBS from 10 PBS solution.
17. Prepare 0.1% and 0.5% PBST by adding 1 and 5 mL of Triton
X-100 to 999 and 995 mL 1 PBS, respectively. Mix well the
solution prior to use.
18. Primary antibodies (refer to Table 2).
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Yu Shu and Chee Wai Chua
Table 2
Antibodies used for immunofluorescent staining
Antigen
Supplier
Species Dilution Remarks
AR
Sigma #A9853
Rabbit
CK5
BioLegend #905901
Chicken 1:500
Basal marker (formerly Covance
#SIG-3475)
CK5
BioLegend #905501
Rabbit
1:500
Basal marker (formerly Covance
#PRB-160P)
CK8
Abcam #ab14053
Chicken 1:500
Luminal marker (discontinued)
CK8
Abcam #ab53280
Rabbit
1:200
Luminal marker
CK8/18 Developmental Studies Hybridoma Rat
Bank #TROMA-1
1:100
Luminal marker
CK18
Abcam #ab668
Mouse
1:100
Luminal marker
FoxA1
Abcam #ab55178
Mouse
1:100
Epithelial lineage marker
GFP
Abcam #ab13970
Chicken 1:1000
GFP
Abcam #ab290
Rabbit
1:1000
GFP
Roche #11814460001
Mouse
1:100
p63
Santa Cruz #sc-8343
Rabbit
1:50
phospho- Cell Signaling #3787
Akt
Rabbit
1:100
phospho- Cell Signaling #4370
Erk
Rabbit
1:200
Ki67
Rat
1:1000 Proliferation marker
eBiosciences #14–5698
1:1000 With tyramide amplification (less
background staining)
Use 1:500 with tyramide
amplification
19. Alexa-fluor-conjugated secondary antibodies.
20. Tyramide signal amplification kit.
21. 0.5 mg/mL DAPI solution.
22. Antifade mounting medium.
2.8 Growth and Gene
Expression
Assessment
of Organoids
1. CellTiter-Glo 3D Cell Viability Assay (Promega #G9681).
2. TRIzol reagent.
3. MagMAX 96 for Microarray Total RNA Isolation Kit (Ambion,
Life Technologies #AM1839).
4. Superscript First-Strand Synthesis System for RT-PCR (Invitrogen #11904-018).
5. SYBR green master mix reagent.
6. Primers (refer to Table 3 for primer sequences).
Organoid Culture for Prostate Luminal progenitors and Cancer Cells
239
Table 3
Primers used for quantitative real-time PCR
Amplicon
Allele size
Forward
Reverse
Fkbp5 172 bp
5 -TGA GGG CAC CAG TAA CAA 50 -CAA CAT CCC TTT GTA GTG GAC
TGG-30
AT-30
Mme
198 bp
50 -CTC TCT GTG CTT GTC TTG 50 -GAC GTT GCG TTT CAA CCA GC-30
CTC-30
Psca
103 bp
50 -GGA CCA GCA CAG TTG CTT 50 -GTA GTT CTC CGA GTC ATC CTC
TAC-30
A-30
Igfbp3 101 bp
50 -CCA GGA AAC ATC AGT GAG 50 -GGA TGG AAC TTG GAA TCG GTC
TCC-30
A-30
3
0
Methods
The volume of reagents detailed below is for dissociation of an
intact prostate from an 8- to 12-week-old C57BL/6 wild-type
mouse. For larger prostate specimens, such as those from GEM
models of prostate cancer, adjust the volume of reagents proportionally (see Note 11).
3.1 Prostate
Dissection,
Dissociation,
and Preparation
of Single Cell
Suspension
1. In tissue culture hood, prepare 1 collagenase/hyaluronidase
solution by combining 200 μL 10 solution and 1.8 mL
DMEM/F12 supplemented with 5% FBS. Warm solution in
37 C water batch for at least 30 min prior to use.
2. Harvest whole urogenital system and transfer to sterile petri
dish containing cold PBS for prostate dissection. Using a dissecting microscope, fine forceps, and tweezers, remove residual
fats, connective tissues, and unrelated organs such as the bladder, ureters, seminal vesicles, and urethra (see Note 12).
3. Fill 1.5 mL Eppendorf tube with 0.75 mL pre-warmed 1
collagenase/hyaluronidase solution and transfer prostate tissue
into the tube. Using small and sharp sterile scissors, lacerate
prostate tissue by rapidly opening and closing scissors inside the
tube until no visible tissue chunks are seen. Fill up the tube by
adding 0.75 mL pre-warmed 1 collagenase/hyaluronidase.
Incubate in 37 C CO2 incubator for 3 h with periodic shaking
of tube once every hour (see Note 13).
4. Spin down digested tissue at 350 g for 5 min in pre-cooled
centrifuge and discard supernatant. Resuspend pellet in 1.5 mL
cold 0.25% trypsin-EDTA and incubate in 4 C refrigerator for
1 h (see Note 14).
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Yu Shu and Chee Wai Chua
5. During trypsin-EDTA incubation, warm 900 μL dispase in
37 C water bath for at least 10–15 min prior to use. At the
same time, thaw DNase I solution by leaving the tube carrying
the solution on ice. Immediately before use, warm 100 μL
DNase I solution briefly prior to combining with dispase
solution.
6. After trypsinization step, transfer digested tissue in trypsinEDTA into 15 mL Falcon tube containing 3 mL cold HBSS
supplemented with 2% FBS (equal to 2 volume of trypsinEDTA) to quench trypsin reaction. Centrifuge at 350 g for
5 min and discard supernatant.
7. Resuspend pellet with 1 mL pre-warmed dispase/DNase I
solution. Pipette vigorously for 1–2 min using P1000 pipette
to dissociate cells from extracellular matrices (see Note 15).
8. Add 5 mL cold HBSS supplemented with 2% FBS (equal to 5
volume of dispase 5 U/mL) to dilute dispase to <1 U/mL.
Using a 40 μm cell strainer, filter cell suspension into a sterile
50 mL Falcon tube.
9. Spin down at 350 g for 5 min and discard supernatant.
Resuspend pellet in 500 μL cold HBSS supplemented with
2% FBS and transfer to 1.5 mL Eppendorf tube. Perform viable
cells counting using a hemocytometer or automated cell
counter and trypan blue (see Note 16).
3.2 Isolation
of Lineage-Marked
Populations
and Unlabeled
Epithelial or Cancer
Cells through FACS
1. (A) To isolate lineage-marked CARNs or prostate cancer cells
derived from various GEM models of prostate cancer,
resuspend cells in cold HBSS supplemented with 2% FBS
and 0.05 μg/mL DAPI at 0.2–0.4 million cells/100 μL
solution (see Note 17).
(B) Prepare the following tubes for isolation of lineage-marked
population (using CARNs as an example):
(a) No stain control—105 cells isolated from regressed
prostates of C57BL/6 mice or castrated and
tamoxifen-induced Nkx3.1CreERT2/+ mice in 100 μL
HBSS supplemented with 2% FBS
(b) DAPI control—105 cells isolated from regressed prostates of C57BL/6 mice or castrated and tamoxifeninduced Nkx3.1CreERT2/+ mice in 100 μL HBSS supplemented with 2% FBS and 0.05 μg/mL DAPI
(c) Endogenous YFP control—105 cells isolated from
castrated and tamoxifen-induced Nkx3.1CreERT2/+;
R26R-YFP/+ mice in 100 μL HBSS supplemented
with 2% FBS
(d) Actual sample—one million cells isolated from castrated and tamoxifen-induced Nkx3.1CreERT2/+; R26R-
Organoid Culture for Prostate Luminal progenitors and Cancer Cells
241
YFP/+ mice in 100 μL HBSS supplemented with 2%
FBS and 0.05 μg/mL DAPI
Use tubes 1–3 to determine true YFP-positive cells. In
particular, use FACS plot of tube 1 to determine background staining; use FACS plot of tube 2 to exclude
DAPI positive cells, which are the dying cells; and then
compare FACS plots of tubes 2 and 3 to estimate
YFP-positive cells that are not DAPI positive.
2. (A) For isolation of unlabeled mouse prostate epithelial or
cancer cells, divide cell suspension into four 1.5 mL
Eppendorf tubes and perform antibody staining as follows:
(a) Unstained control—105 cells in 100 μL HBSS supplemented with 2% FBS
(b) APC single stain control—add 0.5 μL APC antimouse EpCAM antibody to 105 cells in 100 μL
HBSS supplemented with 2% FBS
(c) PerCP-Efluor710 single stain control—add 0.5 μL
PerCP-Efluor710 anti-mouse/human E-cadherin
antibody to 105 cells in 100 μL HBSS supplemented
with 2% FBS
(d) Actual sample—resuspend cells at one million cells
per 100 μL HBSS supplemented with 2% FBS. For
every million cells, add 2 μL APC anti-mouse
EpCAM antibody (1:50) and 0.5 μL PerCPEfluor710 anti-mouse/human E-cadherin antibody
(1:200) to the cell suspension
(B) Incubate tubes on ice and in dark for 25 min. Subsequently,
centrifuge at 350 g at 4 C for 5 min, discard supernatant, and wash cell pellet with PBS. Repeat centrifugation
again to spin down the cells. Lastly, resuspend cells in
control tubes and tube carrying the actual sample with
250 and 800 μL cold HBSS supplemented with 2% FBS
and 0.05 μg/mL DAPI, respectively.
3. Perform cell sorting through a sterile FACS facility and collect
lineage-marked cells, EpCAM+ and/or E-Cad+ prostate epithelial or cancer cells with sterile 1.5 mL Eppendorf tubes
containing 500 μL cold HBSS supplemented with 2% FBS
and 10 μM ROCK inhibitor Y-27632. Keep the collected
cells on ice until ready to plate (see Note 18).
Refer to Fig. 1 for representative FACS plots and sort gates for
isolation of lineage-marked CARNs and EpCAM+ and/or E-Cad+
prostate epithelial cells.
3.3 Establishment
of Organoid Culture
1. Prepare 50 mL basal organoid culture media by combining the
following components (see Note 19):
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Yu Shu and Chee Wai Chua
Fig. 1 Isolation of CARNs and prostate epithelial cells by flow cytometry. (a)
Representative FACS plot for isolation of lineage-marked YFP-positive CARNs.
(b) Representative FACS plot for isolation of EpCAM+ and/or E-Cad+ prostate
epithelial cells. Figure is adapted from Figs. 1b and 2a in ref. 8
(a) Hepatocyte medium (47 mL).
(b) 10 ng/mL EGF (100 μL of 5 μg/mL stock prepared in
PBS)
(c) 5% heat-inactivated charcoal-stripped FBS (2.5 mL)
(d) 1 Glutamax (500 μL 100 stock solution)
2. On the day of plating, prepare 10 mL (or desired amount)
complete organoid culture medium by combining the following components:
(a) Basal organoid medium (10 mL).
(b) 5% Matrigel (500 μL)
(c) 10 μM ROCK inhibitor Y-27632 (20 μL of 5 mM stock)
(d) 100 nM DHT (100 μL of 105 M stock)
(e) 1 antibiotic-antimycotic (100 μL 100 stock solution;
optional)
Warm prepared organoid culture media at 37 C for 30 min
prior to use (see Note 20).
3. Spin down sorted cells at 350 g for 5 min and then resuspend
in complete organoid culture media at a density of 250–1000
lineage-marked CARNs, 5000–10,000 prostate epithelial cells
or 1000–5000 prostate cancer cells per 100 μL media. For
example, if prostate epithelial cells are collected at 100,000
cells per Eppendorf tube, resuspend the cells in 1–2 mL
media. Transfer resuspended cells to 96-well low attachment
plate at 100 μL per well for desired final plating density.
4. Add 100 μL fresh medium every 4 days. When wells are almost
full after addition of new media for twice, transfer the media
from each well to a 1.5 mL Eppendorf tube. Centrifuge tubes
Organoid Culture for Prostate Luminal progenitors and Cancer Cells
243
at 250 g for 5 min, remove 250 μL supernatant, and add
100 μL fresh media (total volume will be approximately
150 μL) before transferring organoids back to a new 96-well
low attachment plate (see Note 21).
5. At day 7 of plating, calculate efficiency of organoid formation
by averaging numbers of visible organoids in each well under a
10 objective (see Note 22).
Refer to Figs. 2a, b, 3a, b, 4a, c, and 5a, f–j for morphology
of organoids derived from lineage-marked CARNs, EpCAM+
and/or E-Cad+ prostate epithelial cells, lineage-marked basal and
luminal cells as well as various GEM model-derived prostate cancer
cells.
3.4 Organoid
Propagation
and Preparation
of Frozen Stocks
1. After 3–5 weeks of plating, organoids can grow up to >200 μm
in diameter. To passage the organoids, transfer them into
1.5 mL Eppendorf tube, centrifuge at 350 g for 5 min, and
discard supernatant. Wash cell pellet in cold PBS and then
perform another round of centrifugation at 350 g for
5 min to spin down the cells. Repeat washing step until Matrigel is no longer seen on top of pelleted cells (see Note 23).
2. After removal of supernatant, add 1 mL warm 0.25% trypsinEDTA to tube and incubate in a 37 C water bath for
5–10 min. During the incubation period, pipette up and
down for 30 s occasionally with P200 pipette tip to facilitate
cell dissociation. In the case of very large organoids or organoids that are difficult to dissociate, separate dissociated cells
from large organoids by filtering cell suspension with a 70 μm
cell strainer. Quench trypsin reaction on dissociated cells with
addition of HBSS supplemented with 2% FBS (2 volume of
trypsin-EDTA) while continuing trypsinization on large organoids (see Note 24).
3. For single cell dissociation, add dispase/DNase I, pipette up
and down vigorously for 2 min, and filter cell suspension with
40 μm cell strainer.
4. Upon the completion of trypsinization step (with or without
dispase/DNase I step), transfer cell suspension into a 15 mL
Falcon tube prefilled with 2 mL cold HBSS supplemented with
2% FBS. Spin down the cells by centrifugation at 350 g for
5 min and discard supernatant. Resuspend cell pellet in fresh
media and plate onto a new 96-well low attachment plate (see
Note 25).
5. Freeze organoids at any point during a passage cycle by pooling
4–6 wells of organoids, centrifuging at 350 g for 5 min, and
resuspending in 1 mL freezing media containing 80% FBS, 10%
basal hepatocyte culture media, and 10% DMSO in 1.8 mL
cryotubes. Use an insulated cryofreezing container to gradually
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Yu Shu and Chee Wai Chua
freeze the cells to 80 C. Transfer frozen stocks to liquid
nitrogen tank for long-term storage.
6. To thaw frozen organoid stock, warm the cryotube rapidly in a
37 C water bath until only a small piece of ice is seen in the
tube. Immediately transfer 1 mL of organoids containing freezing media to a 15 mL Falcon tube prefilled with 10 mL cold
HBSS supplemented with 2% FBS to dilute DMSO. Spin down
thawed organoids at 350 g for 5 min, resuspend in organoid
culture media, and plate onto a new 96-well low attachment
plate (see Note 26).
3.5 Characterization
and Analysis
of Organoids
3.5.1 Histology
and Immunostaining
Organoid Specimens
Processing and Sectioning
1. To perform histopathological evaluation and molecular characterization, fix organoids with either 10% neutral buffered formalin (for paraffin embedding) or cold 4% PFA (for OCT
embedding) at room temperature (RT) for 1–4 h depending
on organoid size. Wash the fixed organoids with three changes
of cold PBS by repeatedly centrifuging at 250 g for 5 min,
discarding supernatant, and resuspending in fresh PBS.
2. For paraffin embedding, after last washing step, mix spun down
organoids with 20–50 μL cold 9:1 collagen I/setting solution
mixture. Prepare the collagen/setting solution mixture on ice
at all time. When resuspending organoids with collagen solution, use a P200 pipette tip that is cut at the tip region for wider
opening to allow homogeneous mixing of collagen and organoids. Avoid generation of bubbles during pipetting and place
the organoid and collagen solution mixture on parafilm as a
button and leave on a petri dish. Allow the collagen button to
solidify in 37 C incubator for 30 min (see Note 27).
3. Slowly transfer the solidified collagen button into 1.5 mL Eppendorf tube prefilled with 10% neutral buffered formalin and fix
overnight at RT. On the following day, wash the fixed button
with three changes of PBS and then leave the sample in a tissue
processor for dehydration, clearing, and infiltration of paraffin.
Embed the processed collagen button by pressing and leaving the
wider surface of the button on a stainless steel base mold before
filling up with paraffin and placing on a cold platform.
4. For OCT embedding, after washing and spinning down the
fixed organoids, resuspend them in cold 30% sucrose solution.
Incubate at 4 C for overnight or until all organoids sink to the
bottom of tube. Remove sucrose solution and transfer organoids in residual sucrose solution to cryomold for biopsy specimen prefilled with a thin layer of OCT compound. Use forceps
to mix organoid with OCT compound and then place them
close to the center base of cryomold using a dissecting microscope. Once organoids are positioned correctly, fill up
Organoid Culture for Prostate Luminal progenitors and Cancer Cells
245
cryomold with additional OCT compound and place on dry ice
to flash freeze the sample (see Note 28).
5. Cut paraffin- or OCT-embedded organoids at a thickness of
4 μm per section. Use freshly cut paraffin sections for best
immunostaining result. When cutting OCT blocks, perform
serial sectioning until organoids are no longer seen on the sections by checking through a histology microscope. Leave OCT
sections at RT for 30 min before storing at 20 or 80 C
freezer or proceeding with immunostaining (see Note 29).
Hematoxylin and Eosin
(H&E) and
Immunofluorescent (IF)
Stainings
1. Use paraffin sections for H&E staining. Perform paraffin clearing and rehydration of slides by going through two changes of
Citrus Clearing Solvent step (9 min each), two changes of
100% ethanol (7 min each), two changes of 95% ethanol
(3 min each), and one change of 70% ethanol (1 min) before
transferring to Milli-Q water for 5 min. From this point
onward, avoid drying out sections at all time.
