Current Protocols in Mouse Biology (Volume 1)

CURRENT PROTOCOLS in Mouse Biology

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Current Protocols in Mouse Biology

Online ISBN: 9780470942390 DOI: 10.1002/9780470942390 Editors & Contributors

EDITORIAL BOARD Johan Auwerx Ecole Polytechnique Fédérale de Lausanne Stephen D. Brown Harwell Science and Innovation Campus, UK Monica Justice Baylor College of Medicine David D. Moore Baylor College of Medicine Susan L. Ackerman The Jackson Laboratory Joseph Nadeau Institute for Systems Biology CONTRIBUTORS Cristina Antal Mouse Clinical Institute Illkirch, France and Institut d’Histologie, Faculté de Médecine Strasbourg, France Johan Auwerx Institut Clinique de la Souris (ICS) Illkirch, France and Ecole Polytechnique Fédérale de Lausanne Lausanne, Switzerland Abdelkader Ayadi Institut Clinique de la Souris (ICS) Illkirch, France Bernard Baertschi Institute for Biomedical Ethics University of Geneva Geneva, Switzerland Gareth T. Banks Neurobehavioural Genetics MRC Harwell

Harwell Science and Innovation Campus Oxfordshire, United Kingdom Isabelle Barde School of Life Sciences and “Frontiers in Genetics” National Program Ecole Polytechnique Fédérale de Lausanne (EPFL) Lausanne, Switzerland Fernando J. Benavides The University of Texas M.D. Anderson Cancer Center Science Park-Research Division Smithville, Texas Marie-Christine Birling Institut Clinique de la Souris (ICS) Illkirch, France Gemma Brufau Department of Pediatrics Center for Liver, Digestive, and Metabolic Diseases University Medical Center Groningen University of Groningen Groningen, The Netherlands Pierre Chambon Institut de Génétique et de Biologie Moléculaire et Cellulaire Université de Strasbourg, and Collége de France Illkirch, France Nathalie Chartoire Institut Clinique de la Souris (ICS) Illkirch, France Luis E. Donate Spanish National Cancer Research Centre (CNIO) Madrid, Spain Pascal Escher IRO-Institute for Research in Ophthalmology Sion, Switzerland and Department of Ophthalmology University of Lausanne Lausanne, Switzerland Jérôme N. Feige MusculoSkeletal Diseases Novartis Institute for Biomedical Research Basel, Switzerland Giséle Ferrand Ecole Polytechnique Fédérale de Lausanne Lausanne, Switzerland Shumin Gao The University of Medicine & Dentistry of New Jersey New Jersey Medical School Newark, New Jersey Françoise Gofflot Institut Clinique de la Souris (ICS) Illkirch, France and Université Catholique de Louvain Life Science Institute Louvain-la-Neuve, Belgium Isabelle Goncalves da Cruz Institut Clinique de la Souris (ICS) Illkirch, France

Jacob B. Griffin Roche Madison Inc. Madison, Wisconsin Albert K. Groen Department of Pediatrics Center for Liver, Digestive, and Metabolic Diseases University Medical Center Groningen University of Groningen Groningen, The Netherlands and Department of Laboratory Medicine Center for Liver, Digestive, and Metabolic Diseases University Medical Center Groningen University of Groningen Groningen, The Netherlands Jean-Louis Guénet Département de Biologie du Développement Institut Pasteur Paris, France Marcel Gyger EPFL—Center of Phenogenomics Lausanne, Switzerland Yann Hérault Mouse Clinical Institute Illkirch, France and Institut de Génétique et de Biologie Moléculaire et Cellulaire (IGBMC) CNRS/INSERM/Université Louis Pasteur Illkirch, France David Ho The University of Medicine & Dentistry of New Jersey New Jersey Medical School Newark, New Jersey Chull Hong The University of Medicine & Dentistry of New Jersey New Jersey Medical School Newark, New Jersey Neil J. Ingham Wellcome Trust Sanger Institute Wellcome Trust Genome Campus Hinxton, Cambridge, United Kingdom Ralf Kühn German Research Center for Environmental Health Munich, Germany and Technical University Munich Munich, Germany David A. Largaespada Department of Genetics, Cell Biology, and Development University of Minnesota Minneapolis, Minnesota, Center for Genome Engineering University of Minnesota Minneapolis, Minnesota, and Masonic Cancer Center University of Minnesota Minneapolis, Minnesota Pontus Lundberg Department of Biomedicine Experimental Hematology University Hospital Basel

Basel, Switzerland Stefan Marcaletti MusculoSkeletal Diseases Novartis Institute for Biomedical Research Basel, Switzerland Manuel Mark Mouse Clinical Institute Illkirch, France, Institut de Génétique et de Biologie Moléculaire et Cellulaire (IGBMC) CNRS/INSERM/Université Louis Pasteur Illkirch, France and Hôpital Universitaire de Strasbourg Strasbourg, France Daniel Metzger Institut de Génétique et de Biologie Moléculaire et Cellulaire Université de Strasbourg, and Collége de France Illkirch, France Michael S. Minett Molecular Nociception Group Wolfson Institute for Biomedical Research University College London London, United Kingdom and London Pain Consortium Kings College London London, United Kingdom Salvatore Modica Institute of Food, Nutrition, and Health ETH Zurich Schwerzenbach, Switzerland Branden Moriarty Department of Genetics, Cell Biology, and Development University of Minnesota Minneapolis, Minnesota, Center for Genome Engineering University of Minnesota Minneapolis, Minnesota, and Masonic Cancer Center University of Minnesota Minneapolis, Minnesota Antonio Moschetta Laboratory of Lipid Metabolism and Cancer Department of Translational Pharmacology Consorzio Mario Negri Sud Santa Maria Imbaro, Italy and Clinica Medica “A. Murri” Department of Internal and Public Medicine University Aldo Moro of Bari Bari, Italy Francisca Mulero Spanish National Cancer Research Centre (CNIO) Madrid, Spain Stéphanie Muller Mouse Clinical Institute Illkirch, France Stefania Murzilli Laboratory of Lipid Metabolism and Cancer Department of Translational Pharmacology Consorzio Mario Negri Sud

Santa Maria Imbaro, Italy Patrick M. Nolan Neurobehavioural Genetics MRC Harwell Harwell Science and Innovation Campus Oxfordshire, United Kingdom Sandra Offner School of Life Sciences and “Frontiers in Genetics” National Program Ecole Polytechnique Fédérale de Lausanne (EPFL) Lausanne, Switzerland Selina Pearson Wellcome Trust Sanger Institute Wellcome Trust Genome Campus Hinxton, Cambridge, United Kingdom Kathryn Quick Molecular Nociception Group Wolfson Institute for Biomedical Research University College London London, United Kingdom and London Pain Consortium Kings College London London, United Kingdom Hannah G. Radley-Crabb School of Anatomy and Human Biology The University of Western Australia Crawley, Australia Richard R. Ribchester Euan MacDonald Centre for Motor Neurone Disease Research University of Edinburgh Edinburgh, Scotland, United Kingdom Daniel F. Schorderet IRO-Institute for Research in Ophthalmology Sion, Switzerland, Department of Ophthalmology University of Lausanne Lausanne, Switzerland, and EPFL-Ecole Polytechnique Fédérale Lausanne, Switzerland Manuel Serrano Spanish National Cancer Research Centre (CNIO) Madrid, Spain Radek Skoda Department of Biomedicine Experimental Hematology University Hospital Basel Basel, Switzerland Bart Staels Université Lille Nord de France Lille, France Inserm, U1011 Lille, France UDSL Lille, France Institut Pasteur de Lille Lille, France Karen P. Steel Wellcome Trust Sanger Institute

Wellcome Trust Genome Campus Hinxton, Cambridge, United Kingdom Anne Tailleux Université Lille Nord de France Lille, France Inserm, U1011 Lille, France UDSL Lille, France Institut Pasteur de Lille Lille, France Charles Thomas Center of Phenogenomics (CPG) Ecole Polytechnique Fédérale de Lausanne Lausanne, Switzerland Didier Trono School of Life Sciences and “Frontiers in Genetics” National Program Ecole Polytechnique Fédérale de Lausanne (EPFL) Lausanne, Switzerland Dorothy E. Vatner The University of Medicine & Dentistry of New Jersey New Jersey Medical School Newark, New Jersey Stephen F. Vatner The University of Medicine & Dentistry of New Jersey New Jersey Medical School Newark, New Jersey Sonia Verp School of Life Sciences and “Frontiers in Genetics” National Program Ecole Polytechnique Fédérale de Lausanne (EPFL) Lausanne, Switzerland Xavier Warot Institut Clinique de la Souris (ICS) Illkirch, France and Ecole Polytechnique Fédérale de Lausanne Lausanne, Switzerland Benedikt Wefers German Research Center for Environmental Health Munich, Germany Olivia Wendling Institut Clinique de la Souris (ICS) Illkirch, France and Institut de Génétique et de Biologie Moléculaire et Cellulaire CNRS/INSERM/Université Louis Pasteur Illkirch, France John N. Wood Molecular Nociception Group Wolfson Institute for Biomedical Research University College London London, United Kingdom and London Pain Consortium Kings College London London, United Kingdom Christine I. Wooddell Roche Madison Inc. Madison, Wisconsin

Wolfgang Wurst German Research Center for Environmental Health Munich, Germany, Technical University Munich Munich, Germany, Max-Planck-Institute of Psychiatry Munich, Germany, and Deutsches Zentrum für Neurodegenerative Erkrankungen e.V. (DZNE) Munich, Germany Guofeng Zhang School of Anatomy and Human Biology The University of Western Australia Crawley, Australia Xin Zhao The University of Medicine & Dentistry of New Jersey New Jersey Medical School Newark, New Jersey

Characterization and Validation of Cre-Driver Mouse Lines Franc¸oise Gofflot,1,3 Olivia Wendling,1,2 Nathalie Chartoire,1 Marie-Christine Birling,1 Xavier Warot,1,4 and Johan Auwerx1,4 1

Institut Clinique de la Souris (ICS), Illkirch, France Institut de G´en´etique et de Biologie Mol´eculaire et Cellulaire, CNRS/INSERM/Universit´e Louis Pasteur, Illkirch, France 3 Universit´e Catholique de Louvain, Life Science Institute, Louvain-la-Neuve, Belgium 4 Ecole Polytechnique F´ed´erale de Lausanne, Lausanne, Switzerland 2

ABSTRACT Conditional gene manipulations in mice are increasingly popular strategies in biomedical research. These approaches rely on the production of conditional genetically engineered mutant mouse (GEMM) lines with mutations in protein-encoding genes. These conditional GEMMs are then bred with one or several transgenic mouse lines expressing a site-specific recombinase, most often the Cre recombinase, in a tissue-specific manner. Conditional GEMMs can only be exploited if Cre transgenic mouse lines are available to generate somatic mutations, and thus the number of Cre transgenic lines has significantly increased over the last 15 years. Once produced, these transgenic lines must be validated for reliable, efficient, and specific Cre expression and Cre-mediated recombination. In this overview, the minimum level of information that is ideally required to validate a Cre-driver transgenic line is first discussed. The vagaries associated with validation procedures are considered next, and some solutions are proposed to assess the expression and activity of constitutive or inducible Cre recombinase before undertaking extensive breeding C 2011 by John experiments and exhaustive phenotyping. Curr. Protoc. Mouse Biol. 1:1-15  Wiley & Sons, Inc. Keywords: site-specific recombination r conditional mutagenesis r inducible Cre r functional genomic

INTRODUCTION Much of the recent progress in mammalian functional genomics has been driven by the use of genetically engineered mutant mouse (GEMM) lines. Informative mutations can now be generated in almost any mouse gene, either through classic gene targeting (conventional germline knockouts) or, increasingly, through conditional gene targeting, a strategy that allows temporal and spatial control of the onset of gene ablation/modification (Lewandoski, 2001; Metzger and Chambon, 2001; Branda and Dymecki, 2004; Argmann et al., 2005). The most successful approach for conditional gene targeting is based on the Cre-loxP system (Sauer and Henderson, 1989; Lakso et al., 1992; Rajewsky et al., 1996; Nagy, 2000; Collins et al., 2007; Birling et al., 2009), in which the allele of interest is flanked by recognition sites for the Cre DNA recombinase, the loxP sites. When such “floxed” mice are bred with transgenic mice expressing the Cre recombinase in a tissue-specific fash-

ion, the gene of interest is knocked out/altered only in this particular tissue. An added sophistication is the inclusion of temporal control, which can be achieved using ligand-activated chimeric recombinases composed, for instance, of the fusion of the Cre recombinase with the ligand-binding domain of a mutated form of the estrogen receptor (ER), which can be activated only by synthetic ER ligands (e.g., tamoxifen), but not by natural estrogen-like compounds (Feil et al., 1996; Danielian et al., 1998; Schwenk et al., 1998; Vasioukhin et al., 1999; Metzger and Chambon, 2001; Hayashi and McMahon, 2002). This strategy avoids problems with early lethality, developmental effects, and compensatory mechanisms, which are often apparent in classical germline or somatic knockout models. In 2006 large collaborative research efforts were launched by the European Commission, the U.S. National Institutes of Health (NIH), and Genome Canada to establish libraries of mutant mouse ES cell lines, each of which

Current Protocols in Mouse Biology 1: 1-15, March 2011 Published online March 2011 in Wiley Online Library (wileyonlinelibrary.com). DOI: 10.1002/9780470942390.mo100103 C 2011 John Wiley & Sons, Inc. Copyright 

Cre-Driver Mouse Lines

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establishment

carries an altered or “floxed” allele of a single gene (Austin et al., 2004; Auwerx et al., 2004; Collins et al., 2007). These mutant ES cell mutations can be readily transformed into mice using blastocyst injection, and the mutation activated by crossing the mouse bearing the floxed allele with a Cre-driver strain to induce the mutation in spatially and temporally determined patterns. The full power of conditional GEMMs, however, can only be exploited if transgenic mouse lines expressing the Cre recombinase in a tissue-, organ-, and cell-type-specific manner are available to allow the creation of somatic mutations. When searching the literature, investigators will find many Cre-driver transgenic lines that have been used successfully. However, for multiple reasons, data available for mouse Cre lines are often incomplete. Ideally, a minimum level of information should be available to users to allow selection of appropriate Cre transgenic lines for genedeletion experiments–specifically: (1) specificity and efficiency of Cre expression and Cre-mediated deletion; (2) reproducibility of the deletion from animal to animal for the same floxed allele; (3) reproducibility of

transgenic founder transgenic line step 1: Cre expression

validation

the deletion with different floxed alleles; (4) timing of Cre expression and Cre-mediated deletion for noninducible Cre mouse lines; (5) kinetics and efficiency of Cre induction and absence of leakage for inducible Cre mouse lines; and (6) phenotypes caused by either integration-mediated mutagenesis, by Cre “toxicity,” or by passenger genes in the construct. Although community efforts are underway to fully characterize newly produced Credriver mice (e.g., CREATE European Project, http://dev.creline.org/home; Mouse Clinical Institute, http://www.ics-mci.fr/crezoo.html), this ideal level of information is typically not known for currently available mice. To ensure the correct interpretation of resulting phenotypes, it is thus up to investigators to carefully verify the most critical parameters before setting up their experiment. In that context, this overview discusses critical parameters associated with production, validation, and use of Cre-driver transgenic lines, and presents some simple assays that can be used for characterization of these mouse lines. These assays can be combined to characterize newly produced strains, ultimately streamlining the establishment of

no

RT-qPCR

characterization

breeding

yes

G2 - tg

step 2 x Reporter line

no

qPCR

discard

yes

G2 - tg

step 3

step 3: Cre activity anatomical level

x colorimetric reporter line

step 4: phenotyping

clinical phenotyping SHIRPA/biochemistry

histology (␤-gal –AP)

upload information in database

Cre-Driver Mouse Lines

G1 - tg

analysis

discard step 2: Cre activity DNA level

G0 - tg

G2 - tg

ready to be used

Figure 1 Flow chart combining simple assays allowing one to characterize Cre recombinase expression and activity in transgenic Cre lines from the genomic to the cellular level. Each step can also be applied to complete or confirm available data on Cre-driver lines. G, generation; tg, transgenic; β–gal, β-galactosidase; AP, alkaline phosphatase; RT-qPCR, quantitative real-time reverse transcriptase PCR; qPCR, quantitative PCR.

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homogeneously validated Cre lines, or individual assays can be applied to existing lines for which available information is incomplete and/or requires confirmation. A flow chart summarizing the appropriate application of the strategies described in this article is shown in Figure 1.

SPECIFICITY AND EFFICIENCY OF Cre EXPRESSION Because the key feature of conditional gene targeting is its spatial or temporal restriction, the first parameter to control is the fidelity of Cre recombinase expression. Indeed, all systems used to generate tissue-specific Creexpressing mice rely on appropriate promoters and/or enhancers to control the expression

of Cre in a specific cell lineage. However, recombination could occur in cells outside of the desired target tissue due to unexpected or inappropriate expression of the transgene. This can be due to incomplete information about the gene whose promoter is used to drive recombinase expression, a transgene-insertion effect, or, in the case of inducible Cre, a “leakage effect” of the construct in other tissues. A broad, rapid, and cost-effective screen to verify the full expression pattern of the Cre recombinase in the selected transgenic lines can be performed by quantitative real-time reverse transcriptase PCR (RT-qPCR). By testing a large range of organs, information on ectopic or unexpected expression can be obtained rapidly before starting crosses with the floxed-allele transgenic mice.

Table 1 List of 25 Samples for Analysis of Cre Expression by RT-qPCR

System

Organ

Vascular

Aorta Heart

Digestive

Jejunum Colon Liver Pancreas Stomach

Skeletal

Bone

Metabolism

BATa WATb Muscle

Respiratory

Lung

Hematopoietic

Spleen Skin

Nervous

Olfactory bulbs Cortical and subcortical area Hypothalamus Hippocampus and thalamus Cerebellum Brainstem Spinal Cord

Urogenital

Ovary/Testis Kidney

Sensory a BAT, brown adipose tissue. b WAT, white adipose tissue.

Eye Cre-Driver Mouse Lines

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A pX

ERT2

Cre

RT-qPCR Primers

pA

␤-globin

Ex Ex

In

Ex

B E10

E9

E9 Cre excision

*

Marker loxP

E10

WT allele

RXR␣⌬AF2(LNL)

LNL allele

RXR␣⌬AF2(L)

L allele

loxP

*

E9

RXR␣

E10 loxP

Figure 2 (A) Schematic representation of the CreERT2 transgene used for pronuculei injection (Feil et al., 1997; Indra et al., 1999). The pair of primers used for RT-qPCR amplification of Cre transcripts is located in the β-globin region between a tissue-specific promoter X (pX) and the CreERT2 gene. (B) RXRαAF2(LNL) mouse line. This line is used as a floxed reporter line for determination of Cre recombinase activity by qPCR. A floxed Neo cassette is inserted between exon 9 and 10 of the RXRα gene, and a mutation (*) is present in exon 10 (Mascrez et al., 1998). Three pairs of primers have been designed that allow specific amplification of the wild-type (WT, green), floxed (LNL, blue), and excised (L, pink) alleles. Ex, exon; In, Intron; pA, polyadenylation site; pX, promoter of the gene X.