2. Immerse the slides in hematoxylin solution for 2 min and 30 s,
transfer to running tap water for 1 min to wash out excessive
hematoxylin, and then immerse into 0.5% acid alcohol for 30 s
for differentiation. Put the slide in running tap water again for
1 min prior to a bluing step using Scott water for 1 min.
Immerse the slides again in tap water for 2 min before putting
into 95% alcohol for 1 min and then eosin solution for 3 min.
Prior to mounting, dehydrate the slide by going through four
changes of absolute alcohol for 30 s, 1 min, 2 min, and 3 min,
respectively (see Note 30).
Refer to Figs. 2c, d, 3c, d, and 5b, k–o for representative
H&E-stained images for organoids derived from lineagemarked CARNs, EpCAM+, and/or E-Cad+ prostate epithelial
cells and GEM model-derived prostate cancer cells.
3. Use paraffin or OCT section for IF staining. When using
paraffin sections, perform clearing and rehydration steps as
described in step 1. For OCT sections, bring out the sections
from freezer, leave at RT for 30 min to 1 h, and then wash with
two changes of Milli-Q water for 5 min each. From this point
onward, avoid drying out sections at all time. Block endogenous peroxidase activity of organoids with 3% hydrogen peroxide solution for 20 min and then wash in Milli-Q water for
5 min (see Note 31).
4. Perform antigen retrieval by immersing slides into citrate acidbased antigen unmasking solution and leave in a food steamer
for 45 min. Upon the completion of antigen retrieval step, cool
down slides by leaving in refrigerator or cold room for 15 min
and then wash with PBS for 5 min. Subsequently, perform
permeabilization step by incubating paraffin sections with
0.5% PBST for 15 min or OCT sections with 0.1% PBST for
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Yu Shu and Chee Wai Chua
10 min and followed by another washing step with PBS for
5 min.
5. Circle sections with DAKO pen to confine reagents used in the
following steps. Incubate in 10% normal goat serum in PBS
(or serum of species, in which secondary antibody is raised) at
RT for 1 h. Prepare primary antibody of choice (up to three
primary antibodies of different species) in 5% normal goat serum
in PBS. Apply antibody (or mixture of antibodies) to sections
and incubate at 4 C for overnight in a humidified chamber
(refer to Table 2 for information of primary antibodies).
6. Wash sections with one change of 0.1% PBST and two changes
of PBS for 5 min in each change. Prepare fluoresceinconjugated secondary antibody that recognizes primary antibody used (up to three secondary antibodies conjugated with
different fluorochromes) in 5% normal goat serum in PBS with
addition of 1:500 0.5 mg/mL DAPI solution. Apply antibody
(or mixture of antibodies) to sections and incubate at RT for
1 h (see Note 32).
7. In the case of tyramide signal amplification, prepare
HRP-conjugated secondary antibody alone or together with
other fluorochrome-conjugated secondary antibodies in 5%
normal goat serum in PBS with addition of 1:500 0.5 mg/
mL DAPI solution. Apply antibody (or mixture of antibodies)
to sections and incubate at RT for 1 h.
8. Wash sections with one change of 0.1% PBST and two changes
of PBS for 5 min in each change (in the case of tyramide signal
amplification, refer to step 9). Mount sections with antifade
mounting medium and then cover with coverslip. Press on the
coverslip to remove excessive mounting medium and bubbles,
and seal coverslip with nail polish.
9. Prepare tyramide amplification solution as suggested by the
manufacturer’s protocol. Apply tyramide amplification solution
at RT for 6 min (see Note 33). Wash sections with 0.1% PBST/
PBS (5 min, 0.1% PBST—one change and followed by PBS—
two changes). Repeat step 8.
10. Store slides in dark at 4 C. For consistent result, visualize
stainings and take images within 1–2 weeks using a confocal
microscope.
Refer to Figs. 2e–h, 3e–h, 4b, d, f, 5c–e, and 6c, d for IF
stainings of various markers in organoids derived from lineagemarked CARNs, EpCAM+, and/or E-Cad+ prostate epithelial
cells, lineage-marked basal and luminal cells, GEM model-derived
prostate cancer cells, as well as prostate epithelial organoids grown
in the presence or absence of DHT.
Fig. 2 Assessment of organoids derived from CARNs. (a, b) Bright-field (a) and epifluorescent (b) views of
CARN-derived organoids that are either partially filled or with hollow lumen (arrow). (c, d) Representative H&Estained CARN-derived organoids at low (c) and high (d) magnification views. (e–h) Organoid derived from
lineage-marked CARNs demonstrates uniformed YFP expression and high cellular proliferation as shown by
Ki67 immunostaining (arrows, e) and correct localization of basal cells at the outer region as marked by CK5
(arrowheads) and the luminal cells at the internal region as marked by CK8 (f). The organoid also expresses AR
(arrows, g) and epithelial lineage marker, FoxA1 (h). Scale bars in a–c corresponding to 100 μm and in d–h to
50 μm. Figure is adapted from Fig. 1c–j in ref. 8
Fig. 3 Characterization of organoids derived from EpCAM+ and/or E-Cad+ prostate epithelium. (a, b) Low (a)
and high (b) magnification views of organoids derived from EpCAM+ and/or E-Cad+ sorted prostate epithelial
cells at 20 days of plating that demonstrate heterogeneous phenotype. (c, d) Representative H&E-stained
EpCAM+ and/or E-Cad+ sorted prostate epithelial cell-derived organoids. (e–h) Organoid derived from prostate
epithelial cells expresses proliferation marker, Ki67 (arrows, e), demonstrates localization of basal cells at the
outer layer as marked by p63 (arrowheads) and the luminal cells at the internal layers as marked by CK18 (f).
The organoid also contains cells that co-expresses AR and CK8 (arrows, g) as well as epithelial lineage
marker, FoxA1 (arrows, h). Scale bars in a–c corresponding to 100 μm and in d–h to 50 μm. Figure is adapted
from Fig. 2b–f and h–j in ref. 8
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Yu Shu and Chee Wai Chua
Fig. 4 Assessment of organoid-forming ability of lineage-marked basal or luminal prostate epithelial cells. (a)
Organoid derived from YFP-labeled prostate basal epithelial cells. (b) CK5-trace organoid contains mostly
CK5-positive cells, including internal cells (arrowheads). (c) Organoid derived from YFP-labeled prostate
luminal epithelial cells. (d) CK8-trace organoid contains some CK5-positive cells at the outer layer (arrowheads). (e) Comparison of organoid formation efficiency between CK5-trace (n ¼ 4 experiments), CK8-trace
(n ¼ 3 experiments), and CK18-trace (n ¼ 2 experiments) prostate epithelial cells. (f) Organoid derived from
mixing of red CK18-trace luminal cells and green CK5-trace basal cells demonstrates green cells at the outer
layer (arrowheads), consistent with the localization of basal cells. Scale bars in a–d and f corresponding to
50 μm. Figure is adapted from Fig. 3c–e, g, h, and n in ref. 8
Organoid Culture for Prostate Luminal progenitors and Cancer Cells
249
Fig. 5 Generation of tumor organoids from GEM models of prostate cancer. (a–e) Organoids derived from
transformed CARNs isolated from castrated and tamoxifen-induced Nkx3.1CreERT2/+; Pten flox/flox; KrasLSL-G12D/
+
; R26R-YFP/+ (NPK) mice. NPK-CARN tumor organoid is YFP positive with extensive budding (a) and mostly
filled morphology as evidenced by H&E staining (b), highly proliferative as marked by Ki67 (c) and expresses
pAKT (d) and patchy pERK (arrows, e), consistent with the Pten deletion and Kras-G12D activation phenotype.
(f–o) Bright-field (f–j) and H&E-stained sections of tumor organoids derived from various GEM models of
prostate cancer through FACS sorting of EpCAM+ and/or E-Cad+ population. Showings are organoids
generated from 22-week-old TRAMP mice (f, k), tamoxifen-induced Nkx3.1CreERT2/+; Pten flox/flox; p53 flox/flox
(NPP53) mice at 2 months old and assayed after 8 months (g, l), 14-month-old Nkx3.1/ mice (h, m),
9-month-old Hi-Myc transgenic mice (i, n) and 10-month-old Nkx3.1+/; Pten+/ mice (j, o). Scale bars in a,
b, and f–o corresponding to 100 μm and in c–e to 50 μm. Figure is adapted from Figs. 4b–f and 5a–j in ref. 8
3.5.2 Growth and Gene
Expression Assessment
(See Note 34)
CellTiter-Glo Assay
for Growth Assessment
1. To assess cell growth in response to androgen withdrawal,
passage prostate epithelial cell-derived organoids and plate
equal amount of organoids onto 96-well low attachment
plate in the presence or absence of 100 nM DHT (see
Note 35).
2. Assay cell viability at days 1, 3, and 5 after plating using
CellTiter-Glo 3D (Promega) with five technical replicates for
each time point. Briefly, thaw CellTiter-Glo 3D reagent at 4 C
and leave at RT for at least 15 min prior to use. Add 100 μL of
the reagent into each well, which contains approximately
100 μL culture medium.
3. Shake the 96-well low attachment plate vigorously at RT for
10 min and then transfer the mixture to an opaque-walled
96-well plate. Incubate at RT for 10 min prior to measurement
using a luminometer plate reader.
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Yu Shu and Chee Wai Chua
Quantitative Real-Time
PCR Analysis
1. For RNA extraction, transfer four to six wells of organoids to a
1.5 mL Eppendorf tube, centrifuge at 350 g for 5 min, and
discard supernatant. Dissolve pelleted organoids in TRIzol
reagent followed by processing using the MagMAX 96 for
Microarray Total RNA Isolation Kit as suggested by manufacturer’s recommendation.
2. Use 100–200 ng of RNA as template for cDNA synthesis using
the Superscript First-Strand Synthesis System according to
manufacturer’s recommendation. Perform quantitative realtime PCR by mixing cDNA, primers (refer to Table 3 for
primer sequences used) and SYBR green master mix reagent
in the Eppendorf Realplex2 instrument in a triplicate manner
for each culture condition. Use the ΔΔCT method to obtain
expression values and normalize values to GAPDH expression.
Repeat the experiment with additional biological replicates (see
Note 36).
Refer to Fig. 6e for difference of various androgen-responsive
genes between prostate epithelial organoids grown in the presence
or absence of DHT.
Fig. 6 Androgen responsiveness of organoid derived from EpCAM+ and/or E-Cad+ prostate epithelial cells. (a,
b) Organoids were passaged and plated in the presence (a) or absence of (b) DHT. (c, d) In the presence of
DHT, organoids demonstrate strong nuclear AR expression, (c) whereas in the absence of DHT, AR expression
of the organoids is weak and localized at the cytoplasm (d). (e) Quantitative real-time PCR analysis of
androgen-responsive genes, Fkbp5, Mme, Psca, and lgfbp3, in organoids cultured in the presence or absence
of DHT. Scale bars in a and b corresponding to 100 μm and in c and d to 50 μm. Figure is adapted from
Fig. 2l–p in ref. 8
Organoid Culture for Prostate Luminal progenitors and Cancer Cells
4
251
Notes
1. Make sure tamoxifen is fully dissolved in corn oil before filtration by repeatedly warming mixture of tamoxifen powder and
corn oil in 37 C water bath for 10 min and then vortexing
vigorously for 15 min. Filtered tamoxifen solution should be
stored at 4 C and used within a month of preparation.
2. We use identical tamoxifen treatment schedule, route, and dose
for different inducible Cre mice.
3. Different GEM models of prostate cancer require different
durations and genetic backgrounds to progress into hyperplasia
and/or cancer.
4. Aliquot and keep the 5 mM stock solution at 20 C until use.
Avoid repeated freeze and thaw cycles for optimal results. We
recommend using ROCK inhibitor Y-27632 from STEMCELL Technologies for consistent organoid formation.
5. After DAPI is fully dissolved in Milli-Q water, filter, aliquot,
and keep the stock solution at 20 C. Once thawed, the
working stock solution should be in good condition for at
least 3 months if it is kept at 4 C and without extensive light
exposure.
6. This reagent has been regularly on back order. Make sure
sufficient reagent is available prior to the start of a new
experiment.
7. The use of heat-inactivated charcoal-stripped FBS is crucial for
efficient organoid formation from prostate luminal cells. After
the completion of heat inactivation, aliquot and keep the
reagent at 80 C prior to use.
8. Thaw Matrigel by placing the reagent on ice in a 4 C refrigerator or cold room for overnight. Once thawed, Matrigel must
remain on ice at all time to prevent polymerization. For consistent results, we avoid using Matrigel that has undergone more
than two freeze-thaw cycles.
9. Keep DHT solution at 20 C at all time. Avoid leaving
working solution outside the freezer for prolonged period
when preparing culture medium.
10. Make sure there is no precipitation when using setting solution
to ensure proper collagen I neutralization and solidification.
11. For regressed prostate, we use identical volume of reagents that
are used for intact mouse prostate dissociation because prostate
epithelium in castrated mice is high in cell density and more
collagenous.
12. For detailed description of urogenital system harvest and prostate dissection procedure, refer to Lukacs et al. [11].
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Yu Shu and Chee Wai Chua
13. For optimal dissociation, place Eppendorf tube on its side on a
sterile petri dish during incubation to maximize surface area of
prostate tissue exposed to collagenase/hyaluronidase solution.
In addition, periodic shaking of tube helps redistributing
tissues.
14. Place a petri dish on ice and leave tube on the petri dish.
Subsequently, place the whole ice bucket into the refrigerator
or cold room. Avoid use of warm trypsin-EDTA as this will
cause over-trypsinization, consequently leading to extensive
cell death.
15. Vigorous pipetting is crucial for optimal cell dissociation.
Pipette sample until the solution is translucent without visible
tissue fragments. Do not exceed 2 min for this step as this will
cause extensive cell death.
16. Using this protocol, we should expect 1.5–4 million of cells
isolated from intact prostate of an 8- to 12-week-old C57BL/6
mouse.
17. We use DAPI to exclude dead cells because they can be mistakenly identified as YFP-positive cells during flow sorting. However, high concentration of DAPI staining is lethal to cells.
Avoid adding >0.1 μg/mL DAPI in resuspension media.
18. Use sheath pressure not exceeding 12 psi for cell sorting as
higher sheath pressure will cause luminal cell death. It is also
important to include media supplemented with ROCK inhibitor Y-27632 in the collecting tube to prevent cell loss and
death.
19. Basal organoid culture media should be finished within a
month of preparation to ensure consistent results.
20. Avoid extensive warming in 37 C water bath because this may
cause Matrigel to solidify on top of the media.
21. Use P1000 pipette tip to transfer organoids as smaller tips may
damage structure of organoids. If there are too many wells for
media change, pool multiple wells prior to centrifugation and
redistribute evenly when plating.
22. Organoid growth should be evidenced as early as days 2–3
(CARNs or prostate cancer cells) or days 4–5 (prostate epithelial cells) of plating. Larger size organoids will occasionally fuse
together. Therefore, quantification of organoid formation
should be done by days 7 of plating for accurate estimation of
organoid-forming efficiency.
23. Trypsinization step will not be optimal with residual Matrigel,
consequently leading to incomplete cell dissociation.
24. Prostate epithelial cell-derived organoids that are solid and do
not have obvious lumen are harder to dissociate. These
Organoid Culture for Prostate Luminal progenitors and Cancer Cells
253
organoids contain more basal cells and require longer trypsinization duration during dissociation. In addition, prolonged
trypsin-EDTA incubation of more easily dissociated cells will
cause extensive death in this population. Therefore, adoption
of different trypsinization time for different cell lineages can
ensure continuous propagation of different lineage populations in organoid culture.
25. Passage organoids in a ratio of 1:4–1:6, i.e., dissociated cells
from 1 well can be passaged into 4–6 wells.
26. During thawing, do not over-warm the freezing media as warm
DMSO is lethal to the cells.
27. After neutralization by setting solution, collagen solution will
change color from yellow to pink or red once it is solidified.
28. Make sure organoids are mixed well with OCT compound and
embedded into it to provide proper support during tissue
sectioning.
29. Avoid over-trimming when sectioning and always check sections with a histology microscope to confirm the presence of
organoids on sections. Serial sectioning of OCT blocks can
minimize tissue loss as each block may only carry 20–30 4 μm
sections with intact organoid structure.
30. Check quickly the slides using a histology microscope after
bluing step to see whether hematoxylin staining provides
great nuclear details as well as after eosin step to see whether
the counterstaining gives good nuclear/cytoplasmic contrast.
Avoid using aged hematoxylin and eosin stains for optimal
staining results.
31. Exclude this step if HRP-enzymatic reaction is not involved in
the subsequent staining procedures.
32. When using two to three fluorochrome-conjugated secondary
antibodies, avoid selecting combination of fluorochromes that
have overlapped emission spectrums. Use Alexa-fluor 488 and
555 for combination of two antibodies and Alexa-fluor
488, 555, and 647 for combination of three antibodies.
33. Do not exceed 6-min incubation time as longer incubation will
lead to high staining background.
34. The protocols are also applicable for drug assessment on prostate luminal progenitors and cancer cells.
35. Instead of supplementing 5% Matrigel in culture medium, use
hepatocyte medium supplemented with 2% Matrigel for this
assay. With prolonged culture period, higher concentration of
Matrigel can affect accuracy of luminescence reading.
36. Dilute cDNA samples 1:5–1:10 when necessary.
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Yu Shu and Chee Wai Chua
Acknowledgments
We thank M. Shibata, S. Irshad, H.H. Zhu, M. Shen, W.Q. Gao,
and members of the Chua lab (M.Y. Liu, Y. Zhang, X. Cai, and
C.P. Liang) for insightful comments on the manuscript. This work
was supported by grants from the National Natural Science Foundation of China (81672548 and 81874098 to C.W. Chua), the
Program for Professor of Special Appointment (Eastern Scholar),
the Shanghai Institutions of Higher Learning (2016012 to
C.W. Chua), the State Key Laboratory of Oncogenes and Related
Genes (90-17-01 to C.W. Chua), and the Shanghai Municipal
Education Commission-Gaofeng Clinical Medicine Grant Support
(20171917 to C.W. Chua).
Author Contributions: Together with M. Lei, C.W. Chua developed
and optimized the organoid culture protocol in M. Shen’s lab at the
Columbia University. C.W. Chua and Y. Shu wrote the book chapter manuscript, and prepared the tables as well as the figures that
were adapted from original figures in Chua et al. [8].
References
1. Debnath J, Brugge JS (2005) Modelling glandular epithelial cancers in three-dimensional
cultures. Nat Rev Cancer 5:675–688
2. Clevers H (2016) Modeling development and
disease with organoids. Cell 165:1586–1597
3. Fatehullah A, Tan SH, Barker N (2016) Organoids as an in vitro model of human development and disease. Nat Cell Biol 18:246–254
4. Shen MM, Abate-Shen C (2010) Molecular
genetics of prostate cancer: new prospects for
old challenges. Genes Dev 24:1967–2000
5. Peehl DM (2005) Primary cell cultures as
models of prostate cancer development.
Endocr Relat Cancer 12:19–47
6. Irshad S, Abate-Shen C (2013) Modeling prostate cancer in mice: something old, something
new, something premalignant, something metastatic. Cancer Metastasis Rev 32:109–122
7. Wang X, Kruithof-de Julio M, Economides
KD, Walker D, Yu H et al (2009) A luminal
epithelial stem cell that is a cell of origin for
prostate cancer. Nature 461:495–500
8. Chua CW, Shibata M, Lei M, Toivanen R, Barlow LJ et al (2014) Single luminal epithelial
progenitors can generate prostate organoids
in culture. Nat Cell Biol 16:951–961
9. Chua CW, Shibata M, Lei M, Toivanen R, Barlow LJ, Shen MM (2014) Culture of mouse
prostate organoids. Protoc Exch. https://doi.
org/10.1038/protex.2014.037
10. Karthaus WR, Iaquinta PJ, Drost J,
Gracanin A, van Boxtel R et al (2014) Identification of multipotent luminal progenitor cells
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159:163–175
11. Lukacs RU, Goldstein AS, Lawson DA,
Cheng D, Witte ON (2010) Isolation, cultivation and characterization of adult murine prostate stem cells. Nat Protoc 5:702–713
Chapter 18
Isolation, Purification, and Culture of Mouse Pancreatic
Islets of Langerhans
Youakim Saliba and Nassim Farès
Abstract
Pancreatic islets constitute an important tool for research and clinical applications in the field of diabetes.
They are used for transplantation, unraveling new mechanisms in insulin secretion, studying pathophysiological pathways in diseased cells, and pharmacological research aimed at developing improved therapeutic
strategies. Therefore, fine-tuning islet isolation protocols remains an important objective for reliable
investigations. Here we describe a relatively simple mouse islet isolation protocol that relies on enzymatic
digestion using low-activity collagenase and several sedimentation and Percoll gradient steps.
Key words Islets of Langerhans, Pancreas, Isolation protocol, Mouse
1
Introduction
Islets of Langerhans constitute an important experimental tool in
the field of diabetic research, and achieving successful cell isolation
and culture is the most important requisite for reliable studies.
Since Lacy’s group described a new enzyme-based method for
islet isolation in 1967 [1], different protocols from rodents to
humans have been proposed and fuelled the advances in islet
research [2–16]. The main difference between all these procedures
resides in the enzyme blends and their delivery route, with the
common bile duct as the most commonly used [10, 13, 17,
18]. Islet separation and purification are then performed by Percoll
[19, 20] or Ficoll gradient separation [15], filtration [21], or
magnetic retraction [4, 22] and handpicking under a microscope
[8, 10, 12, 15, 18, 23–25]. However, many hurdles still persist in
these isolation procedures rendering it difficult to achieve reproducible maximum yield of viable islets that retain their in vivo
characteristics. Besides, the fine details to perform the isolation
that usually requires delicate microsurgery maneuvers are often
lacking, leading to sometimes conflicting data when it comes to
yield and function [26, 27]. Therefore, refining and revisiting islet
Ivan Bertoncello (ed.), Mouse Cell Culture: Methods and Protocols, Methods in Molecular Biology, vol. 1940,
https://doi.org/10.1007/978-1-4939-9086-3_18, © Springer Science+Business Media, LLC, part of Springer Nature 2019
255
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Youakim Saliba and Nassim Farès
isolation protocols remain important for more and more reliable
research.
In the protocol described here, mouse pancreas was excised and
digested with a low-activity collagenase eliminating the need for
microsurgery expertise. Since Percoll is chemically inert with minimum cellular toxic effects as compared to other cell separation
gradient media [9, 21, 28], it was used here to purify the islets
from the isolated pancreatic tissue. This purification step was preceded by several sedimentations and repeated to further purify the
islets without time-consuming handpicking [5, 10, 13]. Islets were
cultured in RPMI 1640 medium that has been shown to be optimal
for keeping islets into culture [2]. Finally, variations in the islet
numbers have been shown to exist between inbred mice [29], and
age differences have been also noted [30–34]. In the current protocol, we isolated islets from male and female mice of different ages
and species and obtained similar yields ranging from 750 to 1200
healthy islets per pancreas.
2
Materials
All chemicals used in the protocol are of reagent grade (>99%).
Deionized water with a resistivity of 18.2 mΩ cm at 25 C was used.
2.1 Modified Tyrode
Solution
1. Add approximately 500 mL water in 1 L autoclaved glass
beaker. Weigh each of the following and add consecutively to
the beaker while stirring with a magnetic stir bar: 8.18 g NaCl,
0.425 g KCl, 0.346 g MgCl2·6H2O, 0.37 g NaHCO3, 0.204 g
KH2PO4, 2.383 g HEPES, 1.492 g creatine monohydrate,
2.5 g taurine, and 2.11 g D-glucose. Adjust the pH to 7.2
with 1 M NaOH, and then make up the solution to 1 L. This
solution will have the following composition: 140 mM NaCl,
5.7 mM KCl, 1.7 mM MgCl2, 4.4 mM NaHCO3, 1.5 mM
KH2PO4, 10 mM HEPES, 10 mM creatine monohydrate,
20 mM taurine, and 11.7 mM D-glucose.
2. Filter the Tyrode solution on a sterile glass filter beaker using
0.22 μm membrane filter (Millipore, Merck, Massachusetts,
USA) under a laminar flow hood and store it at 4 C (see
Note 1).
2.2 Collagenase
Solution
1. Prepare 10 mL of collagenase solution per mouse pancreas.
Weigh in a 15 mL conical tube using the following: 10 mg
collagenase A (activity >0.15 U/mg) (Roche Diagnostics,
Basel, Switzerland) kept at 20 C and 10 mg bovine serum
albumin (BSA) kept at 4 C (see Note 2).
Mouse Islet Isolation
257
2. Under a laminar flow hood, filter the enzyme solution using
0.22 μm membrane syringe filter, transfer it in a new sterile
15 mL conical tube, and keep it on ice until used (see Note 3).
2.3
Percoll Solution
1. Percoll is a colloidal PVP-coated silica for cell separation with a
density of 1.13 0.005 g/mL.
2. Prepare 500 mL of a 10 concentrated NaCl solution (1.5 M)
by diluting 43.83 g NaCl in water. Prepare 500 mL of a 1
concentrated NaCl solution (0.15 M) by either diluting the
10 concentrated NaCl or by formulating 1 concentrated
NaCl solution directly from powder.
3. Prepare a stock isotonic Percoll (SIP) solution by adding
9 parts of Percoll to 1 part of 1.5 M NaCl (10 concentrated)
(see Note 4). In the following calculations, we will use an
example of 10 mL SIP solution, that is, 9 mL Percoll and
1 mL 1.5 M NaCl.
4. Calculate the density of the SIP solution from the following
formula:
V X ¼ V 0 ðρ0 ρi Þ=ðρi ρ10 Þ so ρi
¼ ½ðV 0 ρ0 Þ þ ðV X ρ10 Þ=ðV X þ V 0 Þ
where VX is the volume of diluting medium (mL)
V0 is the volume of undiluted Percoll (mL)
ρ0 is the density of Percoll (1.13 0.005 g/mL)
ρ10 is the density of 1.5 M NaCl (1.058 g/mL)
ρi is the density of the SIP solution (g/mL)
The density of SIP obtained is 1.123 g/mL
5. Dilute the SIP solution to a final density of 1.045 g/mL (see
Note 5) by adding 0.15 M NaCl using the following formula:
V y ¼ V i ðρi ρÞ= ρ ρy
where Vy is the volume of diluting 0.15 M NaCl (mL)
Vi is the volume of SIP (mL)
ρi is the density of SIP (g/mL)
ρy is the density of 0.15 M NaCl (1.0046 g/mL)
ρ is the density of final diluted Percoll solution (g/mL)
In order to dilute all the SIP solution (10 mL) to a final density
of 1.045 g/mL, proceed by adding the following:
Vy ¼ 10 (1.123–1.045)/(1.045–1.0046) ¼ 19.3 mL of
0.15 M NaCl (see Note 6).
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Youakim Saliba and Nassim Farès
Mouse Sacrifice
Male and female adult and aged C57BL/6 mice (3 and 10 months
old) as well as male adult BALB/C mice (3 months old) are used in
the protocol (Janvier Labs, Le Genest-Saint Isle, France). The
animals are kept at a stable temperature (25 C) and humidity
(50 5%) and are exposed to a 12:12-h light-dark cycle. They are
fed ordinary rodent chow, have free access to tap water, and are
acclimatized for at least 1 week under these conditions before the
start of the experiments (see Note 7).
1. Anesthetics: Ketamine hydrochloride 50 mg/mL (RotexMedica, Trittau, Germany) and Xyla, xylazine 2% injection (Interchemie, Waalre, Holland) kept at 4 C
2. Sterile syringe for injections (1 mL)
3. Ethanol 100%
4. Betadine solution, 10% povidone iodine
5. Sterilized surgical scissors
2.5 Pancreas
Removal and Islet
Isolation
1. Three sterile petri dishes (60 mm) kept on ice
2. Serological pipettes (2 mL)
3. Water bath incubator (37 C)
4. One sterilized Erlenmeyer flask (50 mL)
5. Eight sterile conical tubes (4 of 15 mL and 4 of 50 mL)
6. Pure oxygen canister
2.6 Cell Viability
Tests
1. Trypan blue stock solution (0.4%), prepared in 0.81% sodium
chloride and 0.06% potassium phosphate, dibasic
2. Propidium iodide stock solution (1 mg/mL in water)
2.7
Islet Culture
1. RPMI 1640 medium (Lonza, Basel, Switzerland) supplemented with 2 mM L-glutamine.
2. Fetal bovine serum.
3. Prepare a penicillin/streptomycin stock solution (100 concentrated). This solution contains 10,000 penicillin units and
10,000 μg streptomycin per mL. For a 50 mL stock volume,
weigh 33.33 mg of penicillin G sodium salt (1500 U/mg) and
50 mg of streptomycin sulfate and complete with water to the
final volume. The solution is aliquoted and kept at 20 C (see
Note 8).
4. Cell culture plates (6, 24, or 96 wells).
5. 5% CO2 incubator at 37 C.
Mouse Islet Isolation
3
259
Methods
3.1 Mouse Sacrifice
and Pancreas Removal
1. Anesthetize the animals by an intraperitoneal injection of a
mixture of ketamine (75 mg/kg) and xylazine (10 mg/kg).
To make sure of adequate depth of anesthesia, pedal withdrawal reflex (footpad pinch, on two feet) is performed (see
Note 9).
2. When animals are completely nonresponsive to toe pinching,
apply betadine and ethanol on the abdomen in order to keep
the conditions as sterile as possible and to reduce the chance of
hair contamination in the peritoneal cavity during subsequent
steps.
3. Perform a V-shaped abdominal incision starting from the lower
abdomen and extending to the lateral parts of the diaphragm in
order to expose all organs in the peritoneal cavity.
4. Locate the spleen (dark red) which is about 1 cm below the
diaphragm (Fig. 1). The pancreas is surrounded by the stomach, the duodenum and proximal jejunum, and the spleen. In
the mouse, the duodenum wraps around the head of the pancreas that is a soft and diffuse organ as compared with the
human pancreas.
5. Remove the pancreas from the mouse and place it into a petri
dish containing ice cold Tyrode solution. Removal begins by
snipping the common bile duct by scissors and gently detaching the pancreas from the intestines. Continue removing the
pancreas from the stomach and finally the spleen (see Note 10).
Fig. 1 Mouse pancreas anatomical location. The pancreas is surrounded by the
stomach, the duodenum and proximal jejunum, and the spleen. The duodenum
wraps around the head of the pancreas that is a soft and diffuse organ as
compared with the human pancreas. (1) Pancreas, (2) duodenum, (3) stomach,
(4) spleen, (5) liver
260
Youakim Saliba and Nassim Farès
The usual number of islets obtained per pancreas is around
1000; however, if a large number of islets are required for the
assays, use three or four mice and repeat the above steps for
each animal.
6. Gently cut the pancreas into small pieces (2 mm) in ice cold
Tyrode solution. Wash the tissue pieces three times in the same
solution by transferring them in consecutive 60 mm petri
dishes containing 5 mL Tyrode solution. Repeat this three
times or until all blood and fat tissue is removed (fat floats
easily in solution).
3.2 Pancreas
Digestion and Islet
Purification
1. Transfer the pieces into the Erlenmeyer flask and discard the
remaining Tyrode solution. Add the collagenase A solution to
the Erlenmeyer flask (see Note 11) and incubate in a shaking
water bath at 37 C for 1 h at 100 rpm (see Note 12). Supply
the solution by pure oxygen (see Note 13).
2. Stop digestion by transferring the cell suspension into a 50 mL
tube containing an equal volume of ice cold Tyrode solution to
stop the activity of collagenase. Shake the tube vigorously by
hand to mechanically dissociate the digested pancreas and liberate the islets (see Note 14).
3. Leave the cell suspension on ice for 3 min in order to sediment
the islets. Discard the supernatant and add on another 20 mL
ice cold Tyrode solution. Repeat this sedimentation step for
three times (see Note 15).
4. Remove as much liquid as possible from the final pellet and
then purify the cells on the Percoll solution (see Note 16).
Aspirate the pellet with a 1 mL tip and carefully place it on
top of the Percoll solution in a 15 mL tube (see Note 17).
5. Allow the islets to sediment for 5 min (see Note 18). Meanwhile, prepare another two 15 mL tubes with Percoll solution.
6. Collect the pellet from the bottom with a 2 mL pipette and
place it on top of the second Percoll solution. Repeat this again
with the final pellet.
7. Collect the final pellet with a 2 mL pipette and transfer it into a
new 15 mL tube.
8. Add 10 mL RPMI 1640 supplemented with 10% fetal bovine
serum and 2 mM L-glutamine and let islets sediment for 5 min.
9. Discard the supernatant and add another 10 mL RPMI 1640
medium (as above) on the pellet.
10. Gently resuspend the islets.
3.3 Islet Viability
Assessment
Trypan blue exclusion test and propidium iodide are used to assess
islet cell viability. Following the pancreas digestion, perform the
following:
Mouse Islet Isolation
261
1. Transfer 500 μL of islet suspension into a well of a 24-well plate
to perform trypan blue labeling.
2. Add 500 μL trypan blue stock solution to the previous suspension, and incubate for 2 min at room temperature.
3. Count trypan blue-positive and trypan blue-negative islets
under a microscope. Trypan blue-negative islets are regarded
as viable ones. Calculate the percentage of viable islets according to the following formula: total viable cells (unstained)/
total cells (stained and unstained) 100.
4. Transfer another 500 μL of islet suspension into a new well to
perform propidium iodide labeling.
5. Add appropriate volume of propidium iodide stock solution to
reach a final 3 μM concentration in the well, and incubate for
15 min at room temperature.
6. Count fluorescent cells in three view fields under an epifluorescence microscope at 620 nm. Propidium iodide-negative
islets are regarded as viable ones. Calculate the percentage
of viable islets as previously described for trypan blue (see
Note 19).
3.4
Islet Culture
1. Homogenize the cell suspension.
2. Remove 500 μL of the suspension and place in a petri dish to
count the islets under a microscope (see Note 20). The total
number of islets is then obtained by a simple rule of three
calculator.
3. Distribute islets in equal number in 6-, 24-, or 96-well plates as
required, in 2 mL, 500 μL, or 100 μL, respectively
4. Incubate in ambient air with 5% CO2 incubator at 37 C till the
time of experiment (see Note 21).
4
Notes
1. It is important to check the Tyrode solution for any bacterial or
fungi contamination prior to each use. Therefore, it is better to
store sterile solutions in smaller volumes and for short-term
use. Alternatively, add phenol red to the Tyrode solution prior
to filtration, and discard the solution when its color turns
yellow at ph <6.8 for this is a marker for loss of sterility.
2. BSA is added to the collagenase solution in order to protect the
cells from over-digestion. Coating the tissue and vessels with
BSA will prevent the islets from sticking to surfaces they come
into contact with and improve their viability. Adding albumin
increases yield of healthy islets.
262
Youakim Saliba and Nassim Farès
3. It is better to prepare fresh collagenase solution on the day of
isolation, and keep it on ice till the time of tissue removal.
4. In order to make Percoll isotonic and suitable for cell separation, the osmolality of undiluted Percoll must first be adjusted
with saline or cell culture medium. Adding 9 parts of Percoll to
1 part of 1.5 M NaCl or 10 concentrated cell culture medium
is a simple way of preparing an SIP solution.
5. It has been shown that rodent pancreatic exocrine tissue possesses a density lower than 1.045 g/mL, whereas islets typically
have a higher density (1.065–1.07 g/mL) [19, 20].
6. Phenol red can be added to the final Percoll solution in order to
differentiate between the Percoll solution and the pancreatic
tissue mixture that will be layered over it, more easily observing
the interface between the cell suspension and the Percoll density medium.
7. The present study was approved by the ethical committee of
Saint Joseph University. The protocols were designed according to the Guiding Principles in the Care and Use of Animals
approved by the Council of the American Physiological Society
and were in adherence to the Guide for the Care and Use of
Laboratory Animals published by the US National Institutes of
Health (NIH Publication no. 85-23, revised 1996) and according to the European Parliament Directive 2010/63 EU.