Experiments can be performed on one male and one female per line, with a minimum number of 25 samples, as suggested in Table 1, in order to cover the major body systems. However, depending on the expected tissue specificity of Cre recombinase expression, additional samples can be added. According to the Cre transgene used to generate the selected Cre-driver line, a set of primers is designed in a common part of the Cre cassette. In the case of the CreERT2 cassette, which has been largely used to produce tamoxifen-inducible Cre expression (Feil et al., 1997; Indra et al., 1999), specific primers can be designed in the β-globin intron (Fig. 2A). As a control of sensitivity, the expression of the endogenous gene whose promoter is used to drive Cre expression should also be measured. Standard validated procedures for RT-qPCR can be applied, as described elsewhere (Bookout et al., 2006; Gofflot et al., 2007; http://empress.har. mrc.ac.uk/), and in accordance with Minimum

Cre-Driver Mouse Lines

Information for Publication of Quantitative Real-Time PCR Experiments (MIQE) guidelines (Bustin et al., 2009; http://www.genequantification.de/miqe.html). As an example of the predictive value of this assay, analysis in the Ppm1a-CreERT2 line is described here, in which CreERT2 expression is driven by the promoter of the ubiquitous “protein phosphatase 1A, magnesiumdependent, alpha isoform” gene (LifschitzMercer et al., 2001). Also described is the Vil1-CreERT2 mouse line, in which the recombinase is targeted to the epithelial cells of the intestinal crypts (Meseguer and Catterall, 1987; Pinto et al., 1999; Robine et al., 1997). Both lines are produced and available at the Mouse Clinical Institute (http://www.icsmci.fr/crezoo.html). Several lines generated after microinjection of the same construct were analyzed, and both the expression pattern and expression level were compared. In the Ppm1a-CreERT2 line, Cre transcripts were

Figure 3 (figure appears on next page) Characterization of Cre expression in the ubiquitous Ppm1a-CreERT2 (A, B) and digestive tract-specific Vil1-CreERT2 (C) mouse lines. (A, C) Comparison of relative Cre expression between different transgenic lines (N is at least 2 for each line) determined by RT-qPCR in 25 tissue samples. (B) Comparison of relative expression of Cre versus the endogenous Ppm1a mRNAs (N = 2).

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0

Figure 3 Line A

Current Protocols in Mouse Biology Muscle Cortex-subcortex

BAT WAT Muscle Cortex-subcortex

WAT

Muscle

Cortex-subcortex

Cerebellum Olfactory bulbs Spinal cord Hypothalamus Hippocampus-thalamus Brainstem Ovary Testis Lung Eye Kidney

Cerebellum Olfactory bulbs Spinal cord Hypothalamus Hippocampus-thalamus Brainstem Ovary Testis Lung Eye Kidney

Cerebellum

Olfactory bulbs

Spinal cord

Hypothalamus

Hippocampus-thalamus

Brainstem Ovary Testis Lung Eye

Kidney

WAT

BAT

Skin

Spleen

Line C

Bones

Stomach

Line B

Pancreas

Liver

Colon

Skin

20

BAT

40

Skin

60 Spleen

80

Spleen

Line B Bones

100

Bones

20

Stomach

40

Stomach

60

Liver

80

Pancreas

Ppm1a

Liver

100

Pancreas

Colon

Duodenum

Heart

Relative expression (%)

Line A

Colon

Duodenum

Cre

Duodenum

C Heart

B

Heart

0 Aorta

0

Aorta

Relative expression (%)

100

Aorta

Relative expression (%)

A Line D

80

60

40

20

(legend appears on previous page)

Cre-Driver Mouse Lines

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amplified in 25 RNA samples obtained from two animals per line. Relative Cre expression was easily detected in most of the 25 samples analyzed, confirming the ubiquitous nature of the promoter selected (Fig. 3A). One transgenic line (line C) expressed the transgene at a higher level, with little variation between individuals and organs. In these lines, the expression level of Cre mRNA was close to that of the endogenous Ppm1a mRNA, as illustrated in Figure 3B. Notably the testis, an organ expressing high levels of the Ppm1a mRNA, expressed the highest amount of CreERT2 mRNA. Cre, and Ppm1a mRNAs were detected at very low levels in the pancreas, a tissue notorious for its high content of RNase, making RNA extraction challenging (Chirgwin et al., 1979). In contrast, amplification of Cre transcripts in 24 organs from two different Vil1-CreERT2 mouse lines revealed significant Cre mRNA expression only in the jejunum, colon, and testis (Fig. 3C), with relative levels of Cre mRNA being higher in line B. For both lines, the level of Cre expression was in the same range as the level of the endogenous Vil1 mRNA (data not shown). Importantly, this analysis revealed the presence of Cre transcripts in the testis, a site of expression that was not expected for the Vil1 promoter. Infidelity of Cre expression and recombination in the germline has previously been reported by others (Schmidt-Supprian and Rajewsky, 2007). This example emphasizes the importance of verifying ectopic/unexpected expression to avoid misinterpretation of phenotype due to recombination in other cells than the desired target.

SPECIFICITY AND EFFICIENCY OF Cre-MEDIATED DELETION

Cre-Driver Mouse Lines

Although RT-qPCR can provide easy and sensitive detection of Cre expression, the information most needed by investigators is about Cre-mediated recombination. As opposed to Cre expression analysis, Cre activity can only be tested in animals that have been crossed with mouse lines harboring a floxed allele and, for inducible Cre lines, that have been injected either with the inducer or vehicle. Although for most published Cre lines recombination properties have been validated by reporter gene studies, users should be aware that the efficiency of recombination can be locus dependent, and, therefore, the recombination pattern obtained with a particular reporter gene does not necessarily predict that of other floxed genes (see Vooijs et al., 2001; J. Becker and B. Kieffer, pers. comm.). Indeed,

the chromatin structure at the locus of interest, the state of DNA methylation, and the transcriptional activity seem to affect the efficiency of recombination. In addition, it has been reported that the ability of a floxed target gene to be recombined could also vary between cell types (Kellendonk et al., 1999). This was potentially explained by differential accessibility of Cre to loxP sites due to cell type- and development-specific chromatin conformations. Before starting an extensive phenotypic analysis, it is thus mandatory to monitor recombination at the target locus. The most common procedures used to examine the pattern of Cre-mediated recombination in various tissues are Southern blot analysis and simple PCR. Although these procedures are robust, they do not provide a quantitative evaluation of the recombination and are of restricted sensitivity, especially when limited samples are available. An alternative to score for both the efficiency and specificity of Cre recombinase deletion at the DNA level is quantitative PCR. This procedure can (i) provide information on Cre excision efficiency with high sensitivity and reproducibility, (ii) evaluate the reproducibility of deletion from animal to animal on the same floxed alleles, (iii) evaluate the reproducibility of deletion between different floxed lines and, (iv) for inducible Cre lines, verify the efficiency of the tamoxifen induction on the CreERT2 transgene activity. This analysis of Cre recombination activity is performed in animals that have been crossed with any mouse lines harboring a floxed allele, either a so-called “reporter line” or the transgenic line with the floxed allele of interest. As an example, at the Mouse Clinical Institute, the reporter line used to test the recombinase excision activity by qPCR is a floxed RXRα transgenic line, the RXRαAF2 (LNL) (Mascrez et al., 1998; Fig. 2B). Crosses between Cre lines and this reporter line are set up to obtain double transgenic mice, and three pairs of primers are used to specifically amplify the wild-type (WT), floxed (LNL), and excised (L) alleles (Figure 2B). In the case of CreERT2 transgenic animals, bigenic mice are injected either with tamoxifen or vehicle before analysis. For this step, the selection of samples can be made on the basis of the RT-qPCR data, if available—i.e., only positive Cre expression samples are analyzed. Additional samples of one positive system may also be analyzed indepth, e.g., more segments of the digestive tract can be analyzed as illustrated below in the case of the Vil1-CreERT2 .

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A 100% 80% 60% 20% 0% % WT

% LNL

%L

100%

Kidney

Ovary/testis

Spinal cord

Celebellum

Brain

Muscle

Skin

Pancreas

Colon

40% 20% 0%

Liver

80% 60%

Aorta

Proportion of alleles

40%

B 100% 80%

Proportion of alleles

60% 40% 20% 0% % WT

% LNL

%L

100% 80% 60% 40%

Kidney

Ovary/testis

Spinal cord

Celebellum

Cortical and subcortical

Muscle

Tail skin

Pancreas

Liver

Colon

lleum

Jejunum

Duodenum

0%

Aorta

20%

Figure 4 Determination of Cre-mediated excision by qPCR in the ubiquitous Ppm1a-CreERT2 (A) and digestive tract-specific Vil1-CreERT2 (B) mouse lines. Comparison of the percentage of excised allele (L) versus floxed (LNL) and wild type (WT) allele in double transgenic animals Ppm1a-CreERT2 /RXRαAF2(LNL) and Vil1-CreERT2 /RXRαAF2(LNL) mice injected with vehicle (top) or with tamoxifen (bottom). For tamoxifen injections, tamoxifen (Sigma, cat. no. T56648) was prepared at 10 mg/ml in sunflower seed oil (Sigma, cat. no. S5007). Intraperitoneal injection of 100 μl of this solution was performed for 5 consecutive days (1 mg/mouse/day) with mice aged 10 weeks old, and whose weight was >20 g. Identical amounts of sunflower seed oil (vehicle) were administered following the same protocol to control mice.

To illustrate the predictive value and sensitivity of this test, Figure 4 shows the analysis of the Ppm1a-CreERT2 and Vil1-CreERT2 mouse lines. In samples dissected from Ppm1aCreERT2 /RXRαAF2 (LNL) mice injected with vehicle, only the WT and floxed alleles were present, while the excised allele was detected in all the 11 tissues analyzed from tamoxifeninjected double transgenic animals (Fig. 4A). The proportion of excision ranged from 5% to

43%, the WT allele being 50%. The highest level of excision was observed in the skin and liver, and the lowest levels in the three samples from the central nervous system (CNS). The relatively low level of excision in the brain of the Ppm1a-CreERT2 mice, with regard to Cre mRNA expression levels, is most likely due to insufficient entrance of tamoxifen into brain cells. Indeed, induction of Cre activity in the brain seems to be slower than in other organs,

Cre-Driver Mouse Lines

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perhaps due to the blood-brain barrier and/or to the slower renewal of cells. For brain targeting, it is thus recommended to perform analysis 1 month, instead of 1 week, after the last injection of tamoxifen (Metzger and Chambon, 2001; Weber et al., 2001). In comparison, in bigenic Vil1-CreERT2 / RXRαAF2 (LNL) mice injected with tamoxifen, the excised allele was detected only in the digestive tract, at a relative proportion of 15% to 30% (Fig. 4B). No excised allele was detected in the reproductive organs of the selected line, despite the fact that Cre expression was measured by RT-qPCR in the testis. This could be due either to the sensitivity of this test or to lower tamoxifen access to this tissue.

ANATOMICAL PATTERN OF Cre-MEDIATED DELETION As most transgenic Cre lines are driven by cell-specific promoters, the required level of information for validation is the location

of Cre excision activity within specific functional or cellular compartments of an organ. The classic way to characterize Cre activity at the cellular level is to cross Cre-driver lines with colorimetric reporter lines, such as the ROSA26, ACZL, and ZAP reporter mouse lines (Fig. 5). In the ROSA26 reporter line, the ROSA26 allele is targeted with a Cre excision– conditional lacZ reporter (Soriano, 1999). In the ACZL reporter line, a floxed CAT transcription unit prevents lacZ expression in absence of Cre-mediated recombination (Akagi et al., 1997). The Z/AP reporter line (Lobe et al., 1999) utilizes two reporters: the lacZ reporter marks cells before excision occurs, while the heat-resistant human placental alkaline phosphatase (hAP) marks cells after Cre-mediated DNA excision. As for the qPCR test, the analysis is performed on samples dissected from double transgenic animals. However, samples need first to be embedded, sectioned, and stained for either Xgal and/or hAP.

No Cre

ROSA26

pROS A26

PGK Neo pA

loxP

LacZ

Cre

no staining

pA

Ref

Soriano (1999)

loxP

pROS A26

LacZ

-Gal

pA

loxP

p Actin

CAT

pA

ACZL

loxP

p Actin

LacZ

no staining

pA

Akagi et al. (1997)

loxP

LacZ

pA

LacZ

pA

-Gal

loxP

eCMV

p Actin

Z/AP

loxP

eCMV

p Actin

AP

pA

-Gal

Lobe et al. (1999)

loxP

AP

pA

AP

loxP

Figure 5 Schematic representation of the construction and activity of the three most popular colorimetric reporter lines used to test for Cre activity in Cre-driver transgenic lines. β-gal, β-galactosidase; AP, alkaline phosphatase; CMV, cytomegalovirus; PGK, phosphoglycerate kinase; CAT, chloramphenicol acetyltransferase; pA, polyadenylation site. Cre-Driver Mouse Lines

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Figure 6 Characterization of Cre activity at histological level in the Ppm1a-CreERT2 mice line. (A) Cre-mediated expression of the reporter gene β-galactosidase in nine organs dissected from double-transgenic Ppm1a-CreERT2 /ROSA26 mice injected with tamoxifen. (B) XGal (ROSA, ACZL) and hAP (Z/AP) staining of muscle sections revealing (i) the reporter expression pattern in three different colorimetric reporter lines crossed with a CMV-Cre deleter mice (bottom row) (Dupe et al., 1997), and (ii) the localization of Cre activity in Ppm1a-CreERT2 mice crossed with each of these reporter lines (top row).

In the case of CreERT2 transgenic animals, bigenic mice are also injected either with tamoxifen or with vehicle 1 week or 1 month before sample collection. The selection of samples could be made on the basis of the RT-qPCR and/or qPCR analysis; in that case, only tissues in which positive Cre expression and/or recombination activity have been ob-

served are further analyzed, limiting the number of samples to be processed. However, for mouse lines in which Cre is targeted not only to a particular tissue but to a specific cell type that may represent only a very small proportion of the organ cellular population, the sensitivity of RT-qPCR or qPCR may be limited, and it is advisable to confirm negative results at the

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histological level to avoid discarding potentially valuable lines. A matter of concern with this procedure is the potential lack of ubiquitous expression of the reporter in existing reporter lines, especially at adult stages. In addition, tissue-specific Cre lines generated by pronuclear injection often exhibit mosaic expression of the recombinase (Schwenk et al., 1998), meaning that only sub-regions of the tissue/organ show active recombination. To illustrate these points, Cre activity was characterized in detail at the cellular level in Ppm1aCreERT2 mice crossed with the ROSA26 reporter line. Bigenic Ppm1a-CreERT2 /ROSA26 mice were injected with either tamoxifen or vehicle. Xgal staining of sections revealed the activity of the Cre recombinase in all but one organ, the muscle, analyzed in tamoxifeninjected double transgenic mice (Fig. 6A). In all organs positively labeled, the area of excision within an organ was mosaic and the cell excision within a tissue was not complete. The liver and skin displayed the highest level of excision, while the lowest level was observed in the brain, in line with the qPCR analysis. No significant Cre activity was detected in mice injected with vehicle (data not shown). The absence of staining observed in the muscle was unexpected in light of the Cre expression and activity data. To confirm this observation, Cre recombination activity was evaluated, and detected, in the muscle after crosses with the ACZL and ZAP reporter mouse lines (Fig. 6B, bottom row). This analysis revealed that absence of Xgal staining in the muscle of Ppm1a-CreERT2 /ROSA26 mice was associated with the ROSA26 reporter line and not with absence of Cre recombination activity. This could be due to the absence or weak expression of the ROSA26 reporter in this tissue, a hypothesis supported by the higher Cre excision level observed in the ACZL and ZAP reporter mouse lines, which express higher levels of the reporter gene in skeletal muscle, as evaluated after crossing with a ubiquitous deleter, the CMV-Cre mouse line (Dupe et al., 1997; Fig. 6B, top row). The cellular characterization of Cre activity in the Ppm1a-CreERT2 underscores the value of using different reporter mouse lines according to the targeted organ/tissue. Among the colorimetric reporter lines classically used in the literature, a preliminary comparison indicated that the ROSA26 line seems the most adequate for the majority of promoters, as it showed reporter activity in the larger number

of organs evaluated after crossing with a ubiquitous Cre-deleter mouse line, while the ACZL and Z/AP displayed expression in a more restricted number of tissues (O. Wendling and D. Metzger, unpub. observ.). Our study, however, revealed the usefulness of these two lines for tissue-specific analysis, e.g., in the skeletal muscle. To address the problem of the lack of ubiquitousness of reporter lines, detection of Cre mRNA or Cre protein at the anatomical level by in situ hybridization (ISH) or immunohistochemistry (IHC) can be used. These two procedures are discussed below.

ADDITIONAL OR ALTERNATIVE PROCEDURES TO DETECT Cre EXPRESSION To map Cre expression at the anatomical level, nonradioactive in situ hybridization (ISH) using digoxigenin-labeled probes can be performed in a standard 5-step procedure: hybridization of the probe to pretreated tissue at 65◦ C; stringent post-hybridization washes; blocking steps to prepare for the immunodetection; primary antibody anti-DIGAP incubation; and colorimetric alkaline phosphatase detection (Chotteau-Lelievre et al., 2006; Gofflot et al., 2007; Knoll et al., 2007). Although this procedure may require a higher level of expertise, it also has several advantages. It can be used in parallel with, or in place of, the RT-qPCR, as it scores for Cre expression and does not require crosses with a transgenic floxed line. As such, it is thus totally independent of any reporter activity, and also independent with respect to cellular access of inducer such as tamoxifen. It can be combined with other detection procedures, either by double in situ hybridization or immunohistochemistry, allowing precise characterization of the cellular population targeted by Cre expression. For those less familiar with ISH, immunodetection procedures for Cre protein have been previously described (Kaelin et al., 2006; Knoll et al., 2007). To illustrate these two procedures, two inducible tissue-specific Cre lines were used: the Tph2-CreERT2 (gift of P. Chambon and D. Metzger) and the Ins1-CreERT2 , in which Cre expression is targeted to highly restricted cell populations, the raphe serotonergic neurons and the β-cells of pancreatic islets, respectively. As illustrated in Figure 7, ISH was successfully used to detect Cre mRNA in the raphe nuclei of the Tph2-CreERT2 mice (Fig. 7A). Cre expression was similar to lacZ staining on brain sections from tamoxifen-injected

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Figure 7 Characterization of Cre activity at histological level in the Tph2-CreERT2 and Ins1CreERT2 mouse lines. (A) Sections through the pons, midbrain, and medulla revealing Cre activity in the raphe nuclei through Cre-mediated expression of the reporter gene β-galactosidase (Tph2CreERT2 /ROSA26 mice injected with tamoxifen, top row) and Cre expression through ISH with a specific Cre probe (Tph2-CreERT2 mice, bottom row). (B) IHC detection of Cre protein and Xgal staining of pancreas sections of tamoxifen or vehicle-injected Ins1-CreERT2 /ROSA26 mice revealed specific Cre protein content and Cre activity restricted to the islets of Langerhans. ISH procedures have been described elsewhere (Chotteau-Lelievre et al., 2006; Gofflot et al., 2007). For IHC, primary rabbit anti-Cre (1:8000 dilution, VWR, cat. no. 69050-3) was used with goat antirabbit antibody coupled to horseradish peroxidase (1:100 dilution; Invitrogen, cat. no. G-21234) as secondary antibody. After washing, Cre was visualized by FITC-tyramide amplification (1/50, 30-min incubation) (PerkinElmer, cat. no. SAT701B).