8. Penicillin and streptomycin are both light sensitive and should
be kept in dark wrapped in aluminum foil.
9. Monitoring the depth of anesthesia can also be done by pinching the base of the tail. However, the latter is less reliable since
responses can be sometimes noticed with toe-pinching even
with the absence of a tail response.
10. It is important to avoid intestinal rupture in order to reduce
the risk of contamination of the cell culture by intestinal bacteria. If the mouse is aged, it becomes easy to confuse between
abdominal fat and pancreatic tissue. However, when the pancreas is removed and put in Tyrode solution, fat tissue easily
floats and is discarded.
11. 10 mL volume is ideal for a 50 mL Erlenmeyer flask. This
results in optimal shaking and tissue dispersion during the
incubation. If more volume is added, the solution will not
shake well, and tissue will remain placid resulting in overdigestion of the outer layers of cells and so lower yield.
12. There are significant differences in the specific activity of different lots of collagenase A. Therefore it is important to do
pilot studies with each lot of collagenase A to determine the
optimal incubation time for tissue digestion. Since collagenase
Mouse Islet Isolation
263
A possesses low activity, tissue digestion typically takes more
than 50 min and can differ significantly with different batches.
13. Gassing with pure oxygen increases yield by attenuating anaerobic metabolism.
14. The resulting solution has a viscous consistency and is free of
large pieces of pancreatic tissue. If, for a reason, a significant
amount of large tissue remains, pellet the tissue suspension,
add on ice cold Tyrode solution, and leave onto ice till another
collagenase solution is prepared. Re-incubate the pellet with
the enzyme while shaking at 37 C as previously described for
an additional 10–15 min.
15. Islets are significantly larger and heavier than exocrine tissue
and will sediment after just 1 min. Leaving the cell suspension
for 3 min will maximize the islet number obtained. If sedimentation is left for a longer time, islets yield will be higher, but
more and more exocrine tissue will be present with the islets
pellet. Sedimentation time depends on the required assays and
thus on the necessary number of islets. If high purity is not an
issue, the damaging effect of exocrine tissue-secreted enzymes
on islet cells has, however, to be taken into consideration.
16. Removing as much as possible of the supernatant ensures that
the Percoll solution is not diluted when the pellet is poured
onto it.
17. When more than three pancreases are digested, attention is
required not to overload the Percoll solution and thus assure
a proper separation of islets from exocrine tissue. In this case,
use another tube with a Percoll solution for the additional
pancreases.
18. Sedimentation time is a compromise between yield and purity
of islets. Shorter sedimentation times result in lower yield but
higher purity, while longer times result in the opposite
outcome.
19. For long-term cultures, 3-(4,5-dimethylthiazol-2-yl)-2,5diphenyltetrazolium bromide (MTT) assay is also useful.
20. When viewed under a light microscope, islets appear spherical
with a typical golden brown color as compared to the relatively
transparent exocrine tissue. They are 50–250 μm in diameter.
Cells protruding from the surface of the islets are a sign of
decreased overall health (Fig. 2). A dark center is sometimes
visible in larger islets due to hypoxia.
21. Trypsin inhibitors from soybean can be added to the cell culture medium in order to block the remnant protease activity of
contaminating exocrine cells’ secretions.
264
Youakim Saliba and Nassim Farès
Fig. 2 Healthy versus over-digested mouse islets microphotographies. (a) Healthy well-rounded islets. (b)
Over-digested islets with cells protruding from the surface. Magnification, 400. Scale bar, 40 μm
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NM (1996) High yield of rodent islets with
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Chapter 19
Identification and In Vitro Expansion of Adult Hepatocyte
Progenitors from Chronically Injured Livers
Naoki Tanimizu
Abstract
The liver performs a number of physiologically important functions. Hepatocytes are the liver parenchymal
cells performing most of those functions. Therefore, it is important to recover functional hepatocytes after
hepatic injury and prepare a mass of hepatocytes for regenerative medicine. We have found that mature
hepatocytes dedifferentiate to hepatocyte progenitors in chronically injured mouse liver. Those hepatocyte
progenitors can be isolated as CD24+EpCAM cells from the CD31CD45 fraction, which clonally
proliferate and efficiently re-differentiate to functional hepatocytes both in vitro and in vivo. Here, I
describe the methods to isolate hepatocyte progenitors from chronically injured liver, to expand them
in vitro, and to induce differentiation into functional hepatocytes.
Key words Liver, Chronic liver injury, Hepatocyte, Hepatocyte progenitors, Liver stem/progenitor
cells, Fluorescence-activated cell sorting
1
Introduction
Healthy epithelial tissues and organs contain tissue-specific stem/
progenitor cells, which continuously supply multiple types of functionally differentiated cells throughout life. Liver stem/progenitor
cells (LPCs) are defined as bipotential cells differentiating into two
types of liver epithelial cells, hepatocytes and cholangiocytes,
whereas hepatocyte progenitors are committed, hepatocyte lineage
restricted progenitors. LPCs have been isolated from normal and
injured livers based on the expression of surface antigens including
epithelial cell adhesion molecule (EpCAM), CD13, and CD133
[1]. On the other hand, recent results demonstrated that mature
hepatocytes (MHs) near the portal vein are dedifferentiate into
hepatocyte progenitors when the liver suffer chronic injuries
induced by 3,5-diethoxycarbonyl-1,4-dihydro-collidine (DDC)
diet or after bile duct ligation (BDL) [2]. Those dedifferentiated
hepatocytes are referred as biphenotypic hepatocytes since they
express some cholangiocyte markers including osteopontin and a
Ivan Bertoncello (ed.), Mouse Cell Culture: Methods and Protocols, Methods in Molecular Biology, vol. 1940,
https://doi.org/10.1007/978-1-4939-9086-3_19, © Springer Science+Business Media, LLC, part of Springer Nature 2019
267
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Naoki Tanimizu
transcription factor Sry HMG box protein 9 (SOX9) [3]. Biphenotypic hepatocytes emerged in chronically injured liver can be
isolated as SOX9+EpCAM cells from SOX9-EGFP mice fed with
DDC diet or after BDL. SOX9+EpCAM hepatocytes are further
fractionated into CD24 and CD24+ cells, and the latter have the
higher capability [4]. Furthermore, CD24+EpCAM cells isolated
from the CD31CD45 liver cell fraction of the wild-type mice fed
with DDC diet are mostly identical to SOX9+CD24+EpCAM
cells.
CD24+EpCAM
hepatocytes
can
expand
and
re-differentiate to functional hepatocytes both in vitro and
in vivo. Therefore, they can be a good source for producing a
large quantity of hepatocytes for basic research and for regenerative
medicine.
2
Materials
2.1 Mouse Injury
Model and Cell
Isolation
1. 3,5-Diethoxycarbonyl-1,4-dihydro-collidine (DDC) diet: Mix
DDC and MF powder (w:w ¼ 1:100) and mold them into
pellets (12 mm diameter) by Oriental Yeast Co. Ltd. (Tokyo,
Japan).
2. Isoflurane.
3. Calcium, magnesium-free Hanks’ balanced salt solution
(HBSS).
4. Pre-perfusion solution: 0.5 mM EGTA in HBSS.
5. Butterfly needle of 23 gauge.
6. Collagenase: Dissolve in HBSS at 1 mg/mL for perfusion and
2 mg/mL for digesting the remaining tissue after collagenase
perfusion.
7. Peristaltic pump.
8. Autoclaved 250 μm nylon mesh.
9. 70 μm cell strainer.
10. L-15 medium.
11. DNase I: Make 1 mg/mL solution and store at 4 C or at
30 C for a long term.
12. Hyaluronidase: Make 75,000 units/50 μL solution and store
at 30 C.
2.2
Cell Sorting
1. Antibodies:
PE-conjugated
anti-mouse
CD24,
APC-conjugated anti-mouse EpCAM, APC-Cy7-conjugated
anti-mouse CD45, and PE-Cy7-conjugated anti-CD31.
2. Wash buffer: Add 2% FBS to PBS. Store at 4 C.
3. Propidium iodide (PI) (1 mg/mL).
Mouse Adult Hepatocyte Progenitors
269
4. Fluorescence-activated cell sorter (FACS): FACS Aria
equipped (BD biosciences) with three lasers (UV, 488 nm,
and 633 nm) or an equivalent system.
2.3
Cell Culture
1. Growth factor-reduced Matrigel® (MG): Thaw the bottle of
MG on ice and make aliquots in 1.5 mL tubes. Freeze those
tubes in liquid nitrogen and store them at 30 C. Thaw MG
in a tube on ice before use (see Note 1).
2. Laminin solution: Thaw laminin-111 in a glass bottle at 4 C.
Transfer it to a 1.5 mL tube and store at 4 C. Dilute it in PBS
at 10 μg/mL. Add 1 mL diluted solution to 35 mm plate. Wait
about 1 h and remove the solution before plating cells (see
Note 2).
3. Gelatin solution: Autoclave PBS containing 0.1% gelatin. Add
300 μL of gelatin solution to each well of 24-well plate. Aspirate the gelatin solution and wash wells with PBS once.
4. Basal medium: Prepare Dulbecco’s Modified Eagle’s Medium
and Ham’s F-12 Nutrient Mixture (DMEM/F-12) medium
supplemented with 10% fetal bovine serum, 10 mM nicotinamide, and 10 μg/mL gentamicin.
5. Growth medium: Add 1 insulin/transferrin/selenium (ITS),
1 107 M dexamethasone (Dex), 10 ng/mL epidermal
growth factor (EGF), and 10 ng/mL hepatocyte growth factor
(HGF) (R&D systems) to the basal medium.
6. Differentiation medium: Add 1 ITS, 1 107 M Dex, and
1% dimethyl sulfoxide (DMSO) to the basal medium. Add
10 ng/mL oncostatin M (OSM) or 5% MG before use.
7. Rho-associated coiled-coil forming kinase (ROCK) inhibitor:
50 mM Y27632 in autoclaved distilled water.
3
Methods
3.1
Cell Isolation
1. Feed C57BL6 mice with DDC diet for 2 weeks. Anesthetize a
mouse by isoflurane. Open the abdomen and expose the portal
vein. Insert a butterfly needle into the portal vein and fix it by a
vascular clump. Inject 25 mL of pre-perfusion solution by
using a peristaltic pump at 6 mL/min. At the end of
pre-perfusion, dissolve 50 mg collagenase in 50 mL perfusion
solution. Inject the perfusion solution containing collagenase
by using a peristaltic pump at 3 mL/min (see Note 3).
2. Isolate the liver and place it on a 10 cm dish. Add 20 mL of
HBSS. Tear off the liver capsule with forceps. Hold the liver
with forceps, and scrape off hepatocytes from undigested liver
tissue with another forceps until the biliary tissue is thoroughly
exposed. (Use undigested tissue for further enzymatic
270
Naoki Tanimizu
digestion in step 4.) Pass the cell suspension through a 250 μm
nylon mesh and then a 70 μm cell strainer. Centrifuge the cell
suspension at 50 g for 1 min. Transfer the supernatant to a
new 50 mL tube and centrifuge at 350 g for 4 min. Resuspend
cells in 1 mL of basal medium (“non-parenchymal fraction”).
3. Mince the undigested tissue into small pieces (<1 mm cubic)
after collagenase perfusion. Suspend tissue pieces in 15 mL of
L-15 medium and transfer them into a 30 mL beaker. Add
30 mg collagenase, 50 μL of DNase I solution, and 25 μL of
hyaluronidase. Stir the solution vigorously at 37 C for 40 min.
Add 10 mL of basal medium to the cell suspension to stop
enzymatic digestion.
4. Pass the cell suspension through a 250 μm nylon mesh and
then a 70 μm cell strainer. Centrifugation at 50 g for 1 min.
Then, centrifuge the supernatant at 350 g for 4 min to collect
dissociated cells. Resuspend cells in 1 mL of basal medium
(“cholangiocyte fraction”).
5. Combine the “non-parenchymal” and “cholangiocyte” fraction into a new 15 mL tube (see Note 4).
3.2
Cell Sorting
1. Centrifuge the cell suspension at 350 g for 4 min. Resuspend
cells (no more than 1 107 cells) in 100 μL of basal medium
(see Note 5). Add 1 μL of anti-CD16/CD32 antibody and
incubate at 4 C for 20 min to avoid non-specific binding of
antibodies by masking Fcγ receptors. Add ice-cold wash buffer
and centrifuge at 350 g for 4 min.
2. Resuspend cells in 100 μL of basal medium. Add 1 μL of
PE-conjugated anti-CD24, 1 μL of APC-conjugated antiEpCAM, 1 μL of APC-Cy7-conjugated anti-CD45, and 1 μL
of PE-Cy7-conjugated anti-CD31. For the isotype control,
add PE-conjugated rat IgG, APC-conjugated rat IgG,
APC-Cy7-conjugated rat IgG, and PE-Cy7-conjugated rat
IgG. Incubate cells with antibodies at 4 C for 20 min. Add
2 mL of ice-cold wash buffer and centrifuge at 350 g for
4 min (see Note 6).
3. Resuspend cells in 300 μL of basal medium containing 1 μg/
mL PI. Pass through 40 μm cell strainer before analysis on
FACS.
4. Select CD31CD45 cells in PI singlet cells by gating on a
FACS density plot. Identify and collect EpCAMCD24+ cells
in the CD31CD45 fraction into a 5 mL tube (Fig. 1) (see
Note 7).
5. Transfer the cell suspension into a 15 mL tube. Centrifuge at
350 g for 8 min. Resuspend cells and add 7 mL of basal
medium. Centrifuge at 350 g for 8 min. Resuspend cells in
1 mL of basal medium and count the number of cells.
Mouse Adult Hepatocyte Progenitors
271
Fig. 1 Isolation of EpCAMCD24+ hepatocyte progenitors. CD31+ endothelial cells and CD45+ hematopoietic
cells are eliminated from PI singlet live cells by gating. EpCAMCD24+ hepatocyte progenitors are identified
in the CD31CD45 fraction
3.3
Cell Culture
1. Resuspend cells in growth medium containing 20 μM Y27632.
Plate 5000 cells in 35 mm dish coated with laminin 111 (see
Notes 8 and 9). For examining clonal proliferation and differentiation potential, fix cells at day 7 in 4% PFA at 4 C for
15 min and use for immunofluorescence analysis.
2. For inducing hepatocyte differentiation, plate 5000
EpCAMCD24+ cells in a well of 24-well plate coated with
gelatin. After cells become confluent, incubate cells in the
differentiation medium containing OSM for 2 days.
3. Replace the medium with one containing 5% MG in order to
induce hepatic maturation. Keep the culture for additional
3 days for analyzing gene and protein expressions and hepatic
functions to examine hepatocyte differentiation (Fig. 2) (see
Notes 10 and 11).
4
Notes
1. If an aliquot contains 500 μL MG, incubate the tube on ice
for more than 2 h ahead of the experiment to thaw the
solution.
2. The diluted laminin solution can be reused up to three times to
coat tissue culture dishes. We usually recover the laminin solution and store it in a 15 mL tube at 4 C.
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Naoki Tanimizu
Fig. 2 Morphological changes during hepatocyte differentiation. EpCAMCD24+ cells proliferate and form a
monolayer, though the cell-cell boundaries are not clearly recognized and atypical nuclear shapes are
observed (a). After sequential treatment of OSM and MG, EpCAMCD24+ cells show polygonal cellular
shapes, round nuclei, and dark cytoplasm under a phase-contrast microscope (b). Binucleated cells, a feature
of mature hepatocytes, are evident (arrowheads in panel b)
3. All animal experiments were approved by the Sapporo Medical
University Institutional Animal Care and Use Committee and
were carried out under the institutional guidelines for ethical
animal use.
4. If the color of pellet is reddish, resuspend the pellet in 5 mL of
hemolysis buffer (15 mM Tris(hydroxymethyl)aminomethane,
100 mM NH4Cl), and incubate it on ice for 5 min. Add 5-7 mL
of basal medium and pass through 70 μm cell strainer. Collect
cells by centrifugation at 350 g for 4 min.
5. If the number of cells is more than 1 107 cells, cells are
resuspended at 1 108 cells/mL and added with antibodies
at 10 μL/mL.
6. CD31CD45EpCAMCD24+ cells are less than 1% of the
total cells. We usually prepare a tube added with PE-conjugated
rat IgG, APC-conjugated rat IgG, APC-Cy7-anti-CD45, and
PE-Cy7-anti-CD31. By comparing this, EpCAMCD24+ cells
can be clearly recognized in bivariate FACS dot plots of the
sample as shown in Fig. 1.
7. If SOX9-EGFP mice are available, CD31CD45GFP+ are
further divided into four fractions based on expression of
CD24 and EpCAM. GFP+EpCAM fraction contains hepatocyte progenitors, whereas GFP+EpCAM+ fraction contains
cholangiocytes and expanded ductular cells. Among
GFP+EpCAM cells, proliferative hepatocyte progenitors are
enriched in CD24+ cells, which are identical to
EpCAMCD24+ cells isolated from the wild-type mice fed
with DDC diet.
Mouse Adult Hepatocyte Progenitors
273
8. For cell transplantation, EpCAMCD24+ cells are clonally
maintained for a month and then infected with lentivirus containing GFP expression cassette. GFP+ cells are isolated by
FACS and expanded for 4 weeks before transplantation.
9. For isolation of progenitor clones, clonal culture is kept about a
month. Each colony is surrounded with a cloning ring, treated
with trypsin, and replated onto a laminin-coated dish.
10. The morphological features of hepatocytes are the roundshaped nucleus and the sharp contrast between the nucleus
and the cytosol: under a phase-contrast microscope, the dark
cytosol is evident because of enriched granules and organelles
(Fig. 2). Usually, hepatocyte progenitors show such
hepatocyte-like morphology by incubation with MG for
2-3 days. If not, replace medium with the fresh basal medium
containing ITS, Dex, and DMSO, and keep the culture for
additional 2-3 days to induce further differentiation.
11. Cultured biphenotypic hepatocytes differentiate to functional
hepatocytes secreting albumin and possessing CYP3A4 activity
in the presence of OSM and MG. On the other hand, they
inefficiently form cysts (spheroids with a central lumen) in
three-dimensional culture, which indicates that they do not
have potential to differentiate to cholangiocytes.
Acknowledgments
I thank Dr. Toshihiro Mitaka and Dr. Norihisa Ichinohe for their
helpful discussion. I also thank Ms. Yumiko Tsukamoto and
Ms. Minako Kuwano for their technical assistances. This work is
supported by the Ministry of Education, Culture, Sports, Science
and Technology, Japan, Grants-in-Aid for Scientific Research
(C) (25460271, 16 K08716), and Grants-in-Aid for Scientific
Research on Innovative Areas “Stem Cell Aging and Disease”
(17H05653).
References
1. Miyajima A, Tanaka M, Itoh T (2014) Stem
Progenitor cells in liver development, homeostasis, regeneration, and reprogramming. Cell Stem
Cell 14:261–274
2. Yanger K, Zong Y, Maggs LR et al (2013)
Robust cellular reprogramming occurs spontaneously during liver regeneration. Genes Dev
27:719–724
3. Tanimizu N, Nishikawa Y, Ichinohe N et al
(2014) Sry HMG box protein 9-positive
(Sox9+) epithelial cell adhesion moleculenegative (EpCAM) biphenotypic cells derived
from hepatocytes are involved in mouse liver
regeneration. J Biol Chem 289:7589–7598
4. Tanimizu N, Ichihohe N, Yamamoto M et al
(2017) Progressive induction of hepatocyte progenitor cells in chronically injured liver. Sci Rep
7:39990
Chapter 20
The Preparation of Decellularized Mouse Lung Matrix
Scaffolds for Analysis of Lung Regenerative Cell Potential
Deniz A. Bölükbas, Martina M. De Santis, Hani N. Alsafadi, Ali Doryab,
and Darcy E. Wagner
Abstract
Lung transplantation is the only option for patients with end-stage lung disease, but there is a shortage of
available lung donors. Furthermore, efficiency of lung transplantation has been limited due to primary graft
dysfunction. Recent mouse models mimicking lung disease in humans have allowed for deepening our
understanding of disease pathomechanisms. Moreover, new techniques such as decellularization and
recellularization have opened up new possibilities to contribute to our understanding of the regenerative
mechanisms involved in the lung. Stripping the lung of its native cells allows for unprecedented analyses of
extracellular matrix and sets a physiologic platform to study the regenerative potential of seeded cells. A
comprehensive understanding of the molecular pathways involved for lung development and regeneration
in mouse models can be translated to regeneration strategies in higher organisms, including humans. Here
we describe and discuss several techniques used for murine lung de- and recellularization, methods for
evaluation of efficacy including histology, protein/RNA isolation at the whole lung, as well as lung slices
level.
Key words Biomaterial, Decellularization, Recellularization, Lung, Precision cut lung slices, Scaffold,
Tissue engineering
1
Introduction
The global prevalence of chronic lung diseases is increasing, posing
serious threats to human health [1, 2]. Despite recent advancements, curative therapies for end-stage lung diseases are still missing [3]. Lung transplantation remains the only option at end-stage
disease, but the current clinical demand exceeds the number of
available lungs [4]. 5-year survival rates after lung transplantation
have remained around 50% due to severe primary graft dysfunction
and are low when compared to other solid organ transplantations
[5, 6]. Therefore, novel solutions are urgently needed to address
these issues for patients with chronic lung disease.
Ivan Bertoncello (ed.), Mouse Cell Culture: Methods and Protocols, Methods in Molecular Biology, vol. 1940,
https://doi.org/10.1007/978-1-4939-9086-3_20, © Springer Science+Business Media, LLC, part of Springer Nature 2019
275
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Deniz A. Bölükbas et al.
Different experimental animal models, including those utilizing lineage tracing of candidate stem and progenitor cell populations, have been developed to better understand disease
pathomechanisms and to evaluate the regenerative potential of
the lung in diseases such as chronic obstructive pulmonary disease
[7], pulmonary fibrosis [8], and bronchiolitis obliterans syndrome
[9]. However, the ability to study cell-matrix interactions in these
models is challenging. Ex vivo models derived from rodent tissue
have been more recently utilized to study lung repair and regeneration and cell-matrix interactions [3]. Such models are primarily
based on cultivation of stem or progenitor cells on biologically or
synthetically derived scaffolds. While three-dimensional culture of
cells in biologically derived materials such as Matrigel and rat-tail
type I collagen has helped to elucidate lung stem and progenitor
cell behavior, including cell-matrix interactions [10–16], Matrigel
is derived from a sarcoma cell line and is known to contain a variety
of growth factors also known to be involved in cancer. Further,
rat-tail type I collagen incompletely mimics the in vivo microenvironment and is also known to contain a variety of other factors
[17]. Thus, alternative models which more closely mimic the
in vivo lung microenvironment are useful for validating the regenerative potential of different cell populations. One way to generate
more physiologic lung microenvironments is through the use of
acellular biologic lung scaffolds which can be derived through a
process termed as decellularization. This can be achieved by perfusion of select solutions through the airways or the vasculature of the
lungs from a variety of species, such as murine, rat, porcine, nonhuman primate, and human, to remove the native cells while retaining the general architecture of the lung and the majority of
extracellular matrix components [3, 18, 19]. Repopulation and
maturation of autologous cells in these scaffolds, termed as recellularization, can aid in understanding the mechanisms involved in
lung regeneration and ultimately be used to obtain functional lung
tissue that would minimize the need for immunosuppressant therapies posttransplantation [20]. Previous studies have used a variety
of different cell types and culturing media for recellularizing acellular murine lung scaffolds. Examples of cell types inoculated to date
include mesenchymal stromal cells (MSCs) derived from mouse
bone marrow or C10 murine lung epithelial cells [21–25], embryonic stem cells derived from cKIT+/CXCR4+ endodermal cells
[26] and A9 murine transformed subcutaneous fibroblasts [27],
and fetal mouse lung cells derived from E17 lung tissue homogenates [28], E14tg2a, or Nkx2-1+ versus undifferentiated embryonic stem cells (ESCs) [29, 30]. However, functional lung tissue
has yet to be obtained and will likely require multiple cell types and
careful control of growth factors (or small molecule mimics) using
advanced bioreactors.
Acellular Lung Scaffolds for Regenerative Potential
277
Alternatively, the use of decellularized lungs offers the possibility to better understand cell-cell and cell-matrix interactions and
their role in pathomechanisms in a more controlled manner. It has
been previously shown that acellular scaffolds derived from diseased
or aged murine, rodent, and human lungs retain their pathological
composition and architecture to a large degree after decellularization [18, 21, 31–37]. In particular, acellular scaffolds derived from
diseased lungs provide an opportunity to study cell-extracellular
matrix interactions in a more in vivo-like setting compared to
traditional cell culture on tissue plastic or on extracellular-matrixcoated tissue culture dishes. Interestingly, cells seeded onto acellular scaffolds derived from pathological or aged lungs adopted disease or aging phenotypes compared to cells seeded onto control
lung scaffolds [21, 35, 38, 39]. Acellular scaffolds also set the stage
for recellularization analyses and more physiologic assessment of
regenerative potential, which are difficult to achieve using other
methods or require specialized microscope settings (e.g., intravital
microscopy) to achieve in vivo. For instance, lineage-tracing technology and live imaging of select cell types and/or select cell
combinations could be employed in murine lung recellularization
models to identify key players in disease remodeling or
regeneration.
In this chapter, we describe the preparation and use of acellular
murine lungs to study the regenerative potential of the lung (cells)
ex vivo (see Fig. 1). Additionally, protocols for monitoring recellularization of the bioengineered scaffolds are described.
2
Materials
All procedures below should be carried out in a sterile environment
(e.g., cell culture hood) with sterile tools (sterilized, i.e., autoclaved
or rinsed with 70% ethanol, forceps, scissors, silk sutures, petri
dishes, beakers, waste containers, syringes, 19-gauge needles, and
cannulas).
2.1 Solutions
and Materials
for Murine Lung
Extraction
All of the solutions used in Subheadings 2.1–2.3 should be
prepared with deionized or ultrapure water and be sterilized before
use.
1. Mice (see Note 1).
2. 70% (v/v) ethanol
3. Loupe microscope with light (Leica M80 8:1 manual zoom
coupled with KL200 LED).
4. Ventilator (Harvard Apparatus, MiniVent 845).
5. 19-gauge needles with blunt ends and polyurethane cannulas
with 0.9 mm diameter (Instech, C20PU-MJV1451; see
Note 2).
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Fig. 1 Murine lung de- and recellularization approaches. De- and recellularization of the lung can be
accomplished by different strategies: (a, b) at the whole lung level through the airways or vascular system
by perfusion of respective cells in bioreactor systems or (c–g) by seeding individual precision cut lung slices
(depicted as rings in a, b) of acellular lung scaffolds with cell preparations in tissue culture plates
6. 2:1 (v/v) mix of ketamine (100 mg/mL) and xylazine hydrochloride (20 mg/mL) (see Note 3).
7. Phosphate-buffered saline (PBS).
2.2 Solutions
and Materials
for Decellularization
1. 100 U/mL penicillin and 100 μg/mL streptomycin in distilled
water (henceforth called DI solution).
2. 0.1% Triton X-100 solution supplemented with 100 U/mL
penicillin and 100 μg/mL streptomycin.
3. 2% (w/v) sodium deoxycholate solution (SDC).
4. 1 M NaCl solution supplemented with 100 U/mL penicillin
and 100 μg/mL streptomycin.
5. DNase solution (i.e., 30 μg/mL porcine pancreatic DNase,
1.3 mM MgSO4, 2 mM CaCl2, 100 U/mL penicillin, and
100 μg/mL streptomycin) freshly prepared or taken from aliquots stored at 20 C.
6. PBS storage solution (i.e., 100 U/mL penicillin and 100 μg/
mL streptomycin, 50 mg/L gentamicin, and 2.5 μg/mL
amphotericin B in PBS without Ca2+ and Mg2+).
Acellular Lung Scaffolds for Regenerative Potential
2.3 Cells
and Materials
for Recellularization
2.3.1 Whole Lung
Recellularization
279
1. An ex vivo perfusion/ventilation system built from the elements below.
2. Peristaltic pump capable of a flow rate of 1 mL/min (Cole
Parmer #EW-73160-20).
3. Screw caps with two hose connections (Aldrich, Duran® GL
45 #Z680419).
4. 100 mL graduated laboratory bottles (Aldrich, Duran®
#Z232076).
5. Silicone tubing with 8 mm inner diameter (Witeg #9316012).
6. Four-way stopcocks with Luer connections (Cole Parmer
#EW-30600-04).
7. Water bath filled with water kept at 37 C.
8. Ventilator (Harvard Apparatus, MiniVent 845).
2.3.2 Physiologic or
Direct PCLS
Recellularization
1. Low gelling temperature agarose (Sigma-Aldrich A2576; see
Note 4).
2. Vibratome (Hyrax V55, Zeiss, Germany; see Note 5).
3. Cyanoacrylate glue.
4. 24-well plates for cell culture
5. Incubator at 37 C with 5% CO2 in humidified air.
6. Biopsy punch (optional; see Note 6).
2.4
Lung Histology
1. 4% paraformaldehyde (PFA).
2. Paraffin.
3. Microtome.
4. Xylene.
5. 70%, 90%, and 100% ethanol series.
6. Water bath filled with water kept around 45 C.
7. Hot plate kept around 65 C.
8. Adhesive glass microscopic slides.
9. Histological stain as needed (for common examples; see
Table 1).
10. Cover slips and mounting medium (e.g., Entellan for brightfield staining, DAKO fluorescent mounting medium for fluorescent staining).
2.5 Materials
for Tissue
Homogenization
1. Liquid nitrogen.
2. Tissue homogenizer (see Note 7).
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Table 1
Common staining examples for histological validation of de- or recellularized lungs
Name
Stains
Type
Hematoxylin and
eosin (H&E)
Hematoxylin stains the nuclei violet, while eosin stains the other
structures in pink/red
Bright-field
Verhoeff-van Gieson Nuclei are stained in blue, elastic fibers in black, collagen in red,
stain (EVG)
and the other tissue elements in yellow
Bright-field
Masson’s trichrome
stain
Nuclei are stained in black/blue, collagen in blue, and the muscle Bright-field
fibers in red
Alcian blue
Nuclei are stained in red, mucin in blue, and the background in
pink
Bright-field
DAPI, Hoechst
Cell nuclei (excitation/emission 350/470 nm)
Fluorescence
Phalloidin
Actin (various absorption)
Fluorescence
2.6 Solutions
for Protein Analysis
1. RIPA lysis buffer (Thermo Scientific #89900).
2. cOmplete™, Mini Protease Inhibitor Cocktail (Sigma
#11836153001).
3. PhosSTOP™ phosphatase inhibitor cocktail tablets (Sigma
#4906837001).
2.7 Solutions
and Materials for RNA
Analysis
3
1. peqGold Total RNA Kit (VWR International #732-2868).
2. Proteinase K (Thermo Scientific #11501515).
3. tris(2-carboxyethyl)phosphine (TCEP; Merck #580561).
Methods
3.1 Murine Lung
Extraction
and Preparation
of the Heart-Lung
Block
1. Anesthetize the mice by injecting ~100 μL of 2:1 (v/v) ketamine/xylazine hydrochloride mixture intraperitoneally.
2. Wait for circa 5 min until the mouse becomes unconscious.
Validate proper anesthetization by checking toe pinch reflex
before starting the surgery.
3. Place the mouse on the operating table; fix in supine position
with tape or needles holding the arms, legs, and tail in an
outstretched position; and spray with 70% ethanol for sterilization (see Fig. 2a).
4. Using surgical blunt scissors, enter peritoneal cavity by a midline incision.
5. Continue the incision cranially until the trachea is exposed.
Make sure the thoracic cavity is not ruptured.
6. Dissect posterior to the trachea and place the suture posterior
to the trachea.
Acellular Lung Scaffolds for Regenerative Potential
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Fig. 2 Murine lung extraction. (a) Incising cranially through the rib cage to expose the thoracic cavity and lungs
of the mouse. Representation of a heart-lung block ready to be processed for decellularization from (b) dorsal
and (c) ventral view
7. Make a small anterior incision between the cartilaginous rings
of the trachea, making sure not to cut through the entire
trachea, and cannulate the trachea by inserting the 19 gauge
needle with blunt ends (described in Subheading 2.1, item 5)
and securing it with the suture (see Note 8).
8. Enter the thoracic cavity by dissecting the diaphragm and
incising through the sternum cranially. Make sure the lungs
are not ruptured (Fig. 2a).
9. Retract the ribs to two sides to expose the thoracic cavity.
10. Dissect the thymus. Make sure the large vessels of the heart are
not damaged.
11. Move the abdominal organs to one side and make an incision at
the inferior vena cava for euthanasia.
12. Slide a suture posterior to the pulmonary artery (PA) in preparation for securing the polyurethane PA cannula.
13. Slide a suture posterior to the aorta in preparation for securing
the polyurethane aorta cannula.
14. Perfuse the lungs by injecting the right ventricle with
10–20 mL of PBS to wash away blood and blood clots from
the lung microvasculature and take care not to overinflate the
lungs (i.e., stop injecting when you feel pressure on the
syringe). Observe the color change in the lung from pink to
white (Fig. 2b, c).
15. Cannulate the PA and the aorta. Secure each with silk sutures.
16. Dissect the heart-lung block from the thoracic cavity carefully
using blunt scissors.
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3.2 Murine Lung
Decellularization
1. After removing the heart-lung block from the thoracic cavity,
incubate the lungs in the DI solution on ice.
2. Perfuse the lungs with 15 mL DI solution through the trachea
and 15 mL DI solution through the right ventricle using a
19-gauge needle. Do the first rinse very carefully by pausing
when injecting to allow solution to come out as lung recoils.
3. Perfuse the lungs with 3 mL 0.1% Triton X-100 solution
through the trachea and 3 mL 0.1% Triton X-100 solution
through the right ventricle using a 19-gauge needle
(Fig. 3a, b).
4. Incubate submerged in 0.1% Triton X-100 solution for 24 h at
4 C (Fig. 3c).
Fig. 3 Murine lung decellularization. Administration of respective decellularization and washing solutions into the lungs via the (a) trachea and (b) the heart and
(c) incubating the heart-lung block for 24 h at 4 C. See Subheading 3.2 for
sequences and volumes of solutions to be instilled on each day of the protocol
and incubation periods. (d) Following each decellularization step, administration
of DI solution into the murine lungs through both the trachea and the heart is
described. In the final step, PBS storage solution should be instilled through both
the trachea and the heart. The final scaffold should be stored submerged in PBS
storage solution at 4 C
Acellular Lung Scaffolds for Regenerative Potential
283
5. Remove the lungs from 0.1% Triton X-100 solution and perfuse with the DI solution as in step 2 (Fig. 3d).
6. Perfuse the lungs with 3 mL 2% SDC through the trachea and
3 mL 2% SDC through the right ventricle using a 19-gauge
needle.
7. Incubate in 2% SDC for 24 h at 4 C.
8. Remove the lungs from 2% SDC solution and perfuse the lungs
with the DI solution as in step 2.
9. Perfuse the lungs with 3 mL NaCl solution through the trachea
and 3 mL NaCl solution through the right ventricle using a
19-gauge needle.
10. Incubate in NaCl solution for 1 h at room temperature.
11. Remove the lungs from NaCl solution and perfuse the lungs
with the DI solution as in step 2.
12. Perfuse the lungs with 3 mL DNase solution through the
trachea and 3 mL DNase solution through the right ventricle
using a 19-gauge needle.
13. Incubate in DNase solution for 1 h at room temperature.
14. Remove the lungs from DNase solution and perfuse the lungs
as in step 2, but this time using PBS storage solution instead of
the DI solution.