Tph2-CreERT2 /ROSA26 mice, revealing that all 9 nuclei of the raphe in the midbrain and medulla were specifically labeled. As demonstrated by the analysis of the Ppm1a-CreERT2 line, the pancreas is an organ for which the isolation of RNA is particularly challenging (Chirgwin et al., 1979) and Cre mRNA could not be detected reliably in pancreas samples of Ins1CreERT2 mice. In that particular case, IHC was used and allowed the detection of Cre protein in the islets of Langerhans of the

Ins1-CreERT2 mice (Fig. 7B). This result was further confirmed at the histological level in Ins1-CreERT2 /ROSA26 mice, in which specific Cre-mediated excision was detected by lacZ staining in the islets of Langerhans of the pancreas.

PHENOTYPIC CHARACTERIZATION OF Cre LINES Transgenic lines produced by conventional transgenesis can develop unexpected

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phenotypes due to integration-mediated mutagenesis or passenger genes in the construct (Lusis et al., 2007). In addition, a high level of Cre protein expression can result in cellular toxicity (Forni et al., 2006; Schmidt-Supprian and Rajewsky, 2007). In that context, successfully characterized lines should also be subjected to a final functional test. Indeed, functional abnormalities in Cre mice could be a confounding factor for the interpretation of the phenotype observed when that Cre line is used to delete a gene of interest. When producing a new Cre-driver transgenic line, a standard and simple behavioral, biochemical, and metabolic phenotyping procedure would allow one to discard Cre mouse lines with interfering phenotypes before distribution and archiving. First, to evaluate the general health and basic neurological status, the modified SHIRPA protocol (http://empress.har.mrc.ac.uk/browser/) can be used, as it is a rapid, high-throughput non-invasive and non-stressful test suited for a global evaluation of the phenotype (Mandillo et al., 2008). Second, clinical and

basal metabolic parameters in G2 mice maintained under basal chow–fed conditions should be monitored (Champy et al., 2004). Finally, some specific tests could be used to evaluate the functions of the organ(s) to which Cre expression is specifically targeted. For example, blood pressure, heart rate, heart weight, and histology could be specifically evaluated in mice with Cre targeted to cardiac muscle, while rotarod test, grip strength, endurance running, and muscle histology could be investigated in mice with Cre targeted to skeletal muscle (see http://empress.har.mrc.ac.uk/browser/ for detailed procedures). Instructive of the importance of such a phenotypic characterization, the Vil1-CreERT2 mice have a severe functional abnormality which is due to the presence of a passenger gene, the G-protein coupled receptor Tgr5 (Thomas et al., 2008) in the BAC construct, precluding its use for metabolic studies (Fig. 8A). Tgr5 mRNA is 5-fold overexpressed in the Vil1-CreERT2 mouse line

A BAC RP23-278N11 Chromosome 1 :74,302,024-74,495,075

193.05 kb

Pnkd ENSMUSG00000026179 Arpc2 ENSMUSG00000006304

Vil 1 ENSMUSG00000026175 Ctdsp1 ENSMUSG00000026176

Tgr5 (Gpbar1) ENSMUSG00000064272 Gm216 ENSMUSG00000073650 Aamp ENSMUSG00000006299 Slc11a1 ENSMUSG00000026177 Tmbim1 ENSMUSG00000006301 Usp37 ENSMUSG00000033364

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Vil1-CreERT2

Figure 8 Phenotypic analysis of the Vil1-CreERT2 transgenic line. (A) Schematic representation of the mouse BAC used for the construction of the Vil1-CreERT2 transgene. (B) Expression of mRNA levels of the GPCR Tgr5 in the ileum of control and Vil1-CreERT2 transgenic mice. (C) Mean ± SD of the area under the curve during an oral glucose tolerance test in control (N = 8) and Vil1-CreERT2 (N = 8) mice fed a high-fat, high-sucrose diet for 12 weeks. (D-E) Insulin and GLP-1 levels measured in the serum 15 min after the administration of the oral glucose load in the control and Vil1-CreERT2 transgenic mice of panel (B). Cre-Driver Mouse Lines

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(Fig. 8B). As Tgr5 controls the expression of GLP-1 in entero-endocrine cells (Thomas et al., 2009), which in turn improves insulin secretion, Vil1-CreERT2 mice have improved glucose tolerance with a reduced area under the curve (AUC) in oral glucose tolerance tests (Fig. 8C). The higher insulin release as a consequence of the increase of GLP-1 levels after a glucose challenge explains the improved glucose tolerance (Fig. 8D-E). In other cases, the phenotype could be the consequence of insertional mutagenesis. In such conditions, it can be worth testing Cre lines derived from another founder.

SUMMARY Advances in the sophisticated manipulation of the mouse genome now allow the generation of mutant mice for disease modeling and functional analysis by conditional mutagenesis. In the coming years, site-specific recombination transgenic mice will become necessary tools for most scientists to generate conditional mutations. Although a few initiatives attempt to establish and/or index a range of transgenic mouse lines that express Cre recombinase in different tissues (e.g., Gensat, http://www.gensat.org/index. html; CreXMice, http://nagy.mshri.on.ca/cre/; MUGEN Mutant Mice database, http://bioit. fleming.gr/mugen/mde.jsp; CREATE, http:// dev.creline.org/home), most of the current data are disparate, heterogenous, and incomplete. It will thus be up to Cre users to thoroughly characterize and/or validate the appropriate Cre-driver mouse lines before undertaking extensive breeding experiments and exhaustive mutant analyses that could otherwise lead to inconclusive or incorrect conclusions. As already mentioned, investigators will have to test their selected Cre lines on their own conditional mutants, as efficiency of Cre-mediated excision varies from one allele to another. When bigenic mice are generated and phenotyped, it is mandatory to check in parallel that Cre-mediated excision has indeed occurred in the organ of interest. In this overview, a set of basic assays is described that can be used either to confirm existing data or as a streamlined and standardized characterization scheme for newly established Cre lines. Some of the artifacts and problems that can be associated with Cre-driver mouse lines are also discussed, with emphasis on the importance of the characterization steps. Finally, the more a given Cre line is used, the more of these issues will be evaluated.

Investigators using Cre mouse lines are therefore encouraged to upload their information onto databases where all information is centralized and available to the scientific community, as exemplified by the Cre-X-Mice database (Nagy et al., 2009).

ACKNOWLEDGMENTS The authors acknowledge Fabrice Aug´e, Graziella Neau, Leila El Fertak, and the MCI histopathology service for technical assistance. We are indebted to Kristina Schoonjans and Charles Thomas for the phenotypic analysis of the Vil1-CreERT2 mouse line and to Jabier Gallego-Llamas for IHC studies. We wish to thank Jean-Louis Mandel, Guillaume Pavlovic, Lydie Venteo, and Thomas F. Vogt for critical reading of the manuscript. The “CreZoo” project at the MCI was initiated under the auspices of Professors Pierre Chambon and Daniel Metzger. This work was supported in part by an INCa grant and by the EUCOMM European integrated project.

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Birling, M.C., Gofflot, F., and Warot, X. 2009. Sitespecific recombinases for manipulation of the mouse genome. Methods Mol. Biol. 561:245263. Bookout, A.L., Cummins, C.L., Mangelsdorf, D.J., Pesola, J.M., and Kramer, M.F. 2006. High-throughput real-time quantitative reverse transcription PCR. Curr. Protoc. Mol. Biol. 73:15.8.1-15.8.28. Branda, C.S. and Dymecki, S.M. 2004. Talking about a revolution: The impact of site-specific recombinases on genetic analyses in mice. Dev Cell 6:7-28. Bustin, S.A., Benes, V., Garson, J.A., Hellemans, J., Huggett, J., Kubista, M., Mueller, R., Nolan, T., Pfaffl, M.W., Shipley, G.L., Vandesompele, J., and Wittwer, C.T. 2009. The MIQE guidelines: Minimum information for publication of quantitative real-time PCR experiments. Clin. Chem. 55:611-622. Champy, M.F., Selloum, M., Piard, L., Zeitler, V., Caradec, C., Chambon, P., and Auwerx, J. 2004. Mouse functional genomics requires standardization of mouse handling and housing conditions. Mamm. Genome 15:768-783. Chirgwin, J.M., Przybyla, A.E., MacDonald, R.J., and Rutter, W.J. 1979. Isolation of biologically active ribonucleic acid from sources enriched in ribonuclease. Biochemistry 18:5294-5299. Chotteau-Lelievre, A., Dolle, P., and Gofflot, F. 2006. Expression analysis of murine genes using in situ hybridization with radioactive and nonradioactively labeled RNA probes. Methods Mol. Biol. 326:61-87. Collins, F.S., Rossant, J., and Wurst, W. 2007. A mouse for all reasons. Cell 128:9-13. Danielian, P.S., Muccino, D., Rowitch, D.H., Michael, S.K., and McMahon, A.P. 1998. Modification of gene activity in mouse embryos in utero by a tamoxifen-inducible form of Cre recombinase. Curr. Biol. 8:1323-1326. Dupe, V., Davenne, M., Brocard, J., Dolle, P., Mark, M., Dierich, A., Chambon, P., and Rijli, F.M. 1997. In vivo functional analysis of the Hoxa1 3 retinoic acid response element (3 RARE). Development 124:399-410. Feil, R., Brocard, J., Mascrez, B., LeMeur, M., Metzger, D., and Chambon, P. 1996. Ligandactivated site-specific recombination in mice. Proc. Natl. Acad. Sci. U.S.A. 93:10887-10890. Feil, R., Wagner, J., Metzger, D., and Chambon, P. 1997. Regulation of Cre recombinase activity by mutated estrogen receptor ligandbinding domains. Biochem. Biophys. Res. Commun. 237:752-757. Forni, P.E., Scuoppo, C., Imayoshi, I., Taulli, R., Dastru, W., Sala, V., Betz, U.A., Muzzi, P., Martinuzzi, D., Vercelli, A.E., Kageyama, R., and Ponzetto, C. 2006. High levels of Cre expression in neuronal progenitors cause defects in brain development leading to microencephaly and hydrocephaly. J. Neurosci. 26:9593-9602. Cre-Driver Mouse Lines

Gofflot, F., Chartoire, N., Vasseur, L., Heikkinen, S., Dembele, D., Le Merrer, J., and Auwerx, J.

2007. Systematic gene expression mapping clusters nuclear receptors according to their function in the brain. Cell 131:405-418. Hayashi, S. and McMahon, A.P. 2002. Efficient recombination in diverse tissues by a tamoxifeninducible form of Cre: A tool for temporally regulated gene activation/inactivation in the mouse. Dev. Biol. 244:305-318. Indra, A.K., Warot, X., Brocard, J., Bornert, J.M., Xiao, J.H., Chambon, P., and Metzger, D. 1999. Temporally-controlled site-specific mutagenesis in the basal layer of the epidermis: Comparison of the recombinase activity of the tamoxifen-inducible Cre-ER(T) and Cre-ER(T2) recombinases. Nucleic Acids Res. 27:43244327. Kaelin, C.B., Gong, L., Xu, A.W., Yao, F., Hockman, K., Morton, G.J., Schwartz, M.W., Barsh, G.S., and MacKenzie, R.G. 2006. Signal transducer and activator of transcription (stat) binding sites but not stat3 are required for fasting-induced transcription of agouti-related protein messenger ribonucleic acid. Mol. Endocrinol. 20:2591-2602. Kellendonk, C., Tronche, F., Casanova, E., Anlag, K., Opherk, C., and Schutz, G. 1999. Inducible site-specific recombination in the brain. J. Mol. Biol. 285:175-182. Knoll, J.H., Lichter, P., Bakdounes, K., and Eltoum, I.-E. A. 2007. In situ hybridization and detection using nonisotopic probes. Curr. Protoc. Mol. Biol. 79:14.7.1-14.7.17. Lakso, M., Sauer, B., Mosinger, B. Jr., Lee, E.J., Manning, R.W., Yu, S.H., Mulder, K.L., and Westphal, H. 1992. Targeted oncogene activation by site-specific recombination in transgenic mice. Proc. Natl. Acad. Sci. U.S.A. 89:62326236. Lewandoski, M. 2001. Conditional control of gene expression in the mouse. Nat. Rev. Genet. 2:743755. Lifschitz-Mercer, B., Sheinin, Y., Ben-Meir, D., Bramante-Schreiber, L., Leider-Trejo, L., Karby, S., Smorodinsky, N.I., and Lavi, S. 2001. Protein phosphatase 2Calpha expression in normal human tissues: An immunohistochemical study. Histochem. Cell Biol. 116:31-39. Lobe, C.G., Koop, K.E., Kreppner, W., Lomeli, H., Gertsenstein, M., and Nagy, A. 1999. Z/AP, a double reporter for cre-mediated recombination. Dev. Biol. 208:281-292. Lusis, A.J., Yu, J., and Wang, S.S. 2007. The problem of passenger genes in transgenic mice. Arterioscler Thromb. Vasc. Biol. 27:2100-2103. Mandillo, S., Tucci, V., H¨olter, S.M., Meziane, H., Banchaabouchi, M.A., Kallnik, M., Lad, H.V., Nolan, P.M., Ouagazzal, A.M., Coghill, E.L., Gale, K., Golini, E., Jacquot, S., Krezel, W., Parker, A., Riet, F., Schneider, I., Marazziti, D., Auwerx, J., Brown, S.D., Chambon, P., Rosenthal, N., Tocchini-Valentini, G., Wurst, W. 2008. Reliability, robustness, and reproducibility in mouse behavioral phenotyping: A crosslaboratory study. Physiol. Genomics 34:243255.

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Mascrez, B., Mark, M., Dierich, A., Ghyselinck, N.B., Kastner, P., and Chambon, P. 1998. The RXRalpha ligand-dependent activation function 2 (AF-2) is important for mouse development. Development 125:4691-4707. Meseguer, A. and Catterall, J.F. 1987. Mouse kidney androgen-regulated protein messenger ribonucleic acid is expressed in the proximal convoluted tubules. Mol. Endocrinol. 1:535541. Metzger, D. and Chambon, P. 2001. Site- and timespecific gene targeting in the mouse. Methods 24:71-80. Nagy, A. 2000. Cre recombinase: The universal reagent for genome tailoring. Genesis 26:99109. Nagy, A., Mar, L., and Watts, G. 2009. Creation and use of a cre recombinase transgenic database. Methods Mol. Biol. 530:1-14. Pinto, D., Robine, S., Jaisser, F., El Marjou, F.E., and Louvard, D. 1999. Regulatory sequences of the mouse villin gene that efficiently drive transgenic expression in immature and differentiated epithelial cells of small and large intestines. J. Biol. Chem. 274:6476-6482. Rajewsky, K., Gu, H., Kuhn, R., Betz, U.A., Muller, W., Roes, J., and Schwenk, F. 1996. Conditional gene targeting. J. Clin. Invest. 98:600-603. Robine, S., Jaisser, F., and Louvard, D. 1997. Epithelial cell growth and differentiation. IV. Controlled spatiotemporal expression of transgenes: New tools to study normal and pathological states. Am. J. Physiol. 273:G759-G762. Sauer, B. and Henderson, N. 1989. Cre-stimulated recombination at loxP-containing DNA

sequences placed into the mammalian genome. Nucleic Acids Res. 17:147-161. Schmidt-Supprian, M. and Rajewsky, K. 2007. Vagaries of conditional gene targeting. Nat. Immunol. 8:665-668. Schwenk, F., Kuhn, R., Angrand, P.O., Rajewsky, K., and Stewart, A.F. 1998. Temporally and spatially regulated somatic mutagenesis in mice. Nucleic Acids Res. 26:1427-1432. Soriano, P. 1999. Generalized lacZ expression with the ROSA26 Cre reporter strain. Nat. Genet. 21:70-71. Thomas, C., Pellicciari, R., Pruzanski, M., Auwerx, J., and Schoonjans, K. 2008. Targeting bile-acid signalling for metabolic diseases. Nat. Rev. Drug Discov. 7:678-693. Thomas, C., Gioiello, A., Noriega, L., Strehle, A., Oury, J., Rizzo, G., Macchiarulo, A., Yamamoto, H., Mataki, C., Pruzanski, M., Pellicciari, R., Auwerx, J., and Schoonjans, K. 2009. TGR5-mediated bile acid sensing controls glucose homeostasis. Cell Metab. 10:167-177. Vasioukhin, V., Degenstein, L., Wise, B., and Fuchs, E. 1999. The magical touch: Genome targeting in epidermal stem cells induced by tamoxifen application to mouse skin. Proc. Natl. Acad. Sci. U.S.A. 96:8551-8556. Vooijs, M., Jonkers, J., and Berns, A. 2001. A highly efficient ligand-regulated Cre recombinase mouse line shows that LoxP recombination is position dependent. EMBO Rep. 2:292-297. Weber, P., Metzger, D., and Chambon, P. 2001. Temporally controlled targeted somatic mutagenesis in the mouse brain. Eur. J. Neurosci. 14:1777-1783.