15. Store the acellular lungs in PBS storage solution at 4 C (or remove and fix lobes immediately for residual DNA or histological assessment, respectively; see Note 9).
3.3 Generation
of Acellular PCLS
Alternatively, PCLS can also be prepared from decellularized
murine lungs and be utilized as scaffolds for recellularization studies. The following steps are to be followed for generating acellular
PCLS:
1. Pre-warm a water bath to 41 C.
2. Prepare a 3% low gelling temperature agarose solution in PBS
by warming up the mixture in a microwave for a few minutes
until it boils.
3. In parallel, place PBS in 20 C to prepare ice-cold PBS and
ensure that it does not freeze.
4. Place the 3% agarose solution in the 41 C water bath and allow
to equilibrate for at least 20 min.
5. Remove the ice-cold PBS and pour into a sterile 50 mL beaker
in a tissue culture hood.
6. Use forceps to suspend the acellular lung by the tracheal cannula and submerge in the ice-cold PBS.
7. Slowly administer the agarose solution into the lung scaffold by
injecting through the trachea cannula.
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8. Ligate the trachea with sutures to prevent the agarose from
escaping through the trachea cannula.
9. Place the acellular lung scaffold filled with agarose in a petri
dish and place on ice (or at 4 C) for solidification of the
agarose.
10. Separate the individual mouse lobes and mount them using
cyanoacrylate glue onto the cutting surface. Cut with the
vibratome system to a thickness of 300 μm using a speed of
~1.2 mm/s, a frequency of 100 Hz, and an amplitude of
1 mm [21].
3.4 Confirmation
of Decellularization
and Scaffold
Cytocompatibility
In order to confirm the efficacy of the decellularization protocol
prior to recellularization, scaffolds should be evaluated to ensure
that they meet the generally agreed upon criteria set forth by Crapo
et al. [40] with our suggested addition of confirmation of detergent
removal for cytocompatibility assessment [41]. Lung histology
protocols are described in Subheading 3.5. We have previously
published a detailed protocol for assessment of residual DNA and
detergents and direct the reader there for further details [18].
3.5
1. Fix acellular or recellularized mouse lungs in 4% PFA for 24 h at
4 C.
Lung Histology
2. Place the specimens inside histological cassettes with filter
paper for further tissue processing (see Note 10).
3. Dehydrate the specimens in 70%, 90%, and 100% ethanol for
15 min each, respectively.
4. Continue the dehydration with 100% ethanol changes for
another 15, 30, and 45 min.
5. Clear the dehydrated specimen with 20-, 20-, and 45-min
consecutive xylene incubations.
6. Infiltrate the specimen with wax by 30-, 30-, and 45-min
incubations.
7. Embed the specimen in paraffin and cool the blocks down.
8. Cut 3-μm-thick sections and enlarge the sections by floating
them in a water bath at ~45 C before replacing them on
adhesive glass microscopic slides.
9. Let the slides dry on a hot plate before continuing with the
staining.
10. Deparaffinize and stain sections with any standard histological
stain (see Fig. 4 and Notes 11 and 12).
11. Mount cover slips onto the microscopic slides and let the
mounting medium dry before microscopic observation of the
staining.
Acellular Lung Scaffolds for Regenerative Potential
285
Fig. 4 Histological characterization of the decellularized murine lungs. (a, b) Hematoxylin and eosin (H&E), (c,
d) Verhoeff-van Gieson (EVG), (e, f) Alcian blue, (g, h) Masson’s trichrome staining of native (left panel) versus
decellularized (right panel), and murine lungs showing efficacy of the decellularization process (Reprinted with
permission from ref. 25)
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3.6 Murine Lung
Recellularization
Recellularization of the lung can be accomplished by the following
different strategies depending on the experimental questions or
desired endpoint analyses: (1) at the whole lung level through the
airways or vascular system (Fig. 1a, b) or (2) on individual slices of
acellular lung scaffolds. In the instance of recellularization of entire
lobes or entire lungs, the cultivation could either continue as a
whole organ or as lung slices made from the recellularized lungs.
In the second method, recellularization occurs by directly seeding
or incubating cells with lung slices (Fig. 1). Lung slices have previously been generated by hand or through methods similar to those
used for precision-cut lung slices (PCLS). Materials for each strategy are described in separate sections below.
3.6.1 Whole Lung
Recellularization
Cells can be seeded into the acellular lung scaffold IT and/or IV
depending on the cultivation procedure selected. Cell numbers,
amounts, and incubation time needed for attachment will likely
need to be adapted and confirmed for each cell type and combination. Below, the steps to be taken for recellularization at the whole
lung level in an ex vivo perfused/ventilated bioreactor system are
listed (see Note 13).
1. Construct the ex vivo perfusion/ventilation system according
to the setup shown in Fig. 5 (see Note 14).
2. Load the external and the internal lung incubation chamber
(E and I, respectively) with PBS.
3. Place the I chamber in a water bath (w) kept at 37 C.
4. Start the perfusion system by turning on the peristaltic pump
(p). Make sure the stopcock (sc) switch is in proper position
(henceforth called position 1) for the PBS to flow from the E to
the I chamber (a-d line).
5. If air bubbles are observed in the pipeline, remove them by
using the air purge (ap) stopcock, closing the I chamber input
line (d).
6. Once the a-d line is fully filled with PBS, switch the stopcock
(henceforth called position 2) to allow for perfusate to loop
within the I chamber (b-e loop) until it is completely filled
as well.
7. Run PBS through the lung scaffold for 10–15 min in the
b-e loop.
8. Turn off the pump and discard the PBS from the E and I
chambers.
9. Switch the stopcock into position 1 to allow for perfusate to
flow through a-d line again.
10. Load the E chamber with the corresponding cell suspension for
recellularization of the lung vascular tract.
Acellular Lung Scaffolds for Regenerative Potential
287
Fig. 5 Schematic representation of the ex vivo lung perfusion/ventilation system.
System consists of two chambers, i.e., the external chamber (E), the internal
lung incubation chamber (I) connected via tubing, two stopcocks, one (sc) to
control for the external line versus the internal loop, second (ap) to remove the
air bubbles in the tubing before entering the lung incubation chamber, a
peristaltic pump (p) at 0.5 mL/min rate, a water bath (w) at 37 C, as well as
a ventilator (v) for murine lungs with 100 strokes/min and 100 μL stroke volume
11. In parallel, load the I chamber with fresh cell culture medium
(e.g., DMEM/F12 supplemented with 10% FBS and 1% penicillin/streptomycin).
12. Connect the tracheal cannula attached to the heart-lung block
with the ventilation system of the bioreactor and the d line
output to the PA cannula versus the e line input to the aorta
cannula.
13. Instill the corresponding cells selected for recellularization of
the airway tract via the trachea cannula in the heart-lung block.
14. After a certain time of incubation under static conditions for
cell attachment in the airway tract, start running the peristaltic
pump at 0.5 mL/min flow rate to load the lung scaffold with
the vascular tract cell suspension (see Note 15).
15. Once the predetermined amount of perfusate which you want
to instill is loaded into the scaffold (i.e., amount of time to
instill can be determined by dividing the total volume you wish
to instill by the flow rate), turn off the pump, and switch the
stopcock to allow for the perfusate to recirculate through the
internal circuit (b-e loop).
16. Incubate the scaffold again with the cells loaded for cell attachment in the vascular tract.
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17. After incubation, turn on the pump again at 0.5 mL/min flow
rate and start ventilating the scaffold at around 100 strokes/
min with a stroke volume of 100 μL of humidified air (see Note
16).
18. Continue culturing for cellular proliferation and differentiation
for predetermined time-points and exchange the cell culture
media in the I chamber daily.
3.6.2 Airway Route
Recellularization for Slice
Generation
The majority of the reports have exploited the slice technology for
recellularization studies where lung slices are prepared from acellular lungs following IT inoculation of the corresponding cells suspended in low gelling temperature agarose (see Note 17). We have
recently optimized this setup to be compatible with PCLS generation to allow for controlled slice thickness (Fig. 6). The steps to be
followed for such an approach are listed below.
1. Prepare a 3% low gelling temperature agarose solution in cell
culture medium by warming up the mixture (see Note 18).
2. Resuspend 2 mL of the cell suspension in cell culture medium
with 1 mL agarose solution until the cells appear uniformly
dispersed.
3. Administer the cells into the lung scaffold by injecting through
the trachea cannula (see Note 19).
4. Ligate the trachea to prevent the escape of the cell-agarose
solution, and let the recellularized scaffold stay on ice (or at
4 C) for solidification of the agarose (approximately 30 min).
5. Set up the vibratome system and start making lung slices as
described in Subheading 3.3.
6. Place the PCLS samples into cell culture plates submerged in
culture medium.
7. Place the plate in an incubator at 37 C with 5% CO2, and
exchange the culture medium in the wells four times in the first
2–3 h of culturing (see Note 20).
3.6.3 Recellularization
Directly on the PCLS
1. Prepare acellular lung slices as described in Subheading 3.3 and
allow them to float in 24-well plates loaded with fresh media.
2. Place the plates into an incubator at 37 C with 5% CO2 and
exchange the culture medium as stated in Subheading 3.6.2.
3. Before seeding the cells, gently move the PCLS into a new well
and slowly pipette a low-volume cell suspension onto the PCLS
(see Note 21).
4. Let the seeded cells attach within the acellular PCLS by incubating the plate for circa 1 h.
5. Load the wells with fresh medium and continue with the
treatment until the predetermined endpoints.
Acellular Lung Scaffolds for Regenerative Potential
289
Fig. 6 Histological characterization of the murine PCLS recellularization. H&E
staining of the PCLS recellularized via the airway route following precision cut
lung slice generation and culture for 24 h at (a) low and (b) high magnifications
(nuclei shown in violet, proteins shown in pink)
3.7 Tissue
Homogenization
Homogenization of acellular or native tissue must be done under
freezing conditions in order to produce a tissue powder that can be
split into fractions or be fully used for RNA or protein isolation
depending on the availability of tissue.
1. Use approximately one lobe if using a full murine lung, at least
two to three PCLS. Blot tissue to remove extra moisture. Snap
freeze tissue in liquid nitrogen. If homogenization is not performed immediately, store at 80 C in a safe-lock tube with a
small hole in the tube cap for pressure release.
2. Homogenize using one of the devices in Table 2.
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Deniz A. Bölükbas et al.
Table 2
Homogenization of snap-frozen lung tissue in Mikro-Dismembrator S versus TissueLyser II
Mikro-Dismembrator S
TissueLyser II
(a) Adaptor blocks are stored at 80 C
(a) Place 9 mm steel balls in dismembrator tubes
and cool them down in liquid nitrogen or on dry (b) Pre-cool 2 mL safe-lock round-bottom tubes
ice
and beads in liquid nitrogen before starting.
(b) Place lung tissue slices in the cooled tubes
Place adaptor blocks on dry ice during
(c) Cool down sample holder by placing in liquid
preparation
nitrogen
(c) Place sample in cooled 2 mL safe-lock tubes
(d) Place samples in sample holder and quickly
(d) Place 7 mm steel bead on top of sample. Lock
mount the device
lid and place on cold adaptor block
(e) Homogenize the samples for 00:30 s at the
(e) Place adaptor block in TissueLyser II and run at
maximum shaking frequency of 3000/min
frequency of 25/s for 1 min. Quickly return the
(f) Repeat (c–e) three times for each set of samples.
samples to dry ice
Place samples in liquid nitrogen after each cycle (f) Check samples for complete homogenization,
to avoid warming and softening of the tissue
and if samples are not in a powder form,
repeat (d)
3. The produced powder can be divided into several fractions as
needed. Keep sample in liquid nitrogen or on dry ice until
further processing.
3.8 Protein Isolation
from Deand Recellularized
Murine Lung Tissue
1. Homogenize samples as described in Subheading 3.7, and add
200 μL RIPA lysis buffer supplemented with protease and
phosphatase inhibitor cocktails.
2. Vortex sample until it thaws and place on ice for 30 min.
3. Centrifuge samples at 15,000 rpm (16,000 g) for 20 min.
4. Collect the soluble fraction. Save the pellet at
further analysis.
3.9 RNA Isolation
from Deand Recellularized
Murine Lung Tissue
80 C for
1. Homogenize samples as described in Subheading 3.7 and lyse
the frozen samples with 600 μL RNA lysis buffer T containing
6 μL TCEP; mix thoroughly until the sample is liquid and place
it on ice for 1 h. Use smaller volumes of lysis buffer for smaller
samples (e.g., 400 μL for 2 homogenized PCLS).
2. Transfer the lysate into a 1.5 mL tube and add 15 μL proteinase
K to the samples. Vortex thoroughly and incubate at 55 C for
10 min on a thermoblock with a shaking frequency of
700/min.
3. Transfer the lysate into a DNA removing column. Centrifuge at
12,000 g for 1 min.
4. Transfer the flow-through into a new 1.5 mL tube and add one
volume equivalent (600 μL) 70% ethanol.
5. Perform RNA binding and purification according to manufacturer protocol using PerfectBind RNA columns.
Acellular Lung Scaffolds for Regenerative Potential
291
6. Elute with 40 μL sterile RNase-free-dH2O on the membrane.
Centrifuge at 5000 g for 1 min.
7. Re-elute using flow-through and determine concentration
using a NanoDrop or an equivalent device.
4
Notes
1. Various types of mouse models, including those derived from
murine models of disease, have previously been used for lung
de- and/or recellularization. However, each mouse model
must be characterized in depth for the applicability of the
techniques discussed in this chapter.
2. Depending on the instrumentation available, different products can be used for the heart-lung block cannulation. The
most common ones used in the market are available from
Harvard Apparatus [42].
3. Anesthesia should be applied as recommended and approved
by your local guidelines.
4. Several low gelling temperature agarose formulations exist on
the market. The optimal formulation should be chosen according to the physical requirements of the experiment. In particular, variations in the melting temperature of different
formulations can result in alterations in the experiment, as
these would have direct effects on the removal kinetics of the
agarose from the lungs during incubation at 37 C. For
instance, SeaPrep® agarose by Cambrex (used in ref. 25) is
particularly “soft” which results in faster dissociation of the
agarose from the slices under culturing conditions.
5. A number of approaches has been developed to create PCLS
models with the most common ones utilizing Zeiss Hyrax V55
[43] or Krumdieck tissue slicer [44]. Though both methods
are capable of creating PCLS within the same tissue thickness
levels, their cutting strategies are different and may result in
experimental differences.
6. For higher-throughput and paired analysis, punches can be
taken at identical sizes from the PCLS samples.
7. Though any tissue homogenizer can be used, we have found
that a homogenizer with larger beads is sufficient. In this
chapter, we discuss the use of two different homogenizers:
(1) Mikro-Dismembrator S (Sartorius Stedim Biotech
8531609) and (2) TissueLyser II (Qiagen 85300).
8. The cannula may be connected to the ventilation unit once the
mouse is intubated to start external ventilation of the lungs at
rates around 100 strokes of 100 μL air per min.
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9. For histological evaluation of the acellular lungs, the lungs
should be fixed in PFA immediately after the completion of
the decellularization protocol. Otherwise, the lung structure
appears collapsed in histological assessments [25]).
10. Depending on the protocol used for tissue processing, care
should be taken during these steps to ensure that the PCLS
samples do not fold on themselves. In particular, if samples will
be processed in a rotating tissue processor, we have observed
that placing the PCLS in between two biopsy foam pieces in
the cassette helps limit distortion. Alternatively, if your processor has a static chamber, we have found that histological filter
papers are compatible with this processing.
11. The presence of visible red cellular material or “ghost cells” in
H&E or in Masson’s trichrome staining indicates incomplete
cell removal.
12. Retention or loss of elastin and proteoglycans in the lung after
processing can be observed by Verhoeff-van Gieson and Alcian
blue staining.
13. In another model used by Crabbe et al. [24] where the lungs
were cultivated in a rotating wall vessel (RWV) bioreactor, the
lungs were first cultured in static conditions after the seeding of
the cells for 3 days to allow for cell attachment prior to the
dynamic culture conditions in the RWV bioreactors.
14. The bioreactor system discussed in this chapter (Fig. 5) is a
modification of the system previously designed and validated
by Doryab et al. [45].
15. Incubation period of the murine lung scaffold under static
conditions for cell attachment is dependent on the cells used
for the experiment [46].
16. Humidified air is tolerable for short periods of recellularization
studies. However, having humidified air with 5% CO2 is
recommended for long-term cultivation studies.
17. Although the reports to date have utilized recellularization of
lung slices strategy via the airway route, we envision the same
approach can be applied for vascular tract recellularization too.
18. Cool down the agarose solution in a water bath at around
37 C, before resuspending the cells with it.
19. Acellular scaffolds can be ligated at the right bronchus to direct
the cells only to the left lung lobe. In this manner, one can have
paired controls for each individual lung.
20. It is recommended to exchange the media in the wells every
30 min four times at the start of culturing to get rid of the
dissociated agarose and the immediate cytokines released due
to cutting of the lung tissue accumulating in the wells.
Acellular Lung Scaffolds for Regenerative Potential
293
21. In a recent study [26], the PCLS were placed on floating
membranes in culture medium to achieve air-liquid interface
(ALI) conditions.
Acknowledgment
The authors wish to thank all previous lab members who contributed to establishing the methods described here, in particular to
Nicholas Bonenfant, Zack Borg, Dino Sokocevic, Noor Christiaens, Carmela Morrone, and Amelia Payne. Portions of this
work were funded by an ATS Unrestricted Grant (awarded to D.
E.W.) and a Wallenberg Molecular Medicine Fellowship awarded to
D.E.W.
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Chapter 21
Mouse Lung Tissue Slice Culture
Xinhui Wu, Eline M. van Dijk, I. Sophie T. Bos, Loes E. M. Kistemaker,
and Reinoud Gosens
Abstract
Precision-cut lung slices (PCLS) represent an ex vivo model widely used in visualizing interactions between
lung structure and function. The major advantage of this technique is that the presence, differentiation
state, and localization of the more than 40 cell types that make up the lung are in accordance with the
physiological situation found in lung tissue, including the right localization and patterning of extracellular
matrix elements. Here we describe the methodology involved in preparing and culturing PCLS followed by
detailed practical information about their possible applications.