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Standardized Post-Mortem Examination and Fixation Procedures for Mutant and Treated Mice Cristina Antal,1,4 St´ephanie Muller,1 Olivia Wendling,1,2 Yann H´erault,1,2 and Manuel Mark1,2,3 1

Mouse Clinical Institute, Illkirch, France Institut de G´en´etique et de Biologie Mol´eculaire et Cellulaire (IGBMC), CNRS/INSERM/Universit´e Louis Pasteur, Illkirch, France 3 Hˆopital Universitaire de Strasbourg, Strasbourg, France 4 Institut d’Histologie, Facult´e de M´edecine, Strasbourg, France 2

ABSTRACT A procedure for post-mortem examination (or necropsy) of mice is provided. The aim is to obtain a “holistic” picture of organs and systems at the anatomical and histological levels. The major issue is tissue preservation, which is achieved by rapid transfer into a fixative solution, usually neutral buffered formalin. Fixation is the first of the four basic steps in histopathological analyses of tissues, which also include embedding, sectioning, and staining. The protocols provided here describe routine methods for tissue fixation, as these methods are integral parts of any necropsy procedure. There is also a Strategic Planning section that addresses the overall approach to histopathological evaluation, as well as specifics such as age and gender of the mice, cohort size, and controls. Curr. C 2011 by John Wiley & Sons, Inc. Protoc. Mouse Biol. 1:17-53  Keywords: phenotyping r tissue collection r histology r necropsy r pathology

INTRODUCTION After establishing a phenotype in a mouse line through the use of in vivo tests, the subsequent challenge is to refine this phenotype at the organ, tissue, cell, and molecular (DNA, RNA, and protein) levels. This is most easily achieved after sacrificing the mice. This article focuses on collection of murine tissues for standard morphological analyses. A dissection for the purpose of post-mortem analysis, or necropsy, is undertaken to identify, at a macroscopic level (by the naked eye or with a dissecting microscope), morphological defects that characterize the mutant mouse and identify gross lesions that may contribute to morbidity and mortality. After the general condition and the body weight of the animal have been recorded, individual organs are removed, examined, sampled, and fixed in a systematic manner. All lesions should be described with regard to location, color, size, shape, consistency, distribution, and number or percent of involvement of a specific organ. Photographs of lesions provide documentation for records. Tissues that are collected for subsequent histopathological analyses require appropriate handling and preservation to prevent their deterioration. Once death has occurred, tissues undergo a process of self-digestion (autolysis). One major aim of necropsy is to avoid any unnecessary delay in collecting tissue samples, so that they can be optimally preserved through rapid immersion into a fixative solution. The article begins with a Strategic Planning section that addresses the approach to histopathological evaluation of mutant or treated mice. It then provides the information for collecting tissues to obtain a complete, standardized, macroscopic, and histological analysis of the mouse (see Basic Protocol 1). Proper fixation of murine tissues with formalin (see Basic Protocol 2) or Bouin’s solution (see Alternate Protocol) is also described. Current Protocols in Mouse Biology 1: 17-53, March 2011 Published online March 2011 in Wiley Online Library (wileyonlinelibrary.com). DOI: 10.1002/9780470942390.mo100118 C 2011 John Wiley & Sons, Inc. Copyright 

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STRATEGIC PLANNING NOTE: mutant mice are used hereafter as examples, but the proposed screen is applicable to mice treated with test compounds or tumor inducers as well. As histotechnology is time-consuming and labor-intensive, it is important to avoid unnecessary experiments while achieving the highest degree of comprehensiveness. To this end, it is best to perform a first-line screen followed by organ/pathology-specific targeted assays. An in-depth evaluation of organs that have been fixed, processed, and embedded in a systematic manner is used for first-line extended assays. Such assays involve readyto-use tissue samples and are rapidly applied after completion of the systematic screen. In contrast, secondary assays correspond to in-depth evaluations of organs requiring special fixation, tissue processing, and/or embedding procedures, and thus the generation of a new cohort of mice. A first-line histological screen is aimed at detecting the broadest array of tissue abnormalities in the mutant mouse line. Since each of the tissue defects represents a potentially invaluable clue toward understanding the physiological functions of the mutated gene, their inventory should be as exhaustive as possible. However, listing all the histological abnormalities in a given mouse is useless if they are not correlated with the genetic alteration. Thus, the overall goal of histopathology is to characterize the tissue lesions and ensure they are caused by the genetic alteration. This latter goal can only be achieved if proper control mice are analyzed along with those bearing the genetic alteration, and by having a comprehensive knowledge of the phenotype of the background or control strain to recognize true deviations from “normal.”

Importance of performing systematic analyses At least two scenarios can apply to mutant mouse lines entering a histological screen. The “black box” scenario corresponds to situations where the mutant mice have no overt clinical phenotype, when they die post-natally at variable ages for unknown reasons (Turgeon and Meloche, 2009), and/or when the expression domain of the mutated gene is widespread and a large number of tissues are potentially targeted. In contrast, a mutant mouse line entering a systematic histological screen may display consistent clinical and/or histological defects restricted to a single organ or to a small subset of organs expressing the gene under study. However, altering the functions of a single organ may have broad and unexpected secondary consequences on the whole organism. Thus, almost any mutant mouse is eligible for a systematic screen, even when it carries an organ-specific gene alteration generated through somatic mutagenesis. For instance, mice carrying targeted ablation of retinoid X receptors (RXRs) only in the epidermis develop a systemic syndrome, mimicking that observed in atopic dermatitis patients, including lymph node hyperplasia, splenomegaly, and eosinophilic infiltrates in the liver, lung, and heart (Li et al., 2005).

Post-Mortem Examination and Fixation of Mice

Optimal age for analyzing a mutant mouse line Mice are weaned and sexually mature at 3 and 6 weeks, respectively. Likewise, histogenesis is completed at 3 weeks in most tissues and at 6 weeks in reproductive organs. Although a 6-week-old mouse can be considered a young adult, necropsies and histological analyses are in general postponed until ∼4 months, which is the time required for the completion of in vivo phenotyping tests. Four month-old mice are sexually mature, do not have age-related pathology, and are in an age range for which data on inbred strains are readily available. It should be stressed that a number of mutations affect the propensity to develop age-related diseases. Thus, although the cost of maintaining aging mice is prohibitive, it is sometimes necessary to investigate old (i.e., >12 months) and even senescent (i.e., > 24 months) subjects for late-onset diseases such as degenerative tissue changes and cancers (Harvey et al., 1993; Fan et al., 1998; Huang et al., 2002;

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Kim et al., 2002; Khetchoumian et al., 2007). For instance, Trim24-null (Trim24-/- ) mutants show no histopathological abnormalities at 2 months of age. However, monitoring a large cohort of Trim24-deficient mice (n > 100) for a long time period allows detection of hepatic tumors at necropsy in 80% of male and 69% of female Trim24-null mice between months 9 and 29, as compared to only 4% of age-matched, wild-type control (Trim24+/+ ) mice (Khetchoumian et al., 2007). Conversely, if the mutants die before 4 months of age, it will be necessary to determine the window of lethality before selecting an age group for morphological evaluation. It may also be necessary to study the mutant line at several different ages to accurately follow the progression of the phenotypic changes, and thus understand the biological functions of the mutated gene.

Gender The question regarding the use of males or females in histological investigations is not trivial, as male physiology is not subject to the cyclical changes that, in females, can present another source of phenotypic variability. Upon initial consideration, one might imagine systematically analyzing males and restricting the analysis of female organs to those exhibiting sexual dimorphism (reproductive organs, adrenal glands, hypophysis, kidneys, and salivary glands). However, in mutant lines, gender-specific tissue alterations have been reported in organs that do not display sexually dimorphic histological features (Costet et al., 1998; Liu et al., 2000; Khetchoumian et al., 2007). Therefore, in a firstline histological screen, an equal number of males and females should be systematically analyzed. Control mice and the importance of comparative histopathology To interpret histological data, it is important to have a comprehensive knowledge of the phenotypic peculiarities and common lesions present in the inbred strains used to generate the experimental mouse line. The same holds true with respect to the environment of the experimental mouse line (pathogen status of the animal facility, chlorine or antibiotics in drinking water, and so on). This information must be taken into account to interpret possible discrepancies in data from different laboratories. Common pathologies of inbred strains are mentioned in Brayton et al. (2001) and Naf et al. (2002). It is mandatory, when designing a histological screen, to compare mutant mice to control mice that share the closest possible genetic background and that are bred under identical conditions. Size of cohort and the first-line phenotyping strategy The size of a mouse cohort should be sufficient to allow meaningful scientific interpretation of the data. Scientific statements are generally considered acceptable if they are characterized by an uncertainty of <5% (significance, p < 0.05). In this context, knowledge of the pathologies commonly encountered in the inbred strains composing the genetic background of the experimental mice is relevant in the consideration of group size. On a practical level, 24 mice per age group (six mutants of each gender and their controls) appears to be a reasonable number. However, as it takes ∼4 hr to collect, process, and stain the tissues of a single mouse, issues of time and practicality must be taken into account. The benefit of a systematic histological analysis of ∼40 different organs in 24 mice is uncertain for at least two reasons. First, it cannot identify morphological defects associated with subcellular components that are below the resolution level of light microscopes, as well as defects that are not detectable by standard hematoxylin and eosin (H&E) staining. It is noteworthy that there are relatively few features that can be unequivocally identified and quantified from H&E-stained histological sections, i.e., cells/extracellular matrices, cavities/secretions, nuclei/cytoplasm, and cellular contours. Second, a standard histological analysis will not ensure detection of all morphological defects that can be revealed by the routine H&E stain, notably, defects that are heterogeneously distributed in an organ and/or occur at a low frequency in the mutant mouse line. These considerations highlight the need for a phenotyping strategy. Current Protocols in Mouse Biology

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several age groups

subgroup A • 3 mutant males • 3 mutant females • 1 control male • 1 control female

1

• young, ~8–12 weeks old • old, ⬎12 months old • old, ⬎24 months old • n ⫽ 6 per group

subgroup B 1

1 systematic macroscopic evaluation at necropsy

1 15 days 2

preliminary histopathological report

systematic histology 1 paraffin tissue bank

• 3 mutant males • 3 mutant females • 5 control males • 5 control females

2

systematic macroscopic evaluation at necropsy targeted histology

3 targeted histology

1 transient collection of mice cadavers

3 final histopathological report

Figure 1 Proposed flow scheme for standardized, systematic post-mortem analysis of a cohort of 24 mice. Step 1: Each age group is subdivided into two subgroups on the day of necropsy. Mice from subgroup A are subjected to systematic macroscopic and microscopic analyses. What remains of the paraffin blocks after sectioning is archived in a paraffin tissue bank. Mice from subgroup B are subjected to systematic necropsy and to a targeted histological evaluation. At this point, the subgroup B organs destined for paraffin embedding are the ones carrying a macroscopically visible defect in at least one of the mutant or treated mice. The remaining organs and the carcasses of the mice belonging to subgroup B are kept in buffered formalin until the completion of the histological screen (transient collection of mouse cadavers). Step 2:. A preliminary histopathological report will be available in 2 weeks after necropsy. Step 3: Depending on the histological findings, a second round of targeted histological analysis of subgroup B mice may be undertaken with the objective of increasing the statistical significance of the preliminary observations. Organs of subgroup B analyzed by histology at this stage are the ones in which histological defects have been detected in the subgroup A mutant or treated mice. To this purpose, these organs are collected from the bank of mouse cadavers, processed, embedded, and then stained with hematoxylin and eosin.

Post-Mortem Examination and Fixation of Mice

In the flow scheme shown in Figure 1, each age group is subdivided into two subgroups on the day of sacrifice. Subgroup A consists of three mutants of each gender, one control male, and one control female (ideally wild-type littermates). Mice belonging to this subgroup are subjected to a systematic necropsy (i.e., macroscopic analysis and weighing of selected organs) and to a systematic standardized microscopic analysis of ∼40 organs (Table 1 and Fig. 2). Following tissue dissection, all organs, with the exception of testes, are then fixed for a standard period of 24 hr in 10% neutral buffered formalin prior to paraffin embedding. The testes are fixed in Bouin’s fixative, which not only yields a more life-like preservation of the testicular parenchyma than buffered formalin, but also allows one to perform a high-contrast periodic acid Schiff (PAS) stain, which is required to analyze the seminiferous epithelium cycle. Bones (long bones, vertebrae, skull) are decalcified. For each organ, one slide is prepared carrying three consecutive 5-μm-thick histological sections, and is stained with H&E. Therefore, only a small portion of the organ is actually evaluated systematically. What remains of the paraffin block is archived to generate a paraffin tissue bank that can be kept at room temperature for years. These paraffin blocks can be used later for first-line extended assays, including serial sections through the organ, special histological stains, immunohistochemical stains, and cell proliferation and cell death assays. With respect to immunohistochemistry, it is expected that the duration of formalin fixation (24 hr) is short enough to preserve a large number of epitopes.

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Table 1 Organs Systematically Analyzed in a First-Line Histopathological Screen

Cassette

Organs

1 (male)

Preputial glands, salivary glands, pancreas

1 (female)

Mammary glands, salivary glands, pancreas

2

Stomach

3

Duodenum, distal ileum, proximal colon

4

Liver

5

Spleen, kidney

6A (male)

Right testis and epididymis (fixed in formalin)

6B (male)

Left testis and epididymis (fixed in Bouin’s fixative)

7 (male)

Prostates, seminal vesicles, urinary bladder

6 (female)

Ovaries, oviducts

7 (female)

Vagina, uterus body, uterine horns, urinary bladder

8

Adrenal gland, mesenteric lymph nodes, thoracic aorta

9

Trachea, thyroid glands, esophagus, thymus

10

Heart, entire lung

11

Leg muscle, tongue, BAT, WAT

12

Dorsal, tail, footpad, and snout skin

13

Eye, Harderian gland

14

Knee joint

15

Brain

16

Hypophysis

necropsy and fixation: fixed tissue

Figure 2

tissue processing

embedding

paraffin sectioning

data analyses and interpretation: diagnosis, semi-quantification, and imaging

staining (automatic strainer)

Steps following necropsy and fixation in histopathological analyses of mouse tissues.

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Subgroup B comprises the six remaining mutant mice and ten control mice. Mice belonging to this subgroup are subjected to a systematic macroscopic evaluation and a targeted histological evaluation. At this point, the organs that carry a macroscopically visible defect in at least one of the mutants in subgroup B are embedded in paraffin. For instance, if a liver nodule is observed in one mutant mouse, the livers of all twelve mutant mice and all twelve controls are immediately embedded in paraffin for subsequent histological analysis. The remaining, apparently “healthy” organs and carcasses of the mice belonging to subgroup B are kept in 10% neutral buffered formalin until the completion of the first-line histological screen. Following the ∼2-week period required to complete the histological analysis of the subgroup A mice, a second round of targeted histological analysis of subgroup B organs is undertaken. In the second round, the organs of subgroup B that are analyzed by histology are the ones in which histological defects have been detected in the subgroup A mutants (Fig. 1). The first-line screen can lead to four possible conclusions: 1. Tissues from mutant mice, fixed in formalin and stained with H&E, appear normal. This result will end the analysis. 2. Mutant mice display morphological defects that are not correlated with the mutation, as they occur at the same penetrance and with the same severity as in control mice. These defects should be considered spontaneous “background effects” unrelated to the mutation. This will also end the analysis. 3. Mutant mice display morphological defects that are also present among control mice, albeit at a lower frequency. In this case, the first-line screen must be repeated on a larger sample of mice to obtain statistically significant data. 4. Mutant mice display morphological defects that are correlated with the mutation. In this case, the organ defects should be analyzed further using first-line extended and secondary histological assays.

First-line extended assays Organs entering these assays display known abnormalities related to the mutation. The objectives now are to define the penetrance and the variations in severity of each defect in the mouse line, to refine the accuracy of the diagnosis, and to gain insights into the molecular defects associated with the abnormal morphology. First-line extended assays are performed on tissues already collected and preserved during the first-line screening, and can involve any of the following: 1. Serial histological sections, collected consecutively or at regular intervals through the paraffin block. The goal is to increase the chances of detecting a lesion through analysis of organ volumes. 2. Histomorphometry aimed, for example, at determining the relative number of Langerhans cells in the pancreas. These analyses can often be performed with the help of user-friendly, free software (e.g., ImageJ at http://rsb.info.nih.gov/ij/download.html). 3. Cell proliferation and cell death assays. 4. Immunohistochemical assays. 5. Tissue- and pathology-specific stains. Post-Mortem Examination and Fixation of Mice

The limits of the methods that can be applied in these assays are defined by the state of routinely processed organs. For instance, organs in paraffin are unsuitable for detecting

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certain cellular constituents, such as neutral lipids, that are removed by the series of alcohol extractions and paraffin solvents. Paraffin-embedded organs are also unsuitable for electron microscopic studies, which require specific fixation and embedding procedures. Additionally, immunohistochemical analyses on these organs are not always successful, since antigens are often lost during formalin fixation, dehydration, and/or heating of tissues (Werner et al., 2000).

Secondary assays Similar to the first-line extended assays, secondary assays are targeted to specific organs or organ systems. However, they require the use of special protocols for tissue collection (e.g., perfusion fixation) or special histological stains that cannot be adapted to formalin-fixed and paraffin-embedded tissues, and therefore require the generation of a new cohort of mutant and control mice. Secondary assays may include electron microscopic analyses as well as histochemical, immunohistochemical, and in situ hybridization assays. NOTE: All protocols using live animals must first be reviewed and approved by an Institutional Animal Care and Use Committee (IACUC) and must follow officially approved procedures for the care and use of laboratory animals. Regulations concerning the persons allowed to sacrifice a mouse vary between countries. Local authorities should be consulted for the proper procedure.

NECROPSY FOR HISTOPATHOLOGICAL ANALYSIS Mouse identification and sample labeling Confirm the identity of the mouse (Fig. 3), and make sure this matches the corresponding necropsy request (Fig. 4). Use a unique mouse identification number for labeling samples and reporting findings (Fig. 3). Required information for sample labeling at necropsy (i.e., on histological cassettes and slides) includes: cassette number (see necropsy template in Fig. 4 and Table 1), mouse number, sex, treatment group, and project number or necropsy date.

BASIC PROTOCOL 1

External examination The purpose of external examination is to easily record information regarding the state of health for the mouse. This information concerns the general aspects of the mouse (e.g., obese, thin, or malformed), the state of superficial tissues and organs (e.g., skin, eyes), and the natural orifices (which may be informative about some inner-body pathology; e.g., in the mouth, pallor of the oral mucosa is indicative of anemia). Body and organ weights Total body weight as well as the weights of heart, liver, spleen, genital fat pads, and kidney, are systematically determined. Tissue dissection Tissue dissections are not presented in complete detail, and it is assumed that the investigator or technician performing these dissections has had proper training and experience in mouse dissection before attempting these analyses. Although some directions are provided for dissection, the focus of this protocol is the strategy for dissection and proper grouping of tissues. To preserve organs, use attached tissues such as ligaments and mesentery to handle the organs. Trim off adherent tissues before tissue/organs are weighed. Do not attempt to isolate small gross lesions. Additional details for trimming tissues and defining planes of section can be found in the Support Protocol.

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600

3

4

200

6

400

5

7

2

8

1

9

100

10

90

20 30

50 40

60

70

80

Figure 3 Identification of mice by tattooing digits and ears. This figure proposes a numbering code for the permanent identification of 2-week-old to adult mice using forelimb digits for units, hindlimb digits for tens, and ears for hundreds. Units are tattooed by ink injection of forelimb digits, starting the count with 1 on the outermost right (anatomically right digit V) and progressing towards the outermost left (left digit V). Number 5 is obtained by tattooing both digits III and IV. Note that forelimb digits I are much reduced in size and therefore impossible to tattoo. Tens are tattooed on hindlimb digits, starting the count with 10 on the outermost right (anatomically right digit V) and progressing towards the outermost left (anatomically left digit V). As to hundreds, numbers 200 and 400 are indicated by tattooing dots on the right and left ears, respectively. Dots on both right and left ears indicate 600 (i.e., 200 plus 400). Units, tens, and hundreds are then added.