Key words Precision-cut lung slices (PCLS), Ex vivo, Lung tissue functions
1
Introduction
The lung slice represents a lung tissue preparation suitable for a
great variety of studies ranging from airway pharmacology to toxicology [1]. Lung slices are prepared from whole lung by filling the
lung lobes with a low-melting point agarose to be made suitable for
slicing. After slicing, the slices can either be used directly or are
cultured before experimental end-points are determined. Whereas
this paper will focus on the preparation and applications of mouse
lung slices (Fig. 1), lung slices can be prepared from all mammalian
species. We will first briefly review the main strengths, weaknesses,
and applications of the lung slices, followed by a detailed practical
information into the methodologies used.
1.1 Main
Methodological
Strengths
and Weaknesses
of Lung Slices
Probably the main strength of working with lung slices is that the
presence, differentiation state, and localization of the more than
40 cell types that make up the lung are in accordance with the
physiological situation found in the lung, including the right localization and patterning of extracellular matrix elements. Despite
considerable efforts and progress in the area of 3D cell culture
and 3D printing, recapitulation of the complexity of lung tissue in
Ivan Bertoncello (ed.), Mouse Cell Culture: Methods and Protocols, Methods in Molecular Biology, vol. 1940,
https://doi.org/10.1007/978-1-4939-9086-3_21, © Springer Science+Business Media, LLC, part of Springer Nature 2019
297
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Xinhui Wu et al.
Fig. 1 Procedure of precision-cut lung slices (PCLS). This flowchart briefly shows the whole procedure of
precision-cut lung slices
such models is currently not possible. This makes the slice
extremely suitable for studies into the relationships among the
lung, airway or pulmonary vascular structure, and function [2].
Moreover, it has proven extremely difficult to maintain the mature
contractile phenotype of smooth muscle cells in culture [3], and
our understanding of smooth muscle physiology (either airway or
pulmonary vascular) is already restricted in cell culture studies for
that reason alone. Not surprisingly, the first studies conducted with
lung slices were smooth muscle physiology studies [4]. In laboratories working in that area, the slices have taken center stage, mostly
due to the pioneering work of the groups of Christian Martin [4]
and Michael Sanderson [2]. Finally, a significant advantage of the
lung slice model is that multiple slices (up to around 50 in mouse)
can be taken from a single animal for both controls and treatments
study, thereby reducing the use of animals.
However, the lung slice model has weaknesses that should be
taken into account when designing studies and utilizing PCLS.
One of the most significant weaknesses is that it has proven
extremely difficult to culture slices for prolonged periods of
time. This may be due to the diversity of cell types in one lung
slice, each requiring specific culturing media, making it difficult to
find one medium composition that fits all. Research groups working with cultured slices report their use for several days, with
increases in ATP and LDH release, and the presence of dead
cells appearing around day 3–7 [5, 6]. Application of microfluidics
or specific cell culture ingredients may help to prolong this longevity, although virtually no in-depth studies in this area have been
reported.
PCLS Applications in Mice
299
Another significant disadvantage of the lung slice model is that
the tissue is taken out of its context, removing blood supply and
neural network connections to the central nervous system. Accordingly, studies into inflammatory cell recruitment to the lung or
studies into neural reflex control of lung physiology are not possible
in the slice. Having said that, lung slices do contain structural cells
and tissue-embedded inflammatory cells such as macrophages and
mast cells whose function can be studied [6, 7].
1.2
Applications
As discussed above, one of the first applications for the lung slices
was the study of small airway physiology in its natural context of
parenchymal connections. Until lung slices became available, studies into airway physiology were mostly done in tracheal or large
bronchial preparations in traditional organ bath settings. The lung
slices made it possible to study small airway and pulmonary vascular
physiology in response to agonists or electrical field stimulation
(to release endogenous neurotransmitters) [8–10], which is of
great importance as these anatomical regions play a major role in
asthma, COPD, and PAH. Thus, understanding of the receptor
populations and excitation contraction coupling mechanisms in the
distal airways and vessels has provided new insights with important
therapeutic implications. These include the findings that the distal
airways have significant neuronal innervation and functional control and the finding that distal airways have completely different
pharmacology, with large differences in the sensitivity and maximal
response to bronchoconstrictor agents such as methacholine
([8, 9, 11] and own unpublished observations).
The connectivity to the parenchymal structure has additional
benefits as it allows studies into the impact of changes in parenchymal mechanics (e.g., in COPD) on airway mechanics. Such studies
have revealed that the elastin/collagen network in the parenchyma
significantly impacts on airway reactivity and airway reopening,
which is distorted in COPD and in lung slices exposed to elastase
[12–14]. The elastin and collagen fibers can be visualized in the
slices using two-photon confocal microscopy as outlined below.
Strain mapping during bronchoconstriction [15] provides further
insight into these airway-parenchymal interactions and can be used
to map how deep strain penetrates into the lung tissue following
bronchoconstriction.
Moreover, additional applications for the lung slices have been
considered. These include studies into bronchoconstrictioninduced or growth factor-induced airway and lung remodeling.
We have shown that prolonged exposure to bronchoconstrictors
leads to changes in smooth muscle content in the airways [5, 16],
although an important side note here is that this was more readily
measurable in the guinea pig and less so in the mouse. This is
possibly the result of differences in bronchoconstriction to methacholine between these species, as methacholine challenge leads to
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Xinhui Wu et al.
complete airway closure in the guinea pig but only partial closure in
the mouse [13, 16]. Exposure to TGF-β or a mix of growth factors
and mediators involved in lung fibrosis does induce changes in gene
and protein expression of matrix elements and contractile protein in
the mouse and the guinea pig, showing the suitability of the lung
slices for early studies of remodeling and fibrosis as well [17].
The slices may also offer opportunities to study the regenerative capacities of the lung. Alveolar epithelial type II cells (AT II
cells) are difficult to culture in vitro but abundantly expressed in the
lung slices. We have shown that disruption of elastin fibers reduces
the expression of the AT II cell marker pro-SPC in the lung slices
[13], demonstrating the possibility to study the relationships
among ECM structure and airway functions in lung slices. Progenitor cell populations and their response to treatment can also be
studied in the lung slices, as has been done by Uhl et al., who
mapped WNT-active cells in TCF/LEF-H2B:GFP mice, showing
their response to GSK-3 inhibition [6]. Another intriguing application of the lung slices in this area is to decellularize the slices for
subsequent repopulation with progenitor cells to study the impact
of local matrix-derived cues on cell fate decision and differentiation
[18]. This is an active area of research that will likely be expanded
more in the future.
Studies into the immunomodulatory properties of structural
cells can also be done in the lung slices [19], although this method
does not offer the possibility to pinpoint the cellular source of
secreted factors such as cytokines in the medium. The lack of
inflammatory cell recruitment into the lung as would be seen
in vivo also limits the use of the slice for such studies.
Studies of drug metabolism and toxicology can also be done in
the lung slices given the presence of metabolic enzymes such as
P450 in the lung tissue [20–22].
2
Materials
2.1
PCLS Preparation
Prepare all surgery tools (including scissors, tweezers, suture line),
cannula, syringes, cotton pad, ethanol 70%, ice, a 10-cm-diameter
dish, and anesthesia (ketamine and Dexdomitor) and agarose
medium before going to the animal facility. Slicing medium, incubation, and washing medium should be prepared in advance.
A tissue slicer machine (Leica VT 1000 S vibrating blade microtome, Leica Biosystems B.V., Amsterdam, the Netherlands) is used
in this protocol.
1. Anesthetics: Ketamine (40 mg/kg) and
(0.5 mg/kg) are used in the experiments.
Dexdomitor
2. Agarose medium: Agarose powder was dissolved in a solution
composed of CaCl2 (0.9 mM), MgSO4 (0.4 mM), KCl
PCLS Applications in Mice
301
(2.7 mM), NaCl (58.2 mM), NaH2PO4 (0.6 mM), glucose
(8.4 mM), NaHCO3 (13 mM), HEPES (12.6 mM), sodium
pyruvate (0.5 mM), glutamine (1 mM), MEM-amino acid
mixture (1:50), and MEM-vitamins mixture (1:100,
pH ¼ 7.2) within ultrapure water (UP). A final concentration
of 1.5% agarose is n used to fill in the mouse lung (see Note 1).
3. Slicing medium: Medium composed of CaCl2 (1.8 mM),
MgSO4 (0.8 mM), KCl (5.4 mM), NaCl (116.4 mM),
NaH2PO4 (1.2 mM), glucose (16.7 mM), NaHCO3
(26.1 mM), HEPES (25.2 mM), and pH ¼ 7.2 within ultrapure water (UP).
4. Incubation medium: Medium composed of CaCl2 (1.8 mM),
MgSO4 (0.8 mM), KCl (5.4 mM), NaCl (116.4 mM),
NaH2PO4 (1.2 mM), glucose (16.7 mM), NaHCO3
(26.1 mM), HEPES (25.2 mM), pH ¼ 7.2 within ultrapure
water (UP). Add 1% Na-pyruvate (100 mM), 2% nonessential
amino acids (100), 1% MEM-vitamin solution (100), 1%
L-glutamine (100), and 1% Penicillin-Streptomycin (Pen-Strep, 5000 units/mL penicillin and 5000 μg/mL streptomycin, Gibco® by Life Technologies) to the medium before use.
5. Culture medium: DMEM supplemented with sodium pyruvate
(1 mM), MEM nonessential amino acid mixture, gentamycin,
Penicillin-Streptomycin (Pen-Strep, 5000 units/mL penicillin
and 5000 μg/mL streptomycin, Gibco® by Life Technologies), and amphotericin B (1.5 μg/mL; Gibco® by Life
Technologies).
2.2 PCLS
Decellularization
1. Washing solution: Solution composed of sterile ultrapure water
(UP) with 5% Penicillin-Streptomycin (Pen-Strep, 5000 units/
mL penicillin and 5000 μg/mL streptomycin, Gibco® by Life
Technologies).
2. Triton solution: 0.1% Triton X-100 in 5% PenicillinStreptomycin in UP.
3. SDC solution: 2% sodium deoxycholate in 5% PenicillinStreptomycin in UP.
4. NaCl solution: 1 M NaCl solution with 5% PenicillinStreptomycin in UP.
5. Dnase I solution: Solution composed of 30 μg/mL Dnase I,
2 mM CaCl2, 1.3 mM MgSO4.
6. Peracetic acid solution: 0.1% peracetic acid in 40% ethanol.
7. Storage solution: PBS supplemented with 5% PenicillinStreptomycin, 0.1 mg/mL gentamycin (Gibco® by Life Technologies), 25 mg/mL Fungizone.
8. Cytoskeleton Buffer: CB buffer, MES (10 mM), NaCl
(150 mM), EGTA (5 mM), and glucose (5 mM), pH ¼ 6.1.
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9. Cyto-TBS: Tris-base (20 mM), NaCl (154 mM), EGTA
(2.0 mM), and MgCl2 (2.0 mM), pH ¼ 7.2.
10. Cyto-TBST: cyto-TBS containing 0.1% Tween-20.
3
Methods
3.1 Isolation
of Murine Lung
1. Weigh the animal first by using a scale.
2. Euthanize the mouse following subcutaneous injection of ketamine and Dexdomitor (see Subheading 2). Observe the mouse
and check the depth of anesthesia by pressing the feet and eye
reflexes (blink eyes can be checked by approaching a cotton
stick to the eyes). Once the mouse did not show feet and eye
reflexes anymore, pin the animal to a base. Open the abdominal
cavity with scissors by cutting the skin and peritoneum from the
middle of the abdomen up to the jaw. Pull the intestines aside
with forceps and cut the inferior vena cava and aorta abdominalis to exsanguinate the animal. Puncture the thoracic diaphragm with the sharp tip of the scissors to allow expansion
of the rib cage, being careful not to cut the lung [23, 24] to
open the thoracic cavity.
3. Clear the muscle tissue away from the trachea by grabbing the
tissue and manually pulling it away from the underlying trachea
with forceps. Make a small incision in the trachea on the
anterior side of the thickest band of cartilage using fine forceps
or scissors, being careful not to cut off the trachea. Insert the
cannula into the trachea through the incision and use the
suture line to tie the cannula firmly in place.
4. Inflate the lung by injecting approximately 1.5 mL of
low-melting point agarose solution through the cannula
making sure that the distal tips of the lung are also filled with
agarose medium (see Note 2).
5. After agarose injection, tie off the cannulated trachea with the
suture line to prevent the agarose medium from flowing out of
the lung (see Note 3).
6. Keep the cannula inserted in the trachea. Cover the lung with
ice after agarose injection, and place it in the fridge (4 C), for
20 min to let the agarose solidify within the lung (see Note 4).
7. Once the agarose has solidified, remove the cannula from the
trachea, carefully excising the agarose-inflated lung. Cut off the
trachea and remove the front ribs around the heart, and then
remove the connective tissue in the back and take out the lung.
Put the lung in a dish and keep it on ice.
PCLS Applications in Mice
3.2 Preparation
of PCLS
303
1. Separate the lung into individual lobes, and remove the connective tissue between each lobe. Then use each lobe as a
resource to obtain lung slices.
2. A tissue slicer, Leica VT 1000 S vibrating blade microtome
(Leica Biosystems B.V., Amsterdam, the Netherlands) is used
to cut lung slices in this protocol (Fig. 1). Follow the instruction of this slicer machine to cut lung slices (see Note 5).
3. Before cutting, a higher-dose of agarose medium (2–5%) is
recommended to make a gel column around the lung lobe to
facilitate slicing.
4. 250 μm-thick lung slices are cut in slicing medium at 4 C and
collected in incubation medium at 37 C.
5. The lung slices are incubated in a humidified incubator in
atmosphere of 5% CO2/95% air at 37 C. Lung slices are
washed in every 30 min, four times in total, using the incubation medium.
6. Lung slices are placed in incubation medium and cultured at
37 C in 12-well culture plates, using three to four slices
per well.
Lung slices prepared using this procedure can be used in a
number of experimental applications. Previous work from our lab
(unpublished) demonstrates that murine lung slices viability is
preserved for 72 h of culturing, as mitochondrial activity did not
change during this time window. This indicates that the lung slices
are viable for at least 3 days.
3.3 Airway
Narrowing Studies
Airway narrowing can be studied by fixating the lung slices into a
3-well cluster (Fig. 2). These slices are then exposed to a contractile
stimulus following which airway contraction is recorded using a
microscope.
1. 12-well cell culture plates can be cut into small 3-well clusters
which will fit under a microscope. These 3-well clusters can be
used for the airway contraction studies.
Fig. 2 Representative images of airway narrowing studies. (a) Device used in the study of airway narrowing.
(b) Two adjacent airways before treatment, 40. (c) Airway narrowing in response to methacholine (MCh),
40. (d) Airway reopening in response to chloroquine (Cq), 40
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Xinhui Wu et al.
2. Select slices with airways of the desired size. To do so, fill the
wells of the custom-made 3-well clusters with 1 mL of warm
(37 C) incubation medium per well. Put one lung slice to each
well, and inspect these slices using a microscope (we used
Eclipse, TS100; Nikon). It is recommended to inspect a few
slices as not every slice will contain the desired airway size. Only
use slices in which the airways are cut in cross-sectional manner.
Determine the airway size by using the image acquisition software (NIS-elements; Nikon, (see Note 6)).
3. Following medium removal, use a nylon mesh and metal
washer to fix the lung slice, as described previously [25]. The
nylon mesh (which is also washer shaped) should be slightly
bigger than the metal washer to make sure the PCLS tissue
does not come in direct contact with the metal washer. First
place the nylon mesh on top of the slice, then place the metal
washer. The slice is now fixated, while the airway of interest is
still visible through the hole in the middle of the nylon mesh
and metal washer. For ease of preparation, one could carefully
remove the medium from the well using a pipette, leaving the
slice in the well. Following fixation, 1 mL of warm (37 C)
incubation medium has to be added again. Put the plate back
under the microscope, on top of the heating pad, and fix the
plate. Choose the desired magnification and focus the microscope (see Fig. 2).
4. Capture the airway contraction in time-lapse (one frame per
2 s) via the microscope using image acquisition software
(NIS-elements; Nikon). Start the time-lapse and wait for
2 min to record the baseline airway luminal area. Following
2 min, add methacholine in increasing concentration
(109 M–103 M final concentrations) to the well using a
pipette. Be careful not to touch the slice or chamber with the
tip of the pipette. Use an interval of 7.5 min between each dose
(see Note 7). Following the dose response curve for methacholine, it is also possible to dilate the airways again, e.g., using
the bitter taste receptor agonist chloroquine (103 M) (see
Note 8). One could also use other approaches including
β-agonists, but might be less successful.
5. Quantification airway luminal area: Image acquisition software
(NIS-elements; Nikon) could be used to quantify the airway
luminal area.
3.4 Collagen
and Elastin Imaging by
Two-Photon
and Multiphoton
Microscopy
1. Wash the lung slices twice with PBS (500 μL/slice), and leave
the PBS in the well after the second wash. Using a small spatula
with a flat end, scoop one slice out of the well and carefully
place it onto a microscope slide. Make sure the slice does not
fold. Using a piece of paper towel, carefully remove any excessive PBS surrounding the slice. Place a coverslip on top of the
PCLS Applications in Mice
305
Fig. 3 Representative images from two-photon imaging and multiphoton imaging. (a) two-photon and
multiphoton excitation fluorescence imaging are used to visualize α-sm-actin(green) and collagen(red); (b)
two-photon and multiphoton excitation fluorescence imaging are used to visualize collagen (green) and elastin
(red) polymers in lung slices [13]
slice. Seal the edges of the coverslip using transparent nail
polish (see Note 9). As the protocol mentioned above, the
imaging has to be performed straight away if use fresh tissue.