Fixation Once death has occurred, tissues undergo a process of self-digestion (autolysis) that is accelerated by heat and by post-mortem colonization of bacterial flora from the gut (putrefaction). The first goal of fixation is to halt post-mortem tissue alterations, preserving cell structures in forms that resemble the living state. Fixation also hardens the tissue, which helps in subsequent handling. The most commonly used fixative is buffered formalin (Basic Protocol 2), although testes must be fixed in Bouin’s solution (Alternate Protocol). Immediately place fresh tissues in plastic embedding cassettes and immerse in fixative. Most importantly, use a volume of fixative that is 10 to 20 times the volume of the tissue. For most tissues, fixation for 24 hr in 10% neutral buffered formalin is sufficient before trimming and proceeding to paraffin embedding. For head and long bones, fix for 48 hr in 10% neutral buffered formalin; for testis, fix in Bouin’s fixative for 24 hr. For further details on fixation, see Basic Protocol 2. Be aware that special techniques such as histochemistry, including detection of enzymes, immunohistochemistry, and in situ hybridization, require special fixatives and short fixation periods for optimal results (Lillie, 1965; Wilkinson, 1999; Morel and Cavalier. 2000; Polak and Van Noorden, 2003; Renshaw, 2007).

Post-Mortem Examination and Fixation of Mice

Order of dissections The order in which organs are dissected in the systematic necropsy procedure is described below, and the grouping of organs in embedding cassettes is summarized in Table 1. This protocol is indicated for a mouse that has no overt clinical phenotype. However, any organ that, based on gene expression patterns or clinical findings, represents a likely target of the mutation should be protected from autolytic damage as a priority. For example, if the

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Necropsy template

Necropsy date: Operator: Project number:

Mouse n°: sex: Mouse line & background strain: Date of birth: Treatment group:

Death mode:

Found dead

Organ weight Organ

6B 7 7 7 6 6 7 7 7 7 8 8 8 9 9 9 9 10 10 11 11 11 11 12 12 12 13 13 14 15 16

F

Death during anesthesia

: Weight (mg)

Body weight: Heart: Liver: Spleen: Kidneys (left & right): Paragenital WAT:

Cassette Number 1 1 1 1 1 1 2 3 3 3 4 5 5 6A

Euthanasia Clinical symptoms before/at death

M

Organ

Tail biopsy at – 20°C (for genotyping)

Keep cadavers in formalin

Macroscopy/observations

(male) Preputial glands: Salivary glands: Pancreas: (female) Mammary glands: Salivary glands: Pancreas: Stomach: Duodenum: Distal ileum: Proximal colon: Liver: Spleen: Kidney: (male) Right testis and epididymis (fixed in formalin): (male) Left testis and epididymis (fixed in Bouin): (male) Prostates: Seminal vesicles: Urinary bladder: (female) Ovaries: Oviducts: (female) Vagina: Uterus body: Uterine horns: Urinary bladder: Adrenal gland: Mesenteric lymph nodes: Thoracic aorta: Trachea: Thyroid glands: Esophagus: Thymus: Heart: Entire lung: Leg muscle: Tongue: BAT: WAT: Dorsal tail skin: Footpad skin: Snout skin: Eye: Harderian gland: Knee joint: Brain: Hypophysis:

Samples for histology

Picture

yes yes yes yes yes yes yes yes yes yes yes yes yes yes

no no no no no no no no no no no no no no

yes yes yes yes yes yes yes yes yes yes yes yes yes yes

no no no no no no no no no no no no no no

yes

no

yes

no

yes yes yes yes yes yes yes yes yes yes yes yes yes yes yes yes yes yes yes yes y es yes yes yes yes yes yes yes yes yes

no no no no no no no no no no no no no no no no no no no no no no no no no no no no no no

yes yes yes yes yes yes yes yes yes yes yes yes yes yes yes yes yes yes yes yes yes yes yes yes yes yes yes yes yes yes

no no no no no no no no no no no no no no no no no no no no no no no no no no no no no no

Comments

Figure 4

Template for necropsy report.

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hypophysis is a likely target, the necropsy should start with the head, not the subcutaneous glands as proposed below. The entire necropsy procedure should be completed in ∼45 min.

Reporting necropsy findings Use correct anatomical and medical terminology to describe the location, color, size, shape, consistency and distribution and number or percent of involvement of a specific organ of any tissue abnormalities, including absence of organs (see mouse anatomy dictionary at http://www.informatics.jax.org/searches/anatdict form.shtml). Keep descriptions accurate and avoid overinterpretation. A general term such as a mass is preferable to either tumor or abscess, because these more refined diagnoses require microscopic confirmation. Diagrams should be used to provide extra information or clarity, and photographs should be taken of all grossly abnormal organs and external lesions. A ruler should also be included in the photograph for scale. CAUTION: General laboratory safety procedures should be followed, including not eating, chewing gum, drinking, or applying cosmetics in the work area. Laboratory coats must be worn at all times in the work area. Formalin is a carcinogen, and an eye, skin, and respiratory irritant. Avoid contact and inhalation. Work under a fume hood and wear disposable gloves. Collect used fixatives containing formalin in an appropriate waste container for disposal. For further information, consult a local center for occupational health and safety. Clean the instruments and workstation immediately after necropsy work has been completed.

Materials Mice (see Strategic Planning) CO2 source 70% ethanol (see recipe) 10% (v/v) neutral buffered formalin (equivalent to 4% [w/v] formaldehyde; see recipe) Bouin’s fixative solution (e.g., VWR International) for testes 1× phosphate-buffered saline (PBS; see recipe)

Post-Mortem Examination and Fixation of Mice

Box with transparent walls and/or lids Ruler Electric shaver Cork dissecting board and dissecting pins Dissecting microscope equipped with a digital camera for documenting lesions and abnormalities Balance (e.g., Fisher Scientific) Large containers with lids to hold fixing fluids (e.g., Fisher Scientific) Dissection instruments, e.g.: Dissecting forceps including a pair of rat-toothed forceps Fine pointed forceps for delicate dissections Scissors: a pair should be dedicated to cutting bones Scalpels (size 3 with no. 15 blade) Razor blades Spatula to lift the brain and remove the hypophysis Plastic embedding cassettes (e.g., Fisher Scientific) 5-ml syringe and 25-G needles 50-ml conical polypropylene centrifuge tubes with caps (e.g., BD Falcon) Biopsy capsules (Leica, code OEL-039430014) 90-mm Petri dishes Mouse brain and matrix (e.g., stainless-steel for long-term use or acrylic for sporadic use; Harvard Apparatus)

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Euthanize mouse by carbon dioxide inhalation 1. Deliver CO2 into a box with transparent walls and/or lid (euthanasia chamber) gradually so that the mouse will be exposed to a slowly rising concentration of gas (euthanasia is appropriate if CO2 concentration is greater than 70%). Place the mouse into the euthanasia chamber so that it can be observed during exposure to the carbon dioxide. Before proceeding, ensure that death has occurred by testing for the absence of vital signs, including respiration and corneal reflex. The slowly rising concentration of gas ensures that the mouse will lose consciousness because of the effects of the CO2 on the central nervous system, rather than suffocating because of immediate exposure to 100% CO2 . IMPORTANT NOTE: Welfare concerns regarding the use of CO2 for euthanasia have been raised. Among the issues are the optimal concentration of CO2 in the chamber for inducing rapid unconsciousness with a minimum of respiratory distress, avoiding reflex reactions to anoxia, optimal procedure for placing the animal(s) in the euthanasia chamber, and the irritant properties of CO2 when inhaled. CO2 euthanasia should only be performed according to an IACUC-approved SOP (standard operating procedure).

Perform external examination 2. Using a ruler, measure the length of the mouse from the snout to the tail base. Record findings on each of the following: a. State of nutrition and dysmorphologies Thin mouse Obese mouse Dysmorphologies: e.g., craniofacial abnormality, polydactyly, shortened limbs or curly tail b. Skin and subcutaneous tissue Traumatic wound Ulcers Infectious lesions Skin masses Mammary gland masses Edema: swollen, smooth and glossy skin Condition of fur: rough, dry, and hirsute in a serious chronic disease, or depilated areas c. Natural orifices and eyes i. Mouth Oral mucosa: pale mucosa in case of anemia; petechiae (hemorrhages of the submucosa) in infectious diseases; erosions of the mucosa, ulcers, or vesicles Teeth: loss, erosion, or fracture of teeth (damaged or missing teeth may prevent adequate animal feeding and can be a cause of death). Excessive incisor growth can also prevent mice from feeding normally and can cause malnutrition. ii. Nasal openings Discharges Hemorrhages (epistaxis) iii. Eyes Narrowed eyelid opening (blepharophymosis) Corneal and conjunctival exudates and ulcerations. iv. Ear pinnae and external ear canals Discharges v. Anal opening Smeared by feces in animals affected by diarrhea Rectal prolapse Current Protocols in Mouse Biology

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B

A

Figure 5

(A) Mouse with shaved back. (B) Mouse pinned ventral-side-up to cork board.

vi. External genitalia Discharges Prepare mouse 3. Collect a segment of tail (2 to 3 mm in the terminal piece) in a 1.5-ml microcentrifuge tube and store at –20◦ C for regenotyping. 4. Wet the fur with 70% ethanol and shave a portion of the back skin with an electric shaver (Fig. 5A). Shaving with a razor blade may damage superficial skin layers.

5. Lay the mouse on its back and pin the forelimbs and hindlimbs onto a cork board (Fig. 5B). Access to the back will be needed later, but it is easier to shave the back before starting the dissections.

6. Wet the fur with 70% ethanol to minimize contamination with hairs and other potential allergens. 7. Perform a midline incision of the ventral skin, with scissors, from the pubis to the chin. Extend the incision laterally towards inguinal areas to form an upside-down Y, taking care to accurately separate the skin from the underlying musculature (Fig. 6A). Be prepared to describe the abnormalities found at necropsy by the following criteria:

Post-Mortem Examination and Fixation of Mice

Location Number and distribution Color Size Shape Consistency and texture.

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B

A

Figure 6

C

(A) Midline incision with skin reflected. Dissection of (B) salivary glands and (C) male preputial gland.

For example: “The liver displays four white, soft, nodules ranging from 0.2 to 0.5 cm in diameter that are restricted to the left lobe.”

Dissect subcutaneous glands 8. Dissect the three salivary glands (Fig. 6B) and place them into cassette no. 1 (Table 1). 9a. For male: Separate one lobe of the preputial gland (Fig. 6C) and place it into cassette no. 1 with the salivary glands. 9b. For female: Detach the inguinal mammary gland from the overlying skin by scraping gently with scissors (Fig. 7) and place it into cassette no. 1 with the salivary glands.

Dissect abdomen 10. Make a midline incision through the abdominal wall muscles. Dissect the pancreas away from its insertion on the spleen and duodenum (Fig. 8), place it into cassette no. 1, and transfer the cassette to 10% neutral buffered formalin. 11. Working in a fume hood, take up 5 ml of the 10% neutral buffered formalin fixative into a 5-ml syringe. Using a 25-G needle, slowly inject 3 ml fixative into two or three areas along the intestine, into the lumen (Fig. 9), taking care not to burst it. Examine different segments of the intestine and inject additional fixative in any regions that have not been reached by the fixative. Puncture the stomach and inject fixative. Remove the stomach and the intestines from the abdomen and place them into a 50-ml conical polypropylene centrifuge tube containing 40 ml of 10% neutral buffered formalin. Proceed with other tissue dissections while allowing stomach and intestines to fix for 24 hr. A volume of 5 ml of fixative is usually enough to inject the entire digestive tract. Alternatively, proceed to the Swiss roll technique (Moolenbeek and Ruitenberg, 1981). The Swiss roll technique allows scanning of large parts of the intestine. In brief, selected fragments (e.g., ∼5 cm of duodeno-jejunum, ileum, and colon) are split open longitudinally with ball-tip scissors (to prevent mucosa injuries). The feces that were not washed out by the fixative are removed. A toothpick is placed on the side of the slit and the intestinal segments are rolled up such that the mucosa remains on the inside in this “Swiss roll” preparation. The preparation is then carefully detached from the toothpick and placed in a cassette, into fixative.

12. Dissect the liver. Cut the esophagus, ligaments, and omentum that connect the liver to the stomach, diaphragm, and the right kidney (Fig. 10). Weigh the liver. Collect the median lobe and half of the left lateral lobe, place them into cassette no. 4, and transfer into 10% neutral buffered formalin. Place the remaining liver into the 50-ml tube containing formalin, for storage (see step 11).

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A

B

C

Figure 7 Dissection of mammary gland. (A) Insertion of scissors between the skin and the mammary gland, (B) spreading scissors to dissect the mammary gland from the skin, and (C) sectioning remaining adherences.

13. Remove the spleen, weigh it, and place it into cassette no. 5.

Post-Mortem Examination and Fixation of Mice

14. Make a transverse cut through the hilum of the kidney (Fig. 11A) and separate the organ from the dorsal abdominal wall, while leaving in place the dorsal half of its capsule and the adrenal gland. Weigh the kidney. With a razor blade, make a longitudinal cut through the hilum (Fig. 11B) and place the two moieties into cassette no. 5 with the spleen. Close the lid and transfer to 10% neutral buffered formalin.

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liver stomach spleen

pancreas

Figure 8

Dissection of pancreas.

Figure 9

Fixation of intestine.

Dissect the urogenital organs For male 15a. Elevate the right testis from the scrotal sac by gently pulling on the paragenital fat (Fig. 12A). Dissect and weigh the paragenital fat and place it into the same 50-ml plastic tube containing formalin. The paragenital fat will be placed in cassette no. 11, together with brown adipose tissue, skeletal muscle, and tongue. These organs are the gold standard for metabolic/obesity studies. They are also grouped for practical reasons to minimize the number of cassettes, and also because they are of similar size.

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B

A

Figure 10

Dissection of liver.

B

A

Figure 11

Dissection of kidneys.

16a. Cut the thin ligament connecting the tail of the epididymis to the body wall (Fig. 12A). Place the right testis and epididymis into cassette no. 6A and transfer to 10% neutral buffered formalin. Place the left testis and epididymis into cassette no. 6B and transfer to a container with ∼20 volumes of Bouin’s fixative solution. Discard the remaining portion of the vas deferens. 17a. Using forceps, grasp the highest point of the urinary bladder, lift it up, and cut the ligament connecting the urinary bladder to the ventral body wall (Fig. 12B). 18a. Recline both seminal vesicles towards the tail to uncover the ligaments connecting the prostate and dorsal body wall. Cut these ligaments (Fig. 12C), and then make a transverse cut through the pelvic portion of the urethra (Fig.12D). 19a. Place the group composed of the bladder, seminal vesicle, and prostate into the 50ml tube containing 10% neutral buffered formalin. Allow to fix for 24 hr. Proceed to step 20.

For female 15b. Dissect and weigh the paragenital fat (Fig. 13A), and place it into the 50-ml plastic tube containing 10% neutral buffered formalin. Post-Mortem Examination and Fixation of Mice

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The paragenital fat will be placed into cassette no. 11, together with brown adipose tissue, skeletal muscle, and tongue. These organs are the gold standard for metabolic/obesity studies. They are also grouped for practical reasons to minimize the number of cassettes, and also because they are of similar size. Current Protocols in Mouse Biology

A

B bladder epididymis

paragenital fat

ligament

testis

C

ligaments

D

bladder cranial prostate seminal vesicle

Figure 12

ventral prostate urethra

cranial prostate

(A) Dissection of testis, (B) urinary bladder, (C) seminal vesicle, and (D) prostate.

16b. Cut the ligaments that connect the ovaries to the dorsal abdominal wall (Fig. 13B). 17b. Using forceps, grasp the highest point of the urinary bladder (Fig. 13C). Pull the bladder upwards, cut the ligaments that connect the uterus body and vagina to the dorsal abdominal wall (Fig. 13D), and make a transverse cut through the proximal vagina. 18b. Remove the genital tract and ovaries en bloc. Separate the urinary bladder. Make a transverse cut at the junction between the uterus and oviduct, and a second one at the junction between the uterus body and uterine horns (Fig. 14). 19b. Place ovaries with attached oviducts into cassette no. 6, and place the vagina, the uterus body, both uterine horns, and the urinary bladder into cassette no. 7. Close the lids and transfer to 10% neutral buffered formalin. Proceed to step 20. 20. Remove the adrenal gland (Fig. 15A) and the mesenteric lymph nodes (Fig. 15B) and place them into a biopsy capsule placed in a 90-mm Petri dish with 1× PBS. Later in the dissection procedure, this capsule will also contain the thoracic aorta.

Dissect neck and thorax 21. Cut the diaphragm along its insertions on the rib cage, cut the pleura connecting the lung to the diaphragm, and then section the distal portion of the thoracic esophagus. Remove the aorta (Fig. 16) by making a transverse cut through its distal thoracic part and dissecting it away from the vertebral column. Take a 1-cm segment. 22. Place the aorta in the biopsy capsule containing the adrenals and lymph node. Close the capsule, place it into cassette no. 8, and transfer to 10% neutral buffered formalin. Current Protocols in Mouse Biology

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B

A

ovary uterine horns

C

paragenital fat

oviduct

D bladder uterus

vagina

Figure 13 Dissection of (A) paragenital fat in the female, (B) ovaries and oviduct, and (C,D) bladder, vagina, and uterus (C,D)

23. Remove and discard the neck muscles covering the trachea and the thyroid (Fig. 17A-C). 24. Open the rib cage by cutting through the dorsal third of the right ribs, then recline the rib cage and sternum towards the left, and pin it onto the cork board (Fig. 18A). 25. Slowly inject 2 ml of 10% neutral buffered formalin into the lungs by introducing a 25-G needle between the tracheal rings and compressing the trachea with forceps for 30 sec to maintain the fixative in the lungs (Fig.18B). 26. Remove the superior (cervical) part of the trachea, with the thyroid glands and esophagus attached, by cutting transversely at the level of the inferior pole of the thyroid glands (Fig.18C). Place the group into cassette no. 9 together with the thymus. Close the cassette and transfer to 10% neutral buffered formalin. 27. Remove the heart, weigh it, and place it into cassette no. 10.

Post-Mortem Examination and Fixation of Mice

28. Dissect the remaining cervical trachea and esophagus from their adherences to the cervical spine. Extend the dissection into the thorax and remove the lungs en bloc. Put the entire block into cassette no. 10 with the heart, and place the cassette into 10% neutral buffered formalin.

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ovary and oviduct

uterine horn

uterus body

bladder

Figure 14

Dissected ovaries, oviduct, bladder, vagina, and uterus.

B

A

adrenal gland adipose tissue

Figure 15

Dissection of (A) adrenal glands and (B) mesenteric lymph nodes. Post-Mortem Examination and Fixation of Mice

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aorta

Figure 16

Dissection of aorta.