2. Two-photon and multiphoton excitation fluorescence (MPEF)
imaging can be used to visualize collagen and elastin polymers,
respectively, as described previously [26]. Under excitation at
820 nm, the collagen bundles will naturally emit a second
harmonic generation signal which can be collected around
410 nm. Elastin can be visualized by using its endogenous
fluorescence. Elastin images can be generated by using an
infrared laser (excitation wavelength 880 nm). The broadband
emission spectrum ranges from 455 to 650 nm with a peak at
~500 nm (Fig. 3).
3.5 Mean Linear
Intercept (Lmi)
Measurement
The mean linear intercept (Lmi) can be determined as a measurement of alveolar airspace size. This can be measured either by
confocal microscopy or by light microscopy.
1. Lung slices used for contraction experiments or non-used slices
are washed four times with incubation medium. Then they are
transferred to an embedding cassette filled with a biopsy pad
(see Note 10). The slice is placed on the pad, and a second pad
is placed on top of the slice before closing the cassette and
placing it in a formalin solution (10%) for 24 h. The slices are
processed for paraffin embedding and embedded into paraffin
blocks. Sections of 4 μm are cut with a microtome and stained
by H&E staining. Twenty random photomicrographs of the
parenchyma of lung (magnification 200) tissue rather than
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airways or blood vessels are made, as airways or blood will
interfere with the measurement of alveolar airspaces.
2. This method is previously described by van der Strate [27]. In
short, a sheet with vertical lines in three horizontal rows (21 in
total) is placed on the top of the photograph. Whenever an
intercept crosses the parenchymal walls, two points will be
given. When the intercept touches the parenchymal cells, one
point will be given. Importantly, intercepts that cross or touch
blood vessels or airways are not taken into account to prevent
misjudgments. When more than three intercepts are crossing
or touching blood vessels or airways, the picture should not be
taken into account and a different field should be chosen. With
the total scores, the Lmi is calculated as: (n l 2)/m in
which “n” represents the number of intercepts, “l” represents
the length of the individual lines (as calculated with the scale of
microscopic photo), and “m” refers to the amount of points
given.
3. Proteins could be visualized by immunofluorescence
(described below). Fluorescence can be determined with a
confocal laser scanning microscope (CLSM) equipped with
true confocal scanner (TCS; SP8 Leica, Heidelberg, Germany),
using a 200 lens. To avoid bleed-through, sequential scans
need to be performed. Alexa Fluor 488 can be excited using the
488 nm blue laser line, and Cy™3 can be excited using the
552 nm green laser line. Record all images in the linear range,
at an image resolution of 1024 1024 pixels and with a
pinhole size of 1 Airy unit, while avoiding local saturation [28].
4. Single Z-stack images can be used to quantify the Lmi with the
analysis method described above (see Note 11).
3.6
Decellularization
1. Decellularization with detergents (Acellular scaffolds) maintains the architecture and proteins of extracellular matrix for
use as scaffolds in the field of lung tissue engineering or progenitor cell biology. We decellularized the lung slices for
subsequent repopulation with progenitor cells to study the
impact of local matrix-derived cues on cell fate decision and
differentiation (Fig. 4). Place 1–4 slices in each well of a 24-well
plate, and incubate the slices overnight in 1% Triton X-100
medium with 5% Penicillin-Streptomycin (1 mL medium per
well), at 4 C .
2. Wash the slices twice in washing solution for decellularization
(see Subheading 2).
3. Incubate the slices in a 2% sodium deoxycholate (SDC) solution for 3 h at room temperature.
4. Wash the slices twice in washing solution for decellularization.
PCLS Applications in Mice
307
Fig. 4 Representative images of naive and decellularized lung slices. (a) Naive lung slice, 40. (b)
Decellularized lung slice, 40. (c) Masson’s trichrome stain (see Note 14) on naive lung slice, 40; (d)
Masson’s trichrome stain on decellularized lung slice, 40
5. Incubate the slices in 1 M NaCl solution for 1 h at room
temperature.
6. Wash the slices twice in washing solution for decellularization.
7. Incubate the slices in Dnase I solution for 1 h at room
temperature.
8. Wash the slices twice in washing solution for decellularization.
9. Wash the slices in 0.1% peracetic acid for 1 h at room
temperature.
10. Store the slices in storage solution. Slices can be stored in the
storage solution for short periods at 4 C; place the slices at
20 C in storage solution for long term.
11. Slices are now ready to be repopulated with progenitor cells.
3.7 mRNA Isolation
and Real-Time PCR
Total RNA is extracted from PCLS by using the Maxwell 16 instrument and corresponding Maxwell 16 LEV simply RNA tissue kit
(Promega, Madison, USA) for automated purification according to
manufacturer’s instructions. This is an optional method to extract
RNA from PCLS, as the quality of RNA obtained with other
methods including TRIzol and kit extraction is too low to be
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Xinhui Wu et al.
used for experiments. The reverse transcription system (Promega,
Madison, USA) is used to reverse transcribe total RNA (1 μg) into
cDNA. 1 μL diluted cDNA (1:20) is subjected to the Illumina Eco
Personal QPCR System (Westburg, Leusden, the Netherlands)
using FastStart Universal SYBR Green Master (Rox) from Roche
Applied Science (Mannheim, Germany). The cycle parameters used
in real-time PCR system are denaturation at 95 C for 30 s, annealing at 59 C for 30 s, and extension at 72 C for 30 s for 40 cycles
followed by 5 min at 72 C. The amount of target genes could be
normalized to the housekeeping genes such as β-2 microglobulin
(B2M) and ribosomal protein L13A (RPL13). LinRegPCR analysis
software was used to analyze data.
3.8 Immunofluorescence Imaging
Immunofluorescence was performed as described below (Fig. 5).
Primary antibodies and secondary antibodies could be obtained
from various companies according to the research interests. Rabbit
anti-Prosurfactant Protein C (proSP-C, EMD Millipore Corporation, CA,USA) and mouse anti E-cadherin (BD Biosciences, Bedford, MA, USA) were used in our study by this method.
1. Fixation
(a) Wash lung slices twice with cold (4 C) cytoskeleton
buffer (CB buffer).
(b) Incubate the slices for 15 min with 3% paraformaldehyde
(PFA) at room temperature (400 μL/slice).
(c) Incubate the slices for 5 min with 3% PFA + 0.3% Triton
X-100 at room temperature (400 μL/slice).
(d) Wash the slices twice with cold (4 C) CB buffer (see
Note 12).
Fig. 5 Representative immunofluorescence images of mouse PCLS. (a) Blue signals are DAPI, which stained
the nuclei, 63; (b) green signals represent the expression of surfactant protein c (SPC), which is an alveolar
epithelial type 2 cell marker, 63 (c) a merge picture of a and b 63
PCLS Applications in Mice
309
2. Blocking
(a) Prepare blocking buffer (1 cyto-TBS with 1% BSA and
2% normal donkey serum).
(b) Incubate the lung slices with blocking buffer (250 μL/
slice) for 1 h on the shaker at room temperature.
3. Incubation
(a) Dilute primary antibody in cyto-TBST solution.
(b) Incubate the lung slices with primary antibody (use
200 μL/slice) for 1.5 h at room temperature or overnight
at 4 C.
(c) Wash the lung slices with cyto-TBST (500 μL/slice), for
15 min, repeat three times (see Note 13).
(d) Dilute the secondary antibody (1:50) in cyto-TBST
(250 μL/slide) and incubate for 2–3 h at room
temperature.
(e) Wash the lung slices with cyto-TBST (500 μL/slide), for
15 min, repeat three times.
(f) Wash the lung slices with UP water twice quickly.
(g) Wash the lung slices with UP water for 2 min, repeat four
times.
4. Anti-fade staining
(a) Transfer the slices to glass slides.
(b) Add 10 μL/slide anti-fade reagent (Invitrogen, Breda, the
Netherlands) on the glass slide (cover the whole slice) and
cover them with clean microscopic glass plate.
(c) Seal coverslips using the transparent nail polish.
5. Use fluorescence microscope to make images (Fig. 5).
4
Notes
1. The agarose medium must be kept warm in thermal bottle
(around 37 C) so that it will not solidify prior to injection.
2. The volume of agarose is dependent on the size of lung and
should not exceed lung capacity. Since lung tissue is highly
compliant and easily damaged, injection pressure should also
be minimized.
3. If this PCLS model is used to study the arteriole physiology, 6%
gelatin should be used to perfuse the pulmonary arteries [2].
4. This step aims to use the agarose to maintain the shape of the
lung, which makes it easier to cut lung slices.
5. The cutting frequency of 90 Hz, the amplitude of 1.0 mm, and
the sectioning speed of 2.25 mm/s are chosen in this protocol.
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6. The microscope should have a see-through heating pad that
can be kept at 37 C on which the plate with slices can be
placed. This is especially important during the contraction
experiments as these can last up to 1 h.
7. In the described system, medium is not washed away, and
methacholine accumulates in the well with each new dose.
8. In mouse lung slices, β-agonists are less effective in inducing
airway relaxation than chloroquine.
9. This will prevent the coverslip and slice from moving during
the microscopy.
10. The pads are needed to keep the slices flat without wrinkles.
Because the slice is only 250 μm thick, when put in a paraffin
block, it should be very straight; otherwise it is impossible to
make sections.
11. Z-stacks imaging is optional, making it possible to access structures throughout the slice. Image J 1.48d can be used to
further process images.
12. Optional: store in 1 cyto-TBS buffer (200 μL) in sealed
chamber up to maximum 2 weeks.
13. Since the second antibody is labeled by fluorophore, the
experiments should be performed in dark room.
14. Masson’s trichrome stain is a three-color staining protocol
used in histology. Trichrome stain (Masson) kit (SigmaAldrich) is used to stain the lung slices in this study by using
three dyes: hematoxylin (for nucleus), aniline blue (connective
tissue), and Biebrich scarlet (cytoplasm). Cytoplasm and muscle fibers stain red, whereas collagen displays blue coloration.
Acknowledgment
This work is supported by a ZonMW-Vidi grant from the Netherlands Organization for Scientific Research (016.126.307), a grant
for the Dutch Lung Foundation (3.2.08.014) and the China Scholarship Council (File No. 201707720065).
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INDEX
A
Adult stem cells ............................................................... 33
Air-liquid interface (ALI) culture................................158,
159, 164–166, 293
Alkaline phosphatase assay................................. 64, 66, 69
Allergens ........................................................... 10–13, 205
Animal ethics .................................... 16, 18, 20, 113, 272
B
Bleomycin ..................................................................15, 16
Bone marrow transplantation.............................. 129, 132
Brightfield microscopy......................................... 112, 117
Bromodeoxyuridine incorporation .............................. 214
Epidermal keratinocytes
isolation and culture ............................................... 207
proliferation and differentiation assay.................... 205
Extracellular matrix ................................15, 24, 193, 206,
240, 276, 297, 306
F
Fluorescence activated cell sorting (FACS) ........... 43, 54,
55, 66, 69, 131, 135, 136, 138, 172, 174–177,
179, 224, 235, 236, 240–242, 249, 269, 270,
272, 273
Fluorescent microscopy ............... 57, 112, 117, 120, 207
Fluorescent reporter constructs ................................... 119
G
C
Cardiomyocyte isolation ...................................... 193–204
Cell culture
gas phase ...............................................25, 27–29, 219
incubation conditions .........................................25, 29
Chronic liver injury .............................................. 267–273
Collagen coating ................. 97, 161, 164, 166, 207, 219
Colony picking ..........................................................68–70
Colorimetric assay ....................................... 207, 210, 214
CreERT2/LoxP-STOP-LoxP reporter
system .................................112, 113, 119–122
CRISPR-Cas9............................................................77, 79
Gene editing ..................................................... 77–95, 182
Gene transfer ..................................................47, 182, 194
Green fluorescent protein (GFP) .....................50, 54, 55,
58, 59, 78, 79, 86, 91, 92, 124, 145, 147, 149,
152, 153, 182, 238, 272, 273, 300
H
HEK293T cells.............................................48–52, 55, 57
Hematopoietic stem cells (HSC) .....................26, 28, 98,
129–141
Hepatocyte progenitor cells ................................ 267–273
D
I
Decellularised lung scaffold................................. 275–292
Density gradient separation .......................................... 134
Imaging and analysis
image analysis ................................................. 109, 113
immunofluorescent imaging.......................... 123, 126
live imaging ..................................110, 112, 117–122,
124, 277
still imaging ................................... 110, 112, 117–122
Immuno-labelling ......................................................... 134
Immunomagnetic cell separation ........................ 134, 135
Induced pluripotent stem cells (iPSCs) ................. 47, 63,
65, 67–75, 145
Intraocular transplantation .................................. 150, 151
E
Electroporation ..................................... 78, 81, 83–86, 91
Embryonic lung ................................................... 115, 117
Embryonic peripheral blood ..................................97–106
Embryonic stem cells
cryopreservation ..................................................86, 87
passaging....................................................... 40, 86, 87
thawing and recovery............................ 39, 40, 90, 91
Endothelial colony forming cells (ECFC).............97–106
Epiblast stem cells
cryopreservation ........................................................ 87
passaging.................................................................... 87
L
Lacrimal gland epithelial cells.............................. 169–180
Lentiviral transduction
embryonic stem cells...........................................63, 64
Ivan Bertoncello (ed.), Mouse Cell Culture: Methods and Protocols, Methods in Molecular Biology, vol. 1940,
https://doi.org/10.1007/978-1-4939-9086-3, © Springer Science+Business Media, LLC, part of Springer Nature 2019
313
MOUSE CELL CULTURE: METHODS AND PROTOCOLS
314 Index
Lentiviral transduction (cont.)
hematopoietic stem cells....................... 129, 132, 140
mesenchymal stem cells ........................144–147, 149,
150, 153, 154
Lentiviral vectors
concentration ......................................................48, 49
production and titration .............................. 49, 50, 52
titration .................................................. 49, 50, 54, 55
Leukemia inhibitory factor (LIF).....................34, 35, 38,
64, 65, 79, 124, 126
Lineage-tracing ........................................... 232, 276, 277
Liver stem/progenitor cells (LPCs)............................. 267
Lung dissection ........................................... 111, 113, 114
Lung epithelium............................................................ 118
Lung explants ............................ 110–112, 114, 117–119,
121, 122, 124, 126
Lung mean linear intercept measurement .......... 305, 306
Lung mesenchyme ..............................110–114, 117, 118
Lung recellularisation .........................277, 279, 286, 287
M
Mammary epithelial progenitor cells .................. 217–229
Matrigel colony assay .................................................... 219
Medium buffering.............................................. 25–27, 29
Medium pH........................................................ 25–27, 29
Mesenchymal stem cells
cryopreservation ..................................................39, 41
lentiviral transduction .......................... 146, 147, 149,
150, 153
thawing and passaging ................................... 146, 148
Microenvironment .................................. 23–29, 129, 276
Middle ear epithelial cells .................................... 157–166
Mouse embryonic fibroblasts
cryopreservation .................................................. 35–38
harvesting .................................................................. 38
irradiation ............................................................36, 38
thawing and recovery............... 36, 37, 39, 42, 67, 90
Mouse models
chronic liver injury ......................................... 267–273
genetically engineered mice (GEM) ............ 232, 233,
235, 239, 240, 243, 245, 246, 249, 251
mouse strain differences ........................................... 10
respiratory disease models
asthma .............................................................. 7–13
chronic obstructive pulmonary disease
(COPD) ........................................ 7, 13–14, 19
pulmonary fibrosis.............................. 7, 10, 14–17
Mouse retina......................................................... 181–189
O
OP9 stromal cells ...........................................98, 100–106
Organ culture ....................................................... 158, 182
Organoid culture....................... 226, 232, 236, 241–244,
250, 252, 253
Organotypic culture ............................................. 181–189
Osmolarity ................................................. 25, 28, 29, 140
Otitis media (OM) ............................................... 158, 159
Oxygen tension .........................................................27–29
P
Pancreatic islets of Langerhans
culture ............................................................. 255–263
isolation .......................................................... 255–263
PCR amplification .............................................. 82, 87–89
Photoreceptors ...........................181, 182, 184, 186, 189
Plasmids ...................................................... 48, 50, 57, 65,
78, 79, 81–83, 88, 90, 91, 124
Pluripotency ...................... 8, 33, 34, 38, 42, 43, 64, 169
Pluripotency factors
c-Myc ......................................................................... 63
Gbx2 .......................................................................... 34
Klf4 ......................................................................34, 63
Oct4 .............................................................. 42, 43, 63
Sox2 .....................................................................42, 63
Tfcp2l1 ...................................................................... 34
Polyethylene glycol (PEG) ............ 49, 50, 52, 54, 57, 58
Precision-cut lung slices (PCLS) ............... 278, 279, 283,
284, 286, 287, 289–293, 298, 300–304,
307–309
Prostate cancer cells ........................... 231–233, 240–243,
245, 246, 252
Prostate luminal epithelial progenitor cells ........ 231, 248
Puromycin selection..............................78, 79, 81, 84, 91
Q
Quantitative PCR................................................. 239, 250
R
Reaggregated 3D cultures ............................................ 169
Real-time PCR ............................................ 239, 250, 307
Reprogramming ....................... 47, 63–65, 67–69, 71, 75
Retinal explant...................................................... 181–189
Retinal transplant ........................................ 143, 144, 151
S
Smoking mouse model ................................................... 14
N
Neuroprotection .................................................. 143, 145
Neuroretinal cells .......................................................... 182
Neurotrophic factors.......................... 144–146, 149–151,
153, 154, 184
T
Teratoma formation assay..................... 43, 64, 66, 69–74
Tissue dissociation
heart ...............................................193, 194, 196, 203
liver ................................................................. 268, 269
lung ...........................................................15, 115, 290
mammary gland.............................218, 219, 221, 223
pancreas .......................................................... 256, 260
prostate gland ................................232, 236, 239, 251
skin epidermis.......................................................... 208
Tissue engineering ........................................................ 306
Tracheobronchial epithelial (TBE) cells ...................... 158
Two-photon and multiphoton microscopy ................. 304
MOUSE CELL CULTURE: METHODS AND PROTOCOLS
Index 315
U
UVB irradiation..........................205, 206, 212, 213, 215
V
Viral transduction...................................... 54, 55, 58, 153

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