Dissect muscles, skin, fat, and bones 29. Remove the skin from the hindlimb. Insert the scissors under the Achilles’ tendon and spread the blades (Fig. 19A) to separate the gastrocnemius and soleus muscles from the deep muscular layer. Cut the muscles at the level of the knee joint (Fig. 19B) and place them into cassette no. 11. 30. Spread the jaws with the tip of the scissors, cut out the tongue, and place it in cassette no. 11. 31. Turn the mouse over on its ventral side. Make a longitudinal incision in the skin between the two scapulae (Fig. 20A). Spread out the edges of the incision, remove the interscapular brown fat (Fig. 20B,C), and place it into cassette no. 11. 32. Take a piece of white adipose tissue from the tissue stored in the 50-ml plastic tube (step 15a or 15b) and place it into cassette no. 11. Close the lid and immerse the cassette into 10% neutral buffered formalin. 33. Dissect and discard the hindlimb muscles to expose the knee joint. Separate the knee joint by sectioning the bones above and below the knee (Fig. 21). Place it into cassette no. 14 in a fresh container with 10% neutral buffered formalin (different from the one used for the previous organs).

Dissect head 34. Make a midline incision on the dorsal side of the head skin from the neck to the snout. Cut off the head. 35. Optional: Place entire head in 10% neutral buffered formalin fixative for 48 hr. Post-Mortem Examination and Fixation of Mice

The following operations can be performed on fresh tissue or after 48 hr fixation of the whole head.

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A

B

C

Figure 17

Removal of neck muscles (A,B) and view of exposed thyroid and trachea (C).

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A

B

thyroid glands

C

Figure 18 (A,B) Injection of fixative through trachea for fixation of lungs. (C) Lateral view of trachea with esophagus and thyroid.

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B

A Achille’s tendon gastrocnemius muscle

Figure 19

Dissection of gastrocnemius and soleus muscles.

36. Insert the scissors into the foramen magnum and make a midline incision through the occipital bone (Fig. 22). 37. Place the head on the cork board and firmly insert the tips of the scissors at the junction of the frontal and nasal bones, on the midline (Fig. 23A,B). Open the scissors. This action will break the dome of the skull along the medial suture (Fig. 23C).

38. With a pair of forceps, detach the two halves of the dome and expose the brain (Fig. 23D). 39. Using a spatula, remove the brain, taking care not to damage the hypophysis located underneath (Fig. 24). 40. Place the brain in a Petri dish containing 1 ml of 1× PBS. Place the brain, ventral surface up, in the mouse coronal brain matrix (Fig. 25A). The ventral surface of the brain must be parallel to the top surface of the mold.

41. Use the 4th, 6th, and 10th channels, starting from the anterior part of the mold, to generate the brain slices (Fig. 25B). Insert razor blades in the first and last channels, and then insert a razor blade in the middle channel. Remove the three razor blades at the same time, extracting the brain slices from the mold. Place the brain slices in cassette no. 15 and transfer to 10% neutral buffered formalin. The manipulations performed to remove the brain from the skull have broken the skull base. This renders eye removal easy as, in most cases, the orbit is already open.

42. Cut the membrane at the edges of the orbit all around the eyeball. Manipulate the eye using the optic nerve or the skin. Do not pull on these tissues as there is a risk of detaching the retina. 43. Place the eye and the corresponding Harderian gland in cassette no. 13. Close the lid and transfer to 10% neutral buffered formalin. 44. Place what remains of the skull into the 50-ml plastic tube containing 10% neutral buffered formalin and fix for 24 hr.

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A

B brown adipose tissue

C brown adipose tissue

Figure 20

Dissection of brown adipose tissue (BAT).

If there is an area of particular interest, such as a tumor in the organ, trim the tissue so that the area of interest is near one surface of the specimen, and place that surface down in the cassette.

Post-Mortem Examination and Fixation of Mice

Process carcass 45. Transfer the mouse carcass to a large container with 10% neutral buffered formalin. Allow to fix for 24 hr.

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Figure 21

Dissection of knee joint.

Process tissues after 24-hr fixation 46. Separate the stomach by making a transverse section at its junction with the proximal duodenum, and place the stomach in a Petri dish containing 70% ethanol. Cut with scissors along the lesser and greater curvatures and wash out the contents. Place both halves in cassette no. 2 and place the cassette back into 10% neutral buffered formalin. 47. Take a 5-mm segment of the proximal duodenum, a 5-mm segment of the distal ileum closest to the cecum, and the first 5-mm segment of the proximal colon. Place the three segments in cassette no. 3 and place the cassette back into 10% neutral buffered formalin. 48. Collect two 4-mm of pieces of skin from each of the following locations, place into cassette no. 12, and transfer to 10% neutral buffered formalin: a. Back skin: Remove a rectangular piece of dorsal skin, parallel with the longitudinal axis of the body. b. Tail skin: Cut a 1-cm segment of the proximal tail, incise it longitudinally (Fig. 26A), and detach the skin from the vertebra using a scalpel (Fig. 26B). c. Footpad skin: Remove the skin from one of the hindlimb footpads. d. Snout skin: Remove the vibrissae by cutting them with scissors and take a rectangular piece of snout skin. 49. Remove the sample of male urogenital organs (prostate, seminal vesicles, bladder) from the 50-ml tube of fixative (step 19a), trim (see Support Protocol), place in cassette no. 7, and transfer to 10% neutral buffered formalin.

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Figure 22

Incision through occipital bone for dissection of brain.

A

B

C

D

Figure 23 Removal of brain. (A) Placement for insertion of scissors tips, (B,C) splitting of the skull along medial suture, and (D) removal of skull halves.

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trigeminal nerves optic chiasma hypophysis

optic nerves

Figure 24

Brain reflected towards anterior, showing optic chiasma and hypophysis underneath.

A

B

Figure 25 slices.

(A) Placement of brain in brain matrix and (B) position of razor blades for cutting

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A

B

Figure 26

Dissection of tail skin. (A) Longitudinal cut and (B) removal of spine.

50. Remove the skull from fixative (step 44), remove the hypophysis, place in a biopsy capsule in cassette no. 16, and transfer to 10% neutral buffered formalin. 51. Store the remainder of the carcass, together with the remaining organs, in 10% neutral buffered formalin at room temperature for further use. Good histology results are obtained after 1 month and, according to some authors, even after 1 year of storage in formalin. BASIC PROTOCOL 2

FIXATION FOR ROUTINE HISTOPATHOLOGICAL ANALYSES WITH FORMALIN Formalin (i.e., formaldehyde) stabilizes cell structures by introducing covalent crosslinks, primarily between proteins and to a lesser extent between nucleic acids and carbohydrates (Fox et al., 1985). Formalin renders small proteins insoluble by linking them to structural proteins. It has no direct effects on lipids, although it can preserve some of these components by trapping them in the cross-linked protein network. It has damaging effects on DNA and RNA structure (Srinivasan et al., 2002). CAUTION: Formalin is a carcinogen and an eye, skin, and respiratory irritant.

Materials

Post-Mortem Examination and Fixation of Mice

Mice (see Strategic Planning) 10% (v/v) neutral buffered formalin (equivalent to 4% [w/v] formaldehyde; see recipe) 70% ethanol (see recipe) Plastic embedding cassettes (e.g., Fisher Scientific) Large (≥1 liter) containers with lids (e.g., Fisher Scientific)

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1. Excise desired tissue from euthanized mouse, promptly place into a labeled plastic embedding cassette, and close the lid. Do not overfill the cassette (the lid should close without resistance). Overcrowding can impair subsequent processing. Furthermore, it is not possible to position too many tissues in the paraffin block and still obtain good sectioning.

2. Immediately place the cassette in 10 to 20 volumes (relative to tissue volume) of 10% neutral buffered formalin in a large container and close the lid. Tissue should be placed in fixative preferably within 30 min, to avoid autolytic alterations.

3. Fix for 20 to 24 hr at room temperature. 4. If tissues cannot be processed immediately, rinse 10 min in 70% ethanol and store in 70% ethanol at room temperature until use (up to 1 month). Do not reuse the buffered formalin.

FIXATION FOR ROUTINE HISTOPATHOLOGICAL ANALYSES WITH BOUIN’S SOLUTION

ALTERNATE PROTOCOL

Bouin’s solution is used routinely for mouse embryos, fetuses, and placentas, as well as adult testes because it preserves nuclear details and chromosomes better than formalin and causes less tissue distortion (Fig. 27). Bouin’s solution is a mixture of formalin, picric acid, and acetic acid. Whereas formalin introduces covalent cross-links between proteins, picric acid forms picrates with proteins, which are insoluble in alcohols. Compared to buffered formalin, Bouin’s solution improves trichrome stains because of its mordant effect on collagen fibers (Luna, 1992). However, it is unsuitable for immunohistochemistry, unless the antigen is very robust. CAUTION: Bouin’s fixative contains picric acid. Dry picric acid is a shock-sensitive explosive capable of releasing energy on a level similar to dynamite. Care must be taken to maintain the fixative in its liquid state. CAUTION: Formalin is a carcinogen and an eye, skin, and respiratory irritant.

Materials Mice (see Strategic Planning) Bouin’s fixative solution (e.g., VWR International LLC ) 70% ethanol (see recipe) Plastic embedding cassettes (e.g., Fisher Scientific) Large containers with lids (e.g., Fisher Scientific) Rocking platform 1. Excise desired tissue from euthanized mouse, promptly place into a labeled plastic embedding cassette, and close the lid. Do not overfill the cassette (the lid should close without resistance). 2. Immediately place the cassettes in 10 to 20 volumes (relative to tissue volume) of Bouin’s fixative solution in a large container and close the lid. Work as quickly as possible to avoid autolytic changes. 3. Fix for 24 to 48 hr at room temperature. 4. Wash in 70% ethanol for 1 hr at room temperature. 5. Wash again overnight with 70% ethanol under mild agitation on a rocking platform to remove as much picric acid as possible before embedding in paraffin. Proceed to embedding within 1 month.

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A

B

C

D

Figure 27 Applications and limits of routine buffered formalin fixation. In general, a solution of 10% neutral buffered formalin preserves tissues appropriately and provides equal or higher quality for tissue stained with hematoxylin-eosin than formalin-substitute fixatives, the testis presenting a notable exception. (A) Secretion granules in Paneth cells (arrows), located within the intestinal crypts, are clearly identifiable after formalin fixation but (B) are extracted and thus appear as empty vacuoles after immersion in an acetic acid–alcohol proprietary fixative. (C) The seminiferous epithelium of the testis (here stained with hematoxylin and eosin) appears much more “life-like” after Bouin’s fixation than (D) after formalin fixation. Scale bar in D: 20 μm (A-D).

SUPPORT PROTOCOL

TRIMMING ORGANS AND DEFINING PLANES OF SECTION Skin and subcutis, musculoskeletal system Mammary gland No specific orientation is required. Embed and section to obtain the largest cut surface. Knee joint Place the bones along axes parallel to the bottom of the embedding mold to generate longitudinal sections. Muscle Orient the longest axis perpendicular to the bottom of the embedding mold to generate transverse sections. The tendon insertion helps with orientation. Preputial glands Embed and section to obtain the largest cut surface.

Post-Mortem Examination and Fixation of Mice

Skin Embed vertically, the cut surface being parallel to the bottom of the mold, to generate transverse sections through the entire thickness of epidermis, dermis, and hypodermis.

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Abdominal organs Adrenal glands Embed to obtain the largest cut surface. Trim the paraffin block to obtain histological sections showing both cortex and medulla. Digestive tract Orient the two halves of the stomach such that the cut surface is in contact with the embedding mold, to generate longitudinal histological sections through the keratinized (i.e., cardial), glandular, and pyloric regions of the stomach. The three segments or the Swiss roll of intestines are embedded in a vertical position, to generate transverse histological sections.

Fat (brown and white) No orientation required. Embed and section to generate the largest cut surface. Kidney Place the cut surface in contact with and parallel to the bottom of the embedding mold to generate longitudinal histological sections through the renal pelvis. Liver Liver is embedded to obtain the largest cut surface. Three sections are taken, 200 μm apart from one another. Lymph nodes Lymph nodes are embedded and sectioned to obtain the largest cut surface. Ovary Embed and section to obtain the largest cut surface. The paraffin block must be trimmed to generate histological sections through the cortex and the medulla. Pancreas Embed and section to obtain the largest cut surface. Prostates, seminal vesicles, urethra, and bladder The group formed by seminal vesicles, dorsal, ventral, and cranial prostates, bladder, and pelvic urethra is taken out of the fixative and first sectioned with a razor blade according to a sagittal median plane through the urethra and urinary bladder. The seminal vesicles with cranial prostates attached to them are then separated by cutting at ∼3 mm from the midline. The distal parts of seminal vesicles that have no cranial prostate attached to them are discarded. The two halves with bladder, ventral and dorsal prostates, and the two pieces of seminal vesicles and cranial prostates are placed in an embedding cassette. The four organ pieces are positioned with their cut surfaces in contact with and parallel to the bottom of the embedding mold. NOTE: In contrast to the human, the mouse prostate comprises four paired lobes (often referred to as “prostates”) situated circumferentially around the urethra, immediately caudal to the urinary bladder—namely, anterior (AP), dorsal (DP), lateral (LP), and ventral prostate (VP). Often, the dorsal and lateral lobes are thought of in combination and referred to as the dorsolateral (DLP) lobe, as they share a ductal system.

Spleen Generate longitudinal sections through the organ.

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Testis Testis with epididymis is placed in the embedding mold with its longest axis lying horizontally, to generate longitudinal histological sections. Uterus One uterine horn is embedded in paraffin with the long axis in vertical position, to generate transverse histological sections. The second horn is embedded with the long axis lying horizontally, to generate longitudinal histological sections. The group formed by the uterus body and vagina is embedded with its long axis lying horizontally. Cervical and thoracic organs Heart The heart is embedded in the vertical position, with the apex towards the mold bottom. Three transverse sections are taken at 3 mm from the apex. Lungs The lungs are embedded and sectioned to obtain the largest cut surface. Thymus The thymus is embedded and sectioned to obtain the largest cut surface. Trachea, esophagus, and thyroid gland The group formed by these organs has a pyramidal shape. It is embedded in paraffin with its base towards the bottom of the mold, to generate transverse histological sections of the trachea, esophagus, and thyroid glands on the same sections. Aorta The aorta is embedded in a vertical position to generate transverse histological sections. Salivary glands The salivary glands are embedded and sectioned to obtain the largest cut surface. Head Eye and Harderian glands The eye and Harderian glands are laid on the bottom of the embedding mold, with the cornea on the left side and the optic nerve on the right. The paraffin block is trimmed to generate histologic sections through the middle of the eye ball. Tongue The tongue is embedded in the vertical position to obtain transverse histological sections. Brain Brain slices are embedded with the anterior surface toward the bottom of the embedding mold. Hypophysis The hypophysis is embedded with the flat surface towards the bottom of the embedding mold.

Post-Mortem Examination and Fixation of Mice

Others If there is an area of particular interest, such as a tumor, trim the organ so that the area of interest is near one surface of the specimen, and place that surface down in the embedding mold.

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REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps.

Formalin (4% formaldehyde solution), neutral buffered, 10% in PBS Add 100 ml of concentrated (37%) formaldehyde (e.g., Carlo Erba, VWR International) to 900 ml phosphate-buffered saline (PBS; see recipe). Label, date and initial. Store up to 3 months at room temperature.

Ethanol solution, 70% Add 408 ml of water to 1 liter of commercial 96% ethanol Phosphate-buffered saline (PBS) 10× PBS stock solution Dissolve the contents of one bottle of powdered salts (Sigma-Aldrich, cat. no. D5652-10L) into 900 ml of deionized water. Make up to 1 liter with water. Autoclave to sterilize. Store at room temperature as long as it stays clear. 1× PBS working solution Mix 100 ml of PBS stock solution with 900 ml of deionized water. Prepare fresh. COMMENTARY Background Information This article describes tissue collection for a systematic histological screen and is designed for systematic investigation of mutant/treated mice. The advantage of this method is the comprehensive view of the macroscopic and microscopic aspects it provides of almost all tissues and organs of the mouse. Depending on the interests of the investigator, this protocol can be adapted for a smaller number of tissues and organs. Specific experimental considerations might, for instance, call for partial approaches directed toward more specific analysis of individual tissues; approaches requiring more specialized dissection or sampling procedures; or combinations of histological and molecular approaches (for example, fixing and freezing separate parts of a single tissue). Necropsy Necropsies and histopathological analysis are highly technical procedures that can yield a great deal of information. In addition to technical skills, a significant amount of prior knowledge is required for morphological analyses. The investigator must be familiar with characteristics of human and mouse anatomy and human and mouse histology. For this type of background information, the reader is referred to Williams et al. (1989) for human anatomy; Krstic (1984), Fawcett (1994), and Sternberg (1997) for human histology; Popesko et al. (1992) for mouse anatomy; and Maronpot et al. (1999) and Mohr et al. (1996) for mouse histology. Histological protocols usually start with

fixation, followed by tissue processing, embedding, and staining (McManus and Mowry, 1960; Lillie, 1965; Luna, 1992; Carson, 1997; also see Internet Resources for Histosearch and StainsFile). Fixation To minimize post-mortem changes in cell structure, the prefixation time (i.e., the time from the surgical excision of the specimen to immersion in the fixative) should be short. Optimal tissue preservation requires delivering the fixative as closely to each cell as possible. A delay in fixation can induce a decrease in observable mitotic figures, and possibly also the transcription of apoptotic factors because of the ensuing anoxia (Cross et al., 1990). Fixation is routinely accomplished by immersion. The kinetics of formalin fixation are such that the process generally requires 24 to 48 hr to achieve complete fixation (Fox et al., 1985). When an organ is immersed in a fixative, its periphery is fixed more rapidly than central areas, which are reached by the fixative after some delay and may undergo post-mortem alterations (Start et al., 1992). Moreover, and unrelated to the problem of post-mortem changes, there is a gradient in the quality of fixation in immersion-fixed tissues, which may yield experimental artifacts (e.g., uneven shrinkage, hardening) leading to false interpretations. Since ideally all cells should be exposed simultaneously to the same concentration of fixative, successful fixation by immersion requires the use of

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small samples. Despite the inevitable gradient of fixation quality in immersion-fixed organs, there is a minimum size required for the tissue sample due to the risk of introducing mechanical damage at the sample surface. It is important to keep in mind that a given fixative stabilizes tissues in a reproducible manner by introducing a consistent tissue artifact, but that the particular changes in the appearance of cells and extracellular matrices are specific to each fixative. For example, the appearance of stained histological sections from a tissue will be different after formalin or Bouin’s fixation. It is worth noting that a variety of proprietary fixatives have been developed and are now proposed for use in necropsy and surgical pathology. These fixatives contain reduced amounts of formalin or none at all, thus reducing the exposure of laboratory workers to this carcinogen. However, in several studies aimed at assessing histomorphology with different formalin substitute fixatives, formalin fixation yielded the highest quality for most tissue stained with hematoxylin-eosin (Fig. 27; Prento and Lyon, 1997; Titford and Horenstein, 2005; authors’ unpub. observ.).

Post-Mortem Examination and Fixation of Mice

Tissue processing Once stabilized and hardened by fixation, tissue specimens are embedded in a solid medium which will support the production of thin tissue slices for microscopic examination. The paraffin embedding method is the most suitable for routine preparation, sectioning, and storage of large numbers of tissue samples. A fixed tissue cannot, however, be directly embedded in paraffin; its free water must first be replaced by solvents. The technique of getting the fixed tissue into paraffin (or other embedding medium) is commonly referred to as tissue processing (Fig. 2). The three steps in this process are dehydration, clearing, and paraffin infiltration (note that bones need to be decalcified prior to dehydration). Dehydration is usually done by incubating the tissue in a series of alcohols, typically 70%, 95%, and 100% ethanol. There is no benefit to slower stepwise dehydrations. It is important that the first alcoholic solution following the 10% neutral buffered formalin contain no more than 70% alcohol, to prevent precipitation of the buffer salts. If water is left in the tissue as a result of improper dehydration, the clearing agent and thus the paraffin will not penetrate the tissue, which will remain soft and be impossible to cut. In contrast, excessive dehydration will remove bound water, yield-

ing shrunken, hard, and brittle specimens. The clearing step consists of replacing dehydrating agent with a substance that is miscible with the embedding medium. For paraffin, a common clearing agent is xylene. Newer clearing agents that present a reduced health hazard are based on limolene or aliphatic long-chain hydrocarbons. Finally, the tissue is infiltrated with the embedding paraffin, which is subsequently solidified by cooling. Paraffin is an inert mixture of hydrocarbons, and can contain plastics, beeswax, or other additives. Commercial paraffins differ in melting temperature for different hardnesses. Paraffin with a melting point of 56◦ to 58◦ C is commonly used for routine histology. A vacuum can be applied inside the tissue processor to assist penetration of paraffin. Sectioning Sectioning is the production of thin slices from a tissue sample. Tissues embedded in paraffin can be sectioned anywhere from 3 to 15 μm thick, with 5 μm being typical for routine use. During the embedding process, the tissues must be oriented properly in the block of paraffin to generate transverse or longitudinal sections. Once the tissues have been embedded, they are cut into sections that are collected on glass slides for subsequent histological, histochemical, or immunohistochemical staining. Sectioning is done with a microtome (Fig. 2), which is essentially just a very sharp knife attached to a mechanism for advancing the paraffin block at standard distances. Once sections are cut, they are floated on warm water to remove any wrinkles, and then transferred to glass slides. The glass slides are then dried in a warm oven to help the sections adhere to the slide. The use of an adhesive mixture such as a gelatin solution enhances the adherence. Staining Most cells contain no endogenous pigments and are thus transparent. Stains are used to establish contrast to render tissue components visible (Fig. 2). Prior to staining, the embedding process must be reversed to remove paraffin from the tissue and allow water-soluble dyes to penetrate the histological section. Slide-mounted sections are deparaffinized by a series of incubations through solvents (e.g., xylene), graded alcohols, and then water. Routine histology uses the stain combination of hematoxylin and eosin (H&E), which allows for visualization of cellular structures. Other stains, referred to as “special stains,” bind

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selectively to particular tissue components and are used to identify specific structures according to diagnostic needs. It is noteworthy that all stain color is artifactual (i.e., it does not represent the natural color of the tissue). A given structure may exhibit very different colors with different stains. For example, collagen is pink with the routine H&E stain, but blue or green with trichrome stains, and red with the Sirius red stain. Therefore, specific aspects of actual structure (location, size, shape) should be used to identify cells and tissues, rather than color, which is unreliable. It should also be stressed that a histological section must be regarded as a single snapshot of the life of the organ. Information regarding the dynamics of a biological process (cell movement, direction of cell proliferation, or cell fate) cannot be deduced solely on the basis of structural patterns.

Critical Parameters and Troubleshooting Necropsy For good morphological preservation, it is critical to accomplish the full dissection within 45 min following the death of the mouse. Samples should be no thicker than 0.5 cm so that the fixative can rapidly penetrate the entire tissue, but long and wide enough to represent the different areas of a tissue as well as any abnormalities. Tissue manipulation is an important parameter for successful morphology preservation. Do not scrape surfaces of tissues or compress them with forceps: samples should be handled carefully by grasping at the edges or manipulated using noninformative regions of the organ (e.g., ligaments, capsule, etc.) or adjacent tissues (e.g., diaphragm for the liver). Be aware that in organs damaged by rough handling, many of the cells will be distorted beyond recognition. Samples that include abnormal areas and surrounding normal areas are best. Avoid unnecessary tissue manipulation. To minimize tissue damage by crushing, try to finish the dissection of a given organ without having to release it and grasp it again. Do not hesitate to use “plenty” of fixative: the ratio of fixative to tissue (v/v) should not be less than 15:1. If the amount of fixative is insufficient, or the sample is too thick, the interior of the specimens will not be properly fixed. Examine the tissues in the container: fresh tissues tend to stick together and to the bottom of the container, which impairs fixation. Swirl the containers periodically to separate the tissues from each other and from the container bottom. If there is a significant amount

of blood in the fixative after tissue collection, replace the solution with fresh fixative. It is worth noting that expert assistance for pathologic and/or histologic analyses can be obtained from qualified comparative pathologists, veterinary pathologists, and/or colleagues with specific expertise. The continued publication of erroneous descriptions and interpretations of mutant mouse phenotypes by untrained and inexperienced investigators is a real concern. The answer to this problem is to ensure that pathomorphological characterizations of mice are performed by qualified comparative pathologists (Bolon et al., 2008). Incompatibilities between the requirements of tissue processing for histological purposes and tissue collection during molecular biology applications (e.g., DNA and/or RNA sampling) can arise when the procedures are performed simultaneously. For example, RNA cannot be isolated from intestines that were injected with fixative solution at the beginning of the necropsy procedure. To avoid incompatibilities between the two procedures, it is advised to use different mice for necropsy and tissue collection, for molecular biology applications. As in any phenotypic analysis, it is critical that cohorts be carefully planned for a histological analysis, and that mutant animals should be compared with strictly comparable normal animals (i.e., of the same gender, age, strain, circadian phase, etc.). Fixation Formalin. Commercial solutions of concentrated formalin sometimes become turbid during storage through the production of paraformaldehyde, which decreases their concentration of formalin. For the sake of standardization, a turbid solution should be discarded, and any opened bottle of concentrated formalin should be used within 6 months. Formaldehyde has a greater chance for oxidation at low concentrations, and eventually the pH of a 10% neutral buffered formalin solution will start to decrease in spite of the buffer. In addition, 10% buffered formalin solutions will have slowly increasing concentrations of formic acid (a byproduct of aging formaldehyde) upon storage. This promotes the clumping of proteins instead of the crosslinking that formaldehyde normally promotes. Therefore, if the tissue is destined for antibody staining at a later time, 10% buffered formalin solutions should be used within 3 months (McManus and Mowry, 1960; also see Internet Resources). A loss of antigenicity during

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prolonged formalin fixation is caused both by extensive cross-linking of proteins (preventing access of the antibody to the antigen) and by chemical alteration of the antigen (i.e., change in epitope structure) (Leong and Leong, 2007). However, antigens can often survive a short exposure to formalin. On the other hand, the duration of fixation must be long enough to preserve the integrity of the tissue and prevent antigen diffusion. This is a dilemma known as the “immunohistochemical compromise.” Although tissues may be stored in buffered formalin indefinitely without morphological damage, they should not be fixed for more than 24 hr if immunohistochemistry will be performed (Grizzle, 2009). Bouin’s solution. Due to deleterious effects on tissue morphology of prolonged exposure to the acids present in Bouin’s solution, it is recommended not to keep specimens in this fixative for more than 1 month before paraffin embedding

Time Considerations Knowledge of tissue location and experience in tissue dissection and handling are indispensable prior to necropsy to ensure tissue collection in the minimum amount of time. A complete necropsy by an experienced individual requires ∼45 min per mouse. Collecting tissue for RNA analyses should take no longer than 1 to 10 min, depending on the organ. If RNA sampling is desired, the dissections should be performed by two persons, one to dissect the tissues destined for RNA analysis and freeze them in liquid nitrogen, and the other to collect tissues for histological analyses.

Literature Cited Bolon, B., Brayton, C., Cantor, G.H., Kusewitt, D.F., Loy, J.K., Sartin, E.A., Schoeb, T.R., Sellers, R.S., Schuh, J.C.L., and Ward, J.M. 2008. Editorial: Best pathology practices in research using genetically engineered mice. Vet. Pathol. 45:939-940. Brayton, C., Justice, M., and Montgomery, C.A. 2001. Evaluating mutant mice: Anatomic pathology. Vet. Pathol. 38:1-19. Carson, F.L. 1997. Histotechnology: A Self Instructional Text, 2nd ed. ASCP Press, Chicago, Illinois.

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Costet, P., Legendre, C., More, J., Edgar, A., Galtier, P., and Pineau, T. 1998. Peroxisome proliferator activated receptor alpha-isoform deficiency leads to progressive dyslipidemia with sexually dimorphic obesity and steatosis. J. Biol. Chem. 273:29577-29585. Cross S.S., Start R.D., and Smith J.H. 1990. Does delay in fixation affect the number of mitotic fig-

ures in processed tissue? J. Clin. Pathol. 43:597599. Fan, C.Y., Pan, J., Usuda, N., Yeldandi, A.V., Rao, M.S., and Reddy, J.K. 1998. Steatohepatitis, spontaneous peroxisome proliferation and liver tumors in mice lacking peroxisomal fatty acyl-CoA oxidase: Implications for peroxisome proliferator-activated receptor alpha natural ligand metabolism. J. Biol. Chem. 273:1563915645. Fawcett, D.W. 1994. Bloom and Fawcett: A Textbook of Histology, 12th ed. Saunders Co., Philadelphia, Pennsylvania. Fox C.H., Johnson F.B., Whiting J., and Roller P.P. 1985. Formaldehyde fixation. J. Histochem. Cytochem. 33:845-853. Grizzle, W.E. 2009. Special symposium: Fixation and tissue processing models. Biotech. Histochem. 84:185-193. Harvey, M., McArthur, M.J., Montgomery, C.A. Jr., Butel, J.S., Bradley, A., and Donehower, L.A. 1993. Spontaneous and carcinogen-induced tumorigenesis in p53-deficient mice. Nat. Genet. 5:225-229. Huang, J., Powell, W.C., Khodavirdi, A.C., Wu, J., Makita, T., Cardiff, R.D., Cohen, M.B., Sucov, H.M., and Roy-Burman, P. 2002. Prostatic intraepithelial neoplasia in mice with conditional disruption of the retinoid X receptor alpha allele in the prostate epithelium. Cancer Res. 62:48124819. Khetchoumian, K., Teletin, M., Tisserand, J., Mark, M., Herquel, B., Ignat, M., Zucman-Rossi, J., Cammas, F., Lerouge, T., Thibault, C., Metzger, D., Chambon, P., and Losson, R. 2007. Loss of Trim24 (Tif1alpha) gene function confers oncogenic activity to retinoic acid receptor alpha. Nat. Genet. 39:1500-1506. Kim, M.J., Cardiff, R.D., Desai, N., BanachPetrosky, W.A., Parsons, R., Shen, M.M., and Abate-Shen, C. 2002. Cooperatively of Nkx3.1 and Pten loss of function in a mouse model of prostate carcinogenesis. Proc. Natl. Acad. Sci. U.S.A. 99:2884-2889. Krstic, R.V. 1984. Illustrated Encyclopedia of Human Histology. Springer, Berlin, Germany. Leong, T.Y. and Leong, A.S. 2007. How does antigen retrieval work? Adv. Anat. Pathol. 14:129131. Li, M., Messaddeq, N., Teletin, M., Pasquali, J.L., Metzger, D., and Chambon, P. 2005. Retinoid X receptor ablation in adult mouse keratinocytes generates an atopic dermatitis triggered by thymic stromal lymphopoietin. Proc. Natl. Acad. Sci. U.S.A. 102:1479514800. Lillie, R.D. 1965. Histopathologic Technique and Practical Histochemistry, 3rd ed. McGraw-Hill, New York. Liu, J.L., Yakar, S., and LeRoith, D. 2000. Mice deficient in liver production of insulin-like growth factor I display sexual dimorphism in growth hormone-stimulated postnatal growth. Endocrinology 141:4436-4441.

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Luna, L.G. 1992. Histopathological Methods and Color Atlas of Special Stains and Tissue Artifacts. Johnson Printers, Downers Grove, Illinois.

Turgeon, B. and Meloche, S. 2009. Interpreting neonatal lethal phenotypes in mouse mutants: Insights into gene function and human diseases. Physiol. Rev. 89:1-26.

Maronpot, R.R., Boorman, G.A., and Gaul, B.W. 1999. Pathology of the Mouse: Reference and Atlas. Cache River Press, Vienna, Illinois.

Werner, M., Chott, A., Fabiano, A., and Battifora, H. 2000. Effect of formalin tissue fixation and processing on immunohistochemistry. Am. J. Surg. Pathol. 24:1016-1019.

McManus, J.F.A. and Mowry, R.W. 1960. Staining Methods. Histologic and Histochemical. Harper and Row, New York. Mohr, U., Dungworth, D.L., Capen, C.C., Carlton, W.W., Sundberg, J.P., and Ward, J.M. 1996. Pathobiology of the Aging Mouse, Volumes 1 and 2. ILSI Press, Washington, D.C. Moolenbeek, C. and Ruitenberg, E.J. 1981. The “Swiss roll”: A simple technique for histological studies of the rodent intestine. Lab Anim. 15:57-59. Morel, G. and Cavalier, A. 2000. In Situ Hybridization in Light Microscopy (Methods in Visualization). CRC Press, Boca Raton, Florida. Naf, D., Krupke, D.M., Sundberg, J.P., Eppig, J.T., and Bult, C.J. 2002. The Mouse Tumor Biology Database: A public resource for cancer genetics and pathology of the mouse. Cancer Res. 62:1235-1240. Polak, J.M. and Van Noorden, S. 2003. Introduction to immunohistochemistry. Bios Scientific Publishers, Oxford, United Kingdom. Popesko, P., Rajtova, V., and Horak, J. 1992. Colour Atlas of Anatomy of Small Laboratory Animals, Volume 2. Wolfe Publishing, London. Prento, P. and Lyon, H. 1997. Commercial formalin substitutes for histopathology. Biotech. Histochem. 72:273-282. Renshaw, S. 2007. Immunohistochemistry. Scion Publishing, Bloxham Mill, U.K. Srinivasan, M., Sedmak, D., and Jewell S. 2002. Effect of fixatives and tissue processing on the content and integrity of nucleic acids. Am. J. Pathol. 161:1961-1971. Start, R.D., Layton, C.M., Cross S.S., and Smith, J.H. 1992. Reassessment of the rate of fixative diffusion. J. Clin. Pathol. 45:1120-1121. Sternberg, S.S. 1997. Histology for Pathologists, 2nd ed. Lippincott-Raven, Philadelphia, Pennsylvania. Titford, M.E. and Horenstein, M.G. 2005. Histomorphologic assessment of formalin substitute fixatives for diagnostic surgical pathology. Arch. Pathol. Lab. Med. 129:502-506.

Wilkinson, D.G. 1999. In situ hybridization. A practical approach. Oxford University Press, Oxford. Williams, P.L., Warwick, R., Dyson, M., and Bannister, L.H. 1989. Gray’s Anatomy, 37th ed. Churchill Livingstone, London.

Internet Resources http://www.eumorphia.org/ The EUMORPHIA Web site, provides information about understanding human disease through mouse genetics. http://www.pathbase.net/ Pathbase, a database of histopathology photomicrographs and macroscopic images derived from mutant or genetically manipulated mice. http://ctrgenpath.net/static/atlas/mousehistology/ Provides an electronic atlas for the anatomy and histology of the mouse. http://www.deltagen.com/target/histologyatlas/ HistologyAtlas.html Provides an electronic atlas for the histology of the mouse. http://www.dundee.ac.uk/histology/welcome. html A workbook with a set of microscope slides that illustrate the basic tissues and organs of the body. http://www.histosearch.com/ Histosearch is a search engine that searches over 20,000 Web pages from histology-related sites. http://stainsfile.info/StainsFile/jindex.html StainsFile is a useful site with general information and resources for histology. http://swehsc.pharmacy.arizona.edu/exppath/ resources/formaldehyde.html Information on formaldehyde fixatives at the Southwest Environmental Health Sciences Center (SWEHSC), University of Arizona College of Pharmacy. http://www.ccac.ca/ The Canadian Council on Animal Care (CCAC) is the national organization responsible for setting and maintaining standards for the care and use of animals in science in Canada.

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Generation of Spatio-Temporally Controlled Targeted Somatic Mutations in the Mouse Daniel Metzger1 and Pierre Chambon1 1

Institut de G´en´etique et de Biologie Mol´eculaire et Cellulaire, Universit´e de Strasbourg, and Coll`ege de France, Illkirch, France

ABSTRACT The generation of ligand-activated site-specific Cre recombinases has led to the development of cell type–specific temporally controlled targeted somatic mutagenesis in the mouse. We illustrate this technique using K14-Cre-ERT2 transgenic mice that express the tamoxifen (tam)-activatable Cre-ERT2 recombinase in epidermal basal keratinocytes to induce mutations in epidermal keratinocytes of adult mice. Our highly reproducible technique, based on induction of Cre-ERT2 recombinase activity by tamoxifen administration at low doses (once daily 100-μg intraperitoneal injection for 5 days), has allowed the generation of site-directed somatic mutations of numerous genes in mouse epidermal keratinocytes, and several mouse models of human diseases. The present step-by-step protocol describes how to introduce temporally controlled targeted mutations in epiderC 2011 by John mal keratinocytes of adult mice. Curr. Protoc. Mouse Biol. 1:55-70  Wiley & Sons, Inc. Keywords: Cre-ERT2 r tamoxifen r loxP r keratinocytes r skin

INTRODUCTION Major progress in understanding the function of gene products in mammalian development, as well as in adult homeostasis and pathophysiology, was achieved over recent years by studying mice bearing site-directed mutations. However, targeting mutations in the germ line induces, in many instances, developmental aberrations, embryonic lethality, or compensatory effects due to functionally redundant genes. These limitations often preclude the determination of the function of a given gene product in a defined subset of cells at a given time during the life of the animal, and therefore make it hard to discriminate between cell-autonomous and non-cell-autonomous functions. Furthermore, germline mutations are inadequate to generate mouse models of human diseases resulting from somatic mutations, such as most forms of cancer (Metzger and Chambon, 2001; Jonkers and Berns, 2002). To circumvent these problems, methods to achieve conditional gene targeting have been developed. They are based mainly on the properties of the bacteriophage P1 site-specific Cre recombinase, which efficiently and faithfully excises a DNA segment flanked by two loxP sites (floxed DNA) in animal cells. Thus, spatially or temporally controlled somatic mutations can be obtained in mice by controlling its expression with a cell-specific or an inducible promoter, respectively (Rajewsky et al., 1996). However, as these conditional gene-targeting systems can only be either spatially or temporally controlled, they present a number of limitations. The ligand-inducibility of the recombinase activity of chimeric proteins, in which the Cre recombinase is fused to mutated ligand-binding domains of steroid hormone receptors (Cre-ERT , Cre-ERTM , Mer-Cre-Mer, and Cre-PR), has been exploited to

SpatioTemporally Controlled Targeted Somatic Mutations

Current Protocols in Mouse Biology 1: 55-70, March 2011 Published online March 2011 in Wiley Online Library (wileyonlinelibrary.com). DOI: 10.1002/9780470942390.mo100128 C 2011 John Wiley & Sons, Inc. Copyright 

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generate ligand-inducible spatio-temporally controlled targeted somatic mutations in the mouse (Feil, 2007). In this respect, the tamoxifen (tam)-inducible activity of the Cre-ERT2 recombinase, which results from the fusion of the Cre recombinase and a triple-mutated ligand-binding domain of the human estrogen receptor ERα, is of particular interest (Feil et al., 1997; Indra et al., 1999). Indeed, cell-specific expression of Cre-ERT2 in transgenic mice allows, for most “floxed” loci, efficient tamoxifen-inducible Cre-mediated recombination, without background recombinase activity in the absence of tamoxifen (el Marjou et al., 2004; Schuler et al., 2004, 2005; Imayoshi et al., 2006; Slezak et al., 2007; Ratnacaram et al., 2008; Wendling et al., 2009).

STRATEGIC PLANNING To generate temporally controlled targeted mutations of a given gene in specific cell types, it is highly recommended to select well-characterized Cre-ERT2 transgenic lines. In most cases, it is required that such transgenic lines (i) express Cre-ERT2 selectively in the target cells, (ii) exhibit no recombinase activity before tamoxifen administration, and (iii) generate efficient Cre-mediated recombination after tamoxifen administration. It is also important that loxP sites be inserted in appropriate genomic regions of the target gene, to ensure that Cre-mediated deletion of the loxP-flanked DNA segment results in the expected genetic alteration. BASIC PROTOCOL

GENERATION OF TEMPORALLY CONTROLLED MUTATIONS IN SKIN USING MICE EXPRESSING Cre-ERT2 SELECTIVELY IN KERATINOCYTES The skin is a complex organ composed of the epidermis and its appendages, as well as the dermis. The epidermis, a highly dynamic stratified epithelium principally made of keratinocytes, serves as a protective barrier against loss of body fluids and external environmental insults. The epidermal innermost cells that are attached to the basement membrane and express cytokeratins K5 and K14 form the basal proliferative layer, from which keratinocytes periodically withdraw from the cell cycle and commit to terminally differentiate while migrating into the suprabasal layers. Terminally differentiated keratinocytes that form the cornified layer are sloughed off the skin surface and are continuously replaced by newly differentiating cells (Fuchs, 2007). To generate temporally controlled mutations of target genes in skin keratinocytes, K14Cre-ERT2 transgenic mice that express Cre-ERT2 in basal keratinocytes (Indra et al., 2000) are used. The protocol presented here for efficient temporally controlled target gene ablation in epidermal keratinocytes of adult mice involves three major steps: (i) generation of premutant mice bearing the K14-Cre-ERT2 transgene and “floxed” alleles (L2) of the chosen target gene (called hereafter GeneX), (ii) induction of the target gene mutation in epidermal keratinocytes by intraperitoneal tamoxifen administration, and (iii) characterization of the selectivity and efficiency of target gene ablation in epidermis.

Materials

SpatioTemporally Controlled Targeted Somatic Mutations

K14-Cre-ERT2 mice (Indra et al., 2000; Li et al., 2000; US patent no.7112715 and European patent no.1 692 936 cover commercial use of Cre-ERT2 expressing mice) GeneXL2/+ or GeneXL2/L2 mice, which bear one or two floxed target alleles Custom-designed oligo primers (see Table 1 and Figs. 1 and 2; also see recipe) Direct PCR lysis solution (see recipe) Cre PCR master mix (see Table 2) GeneX PCR master mix (see Table 3)

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Ethidium bromide–stained 2.0% agarose gel in TBE electrophoresis buffer (see recipe) (Armstrong and Schulz, 2008) DNA Ladder Gene Ruler (Euromedex, cat. no. SM0331) Tamoxifen ( see recipe) Isoflurane 70% (v/v) ethanol Dispase solution (see recipe) Proteinase K digestion buffer (see recipe) Ethanol (EtOH) Sterile water Tris·Cl, pH 8.0 1:1 phenol:chloroform (see recipe) 1.5-ml microcentrifuge tubes 55◦ C incubator 85◦ C water bath 0.2-ml PCR microtubes (Dominique Dutscher, cat. no. 01600) Thermal cycler (Gene Amp PCR 9700; Applied Biosystems) Gloves (Laboratories Euromedis, cat. no. 127587) Syringe (1-ml equipped with a 25-G needle; Terumo, cat. no. BS-01 H2516) Gas anesthesia station for rodents (TEM) Animal electric shaver Surgical instruments: dissection scissors, straight surgical forceps, needle holder Suture materials (Ethibond Excel polyester 3-0; Ethicon, cat. no. X32040) Microcentrifuge (Eppendorf, cat. no. 5415D) Additional reagents and equipment for running PCR products on an ethidium bromide–stained 2.0% agarose gel in TBE electrophoresis buffer (Armstrong and Schulz, 2008) NOTE: All protocols using live animals must first be reviewed and approved by an Institutional Animal Care and Use Committee (IACUC) and must follow officially approved procedures for the care and use of laboratory animals.

Table 1 Custom Designed Oligonucleotides

For Cre-ERT2 (see Fig. 1) TK139

5 -ATTTGCCTGCATTACCGGTC-3

TK 141

5 -ATCAACGTTTTGTTTTCGGA-3

For internal control (myogenin) ADV28

5 -TTACGTCCATCGTGGACAGC-3

ADV30

5 -TGGGCTGGGTGTTAGCCTTA-3

For GeneX WT, L2 and L- alleles, P1, P2, and P3 See Figure 2 For RXRα alleles P1, BAA239

5 -TCAAGTGAGGTGGACATTAGGATG-3

P2, BAA982

5 -CTGGAAGAGGATGGGCACTATTCT-3

P3, BAA983

5 -AAACTGCAAGTGGCCTTGAGAAGAA-3

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Figure 1 Schematic structure of the K14-Cre-ERT2 transgene. The human K14 promoter, the Cre-ERT2 coding sequence, and the simian virus 40 polyadenylation signal (polyA) are represented by blue, yellow, and purple boxes, respectively. The rabbit β-globin intron and splice donor- and acceptor-sites are depicted by a line and pink boxes, respectively. The position of the PCR primers TK139 and TK141, and the length of the PCR-amplified DNA segment are indicated. The sequence of the K14-Cre-ERT2 transgene is available upon request.

Figure 2 Schematic diagram of a target GeneX WT allele, a floxed GeneX L2 allele, and a GeneX L- allele generated by Cre-mediated excision. Two exons (En and En+1) of GeneX are boxed. P1, P2, and P3 are PCR primers used to identify GeneX WT, L2, and L- alleles are indicated. Arrowheads represent loxP sites.

Generate premutant K14-Cre-ERT2(tg/0) /GeneXL2/L2 mice 1. Generate K14-Cre-ERT2(tg/0) /GeneXL2/L2 premutant mice bearing the K14-Cre-ERT2 transgene (Fig. 1) and two floxed GeneX (L2) alleles (Fig. 2), as well as appropriate control mice, by breeding 7- to 8-week-old hemizygous K14-Cre-ERT2(tg/0) mice with GeneXL2/+ mice or with GeneXL2/L2 mice, bearing one or two allele(s) of the floxed target gene (GeneX), respectively. In the example below, RXRαL2/+ mice (Li et al., 2000) are used.

SpatioTemporally Controlled Targeted Somatic Mutations

2. Design primers to identify K14-Cre-ERT2(tg/0) /GeneXL2/+ pups by PCR-based DNA amplification from tail biopsies. To reveal GeneX wild-type (WT) and L2 alleles, design a forward P1 primer and a reverse P2 primer, located 5 and 3 of the 5 loxP site, respectively, in order to generate two PCR-amplified DNA segments of different length from the two alleles (Fig. 2).

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3. Synthesize GeneX P1 and P2 primers, as well as TK139 and TK141 to identify the K14-Cre-ERT2 transgene, and ADV28 and ADV30 primers to identify myogenin (see Table 1). 4. Take a 3-mm tail biopsy from 1- to 2-week-old offspring and place in a 1.5-ml microcentrifuge tube. Biopsies can be stored for several weeks at −20◦ C.

5. Add 150 μl of direct PCR lysis solution containing 60 μg of proteinase K into the tube containing the tail biopsy. CAUTION: Proteinase K is an irritant; wear gloves and safety glasses when handling. The use of proteinase K–containing direct PCR lysis reagent allows fast sample preparation for efficient genomic DNA PCR amplification.

6. Place the tubes for 5 to 12 hr in an incubator at 55◦ C and then for 45 min in an 85◦ C water bath to lyse the tail cells. Crude lysates may be stored up 1 week at 4◦ C and at −20◦ C for months.

7. Set up PCR amplification reactions to identify the presence of the K14-Cre-ERT2 transgene. For ten reactions, assemble the Cre PCR (10R) master mix as shown in Table 2. 8. Dispense 2 μl of crude tail lysates (about 0.5 μg/μl) from the pups to be genotyped into 0.2-ml thermal cycler tubes and add 28 μl of Cre PCR (10R) master mix. Perform control reactions with 2 μl crude lysates from WT and K14-Cre-ERT2 transgenic mice, as well as with 2 μl direct PCR lysis solution. 9. Set up PCR amplification reactions to identify GeneX WT and L2 alleles. For ten reactions, assemble the GeneX PCR (10R) master mix as shown in Table 3. 10. Dispense 2 μl of crude tail lysates from the pups to be genotyped into 0.2-ml thermal cycler tubes and add 28 μl of GeneX PCR (10R) master mix. Perform control reactions with 2 μl of crude lysates from WT and GeneXL2/L2 mice, as well as with 2 μl of direct PCR lysis solution. Table 2 Cre PCR Master Mix for 10 Reactions to Identify the K14-Cre-ERT2 Transgene

Reagent

Volume of reagent

Initial concentration of reagent

Final concentration of reagent

10×

1×

3 μl

30 μl

10 mM each

100 μM

0.3 μl

3 μl

Forward primer TK139

100 μM

0.33 μM

0.1 μl

1 μl

Reverse primer TK141

100 μM

0.33 μM

0.1 μl

1 μl

Forward primer ADV28

100 μM

0.33 μM

0.1 μl

1 μl

Reverse primer ADV30

100 μM

0.33 μM

0.1 μl

1 μl

5 u/μl

1U

0.2 μl

2 μl

24.1 μl

241 μl

PCR buffer (see recipe) dNTPs (see recipe)

Taq DNA polymerase

a

Water

1 reaction 10 reactions

a 5 U/μl; Sigma, cat. no. D4545.

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Table 3 GeneX PCR Master Mix for 10 Reactions to Identify GeneX WT and L2 Alleles

Reagent

Volume of reagent

Initial concentration of reagent

Final concentration of reagent

1 reaction

10 reactions

10×

1×

3 μl

30 μl

10 mM each

100 μM

0.3 μl

3 μl

Forward primer P1 (i.e., BAA239 for RXRα)

100 μM

0.33 μM

0.1 μl

1 μl

Reverse primer P2 (i.e., BAA982 for RXRα)

100 μM

0.33 μM

0.1 μl

1 μl

5 u/μl

1U

0.2 μl

2 μl

24.3 μl

243 μl

PCR buffer (see recipe) dNTPs (see recipe)

Taq polymerasea Water a 5 U/μl; Sigma, cat. no. D4545.

11. Place the tubes prepared in steps 8 and 10 in a thermal cycler, and run the following cycling program: 1 cycle: 30 cycles:

1 cycle:

5 min 30 sec 30 sec 30 sec 7 min

94◦ C (initial denaturation) 94◦ C (denaturation) 55◦ C (annealing) 72◦ C (extension) 72◦ C (final extension).

PCR products can be stored for several days at 20◦ C and for several weeks at −20◦ C.

12. Run PCR products and DNA ladder gene ruler on an ethidium bromide–stained 2.0% agarose gel in TBE electrophoresis buffer (Armstrong and Schulz, 2008). The K14-Cre-ERT2 transgene is revealed by a 350-bp DNA segment amplified with primers TK139 and TK141 (Fig. 1). A 230-bp DNA segment, amplified from an endogenous mouse gene (myogenin) with primers ADV28 and ADV30 included in the reaction, serves as an internal control to ensure that genomic DNA can be efficiently amplified from each lysate (Fig. 3). Control reactions with lysates from WT and K14-Cre-ERT2(tg/0) transgenic mice, as well as with lysis solution (no DNA), ensure that the K14-Cre-ERT2 transgene is specifically amplified under the used experimental conditions. One PCR product is amplified with primers P1 and P2 from tail biopsy lysates of pups bearing two WT target alleles, whereas two DNA segments, differing by at least 34 bp (i.e., the length of a LoxP site), are amplified from genomic DNA of pups bearing one WT and one floxed target allele. Control reactions with lysates from WT and from GeneXL2/+ mice, as well as with lysis solution (no DNA), ensure that GeneX WT and L2 alleles are specifically amplified under the used experimental conditions (Fig. 4).

13. Breed 7- to 8-week-old K14-Cre-ERT2(tg/0) /GeneXL2/+ mice with GeneXL2/+ mice, and identify K14-Cre-ERT2(tg/0) /GeneXL2/L2 offspring (pre-mutant mice), as well as K14-Cre-ERT2(tg/0) and K14-Cre-ERT2(0/0) /GeneXL2/L2 control littermates, by genotyping as described in steps 4 to 12.

SpatioTemporally Controlled Targeted Somatic Mutations

Induce LoxP-flanked DNA excision 14. Thaw an aliquot of tamoxifen (1 mg/ml in sunflower oil) and of vehicle solution, and mix well. CAUTION: Wear gloves and glasses; avoid exposure as tamoxifen may impair fertility and cause cancer.

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Figure 3 Identification of K14-Cre-ERT2 transgenic mice by PCR-mediated tail genomic DNA amplification. PCR-amplified DNA segments were run on a 2% agarose gel. Lanes 5 to 9, amplification products from 5 pups (Pu 1 to 5). Lanes 2 and 3, amplification products from K14-Cre-ERT2 and WT mice, respectively. Lane 4, PCR reaction without genomic DNA (lysis solution). Lane 1, DNA ladder (L). The size of the DNA segments is given in base pairs. Cre, DNA segment amplified from the K14-Cre-ERT2 transgene with the primer pair TK139 and TK141 (Fig. 1). IC (Internal control): DNA segment amplified from an endogenous mouse gene (myogenin) with the primer pair ADV28 and ADV30. Pu1, Pu3, and Pu4 are transgenic for K14-Cre-ERT2 ; Pu2 and Pu5 are not.

Figure 4 Identification of RXRα WT and L2 alleles by PCR-mediated tail genomic DNA amplification. PCR-amplified DNA segments with the primer BAA239 (P1) and BAA982 (P2) were run on a 2% agarose gel. Lanes 5 to 9, amplification products from 5 pups (Pu 1 to 5). Lanes 2 and 3, amplification products from floxed RXRαL2/L2 and WT mice, respectively. Lane 4, PCR reaction without genomic DNA (lysis solution). Lane 1, DNA ladder (L). The size of the DNA segments is in base pairs. The position of the PCR product amplified with BAA239 and BAA982 from RXRα WT (WT; 158 bp) and L2 floxed alleles (L2; 199 bp) are indicated. Pu1 and Pu5 show RXRα WT alleles; Pu2, Pu3, and Pu4 show one RXRα WT allele and one RXRα L2 allele.

15. Inject 8-week-old K14-Cre-ERT2(tg/0) /GeneXL2/L2 premutant mice intraperitoneally with 100 μl tamoxifen (0.1 mg) daily for 5 days with a 1-ml syringe equipped with a 25-G needle to induce loxP-flanked GeneX excision in epidermal keratinocytes. To control for possible tamoxifen-induced side effects, also inject age- and sex-matched K14-Cre-ERT2(tg/0) /GeneX+/+ and K14-Cre-ERT2(0/0) /GeneXL2/L2 mice with 100 μl tamoxifen solution (0.1 mg). Absence of background recombination is assessed in vehicle (oil)-treated K14-CreERT2(tg/0) /GeneXL2/L2 mice. To avoid possible exposure of oil-treated mice to tamoxifen (e.g., through contact with urine and/or stool from tamoxifen-treated mice), oil- and tamoxifen-treated mice are housed in distinct cages. T2

Note that Cre-ER recombinase activity can also be induced by oral gavage of tamoxifen (Park et al., 2008) or by topical application of 4-hydroxytamoxifen (Ruzankina et al., 2007; see Commentary section).

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Characterize floxed DNA excision 16. Design and synthesize GeneX oligo primer P3. Identification of recombined GeneX allele (L-) by PCR-based DNA amplification requires the design of reverse primer P3, located 3 of the 3 loxP site (see Fig. 2).

17. Prepare the samples for analysis using the following options (depending on whether the main aim is to determine the specificity of DNA excision or its efficiency before phenotypic analyses, respectively): a. Sacrifice tamoxifen- and oil-treated mice (approximately two weeks after the last tamoxifen injection) by cervical dislocation, shave an area 2-cm2 from the back skin, and take dorsal skin, ear, and tail samples, as well as other organs, if required. b. Alternatively, place the mouse in the chamber of a gas anesthesia station containing isoflurane. After the mouse has lost consciousness, shave an area 2-cm2 from the back skin and sterilize the areas to be sampled with 70% ethanol. Take a 0.5-cm2 biopsy of the selected back skin and suture the wound, and/or 2-mm biopsies from the tip of the ear and tail. To check whether the desired genetic modification is present in keratinocytes of all epidermal layers, biopsies should be taken at least 2 weeks after the last tamoxifen administration, as it takes ∼2 weeks for complete renewal of epidermal suprabasal keratinocytes in mouse skin. To ensure that excision of the floxed target gene is efficiently induced in epidermal stem cells, we recommend analyzing skin biopsies several months after tamoxifen administration.

18. Isolate the epidermis from the dermis by removing bone from the tail biopsy and incubating the ear and tail skin in 2 ml of dispase solution (4 mg/ml in PBS) overnight at 4◦ C (or for 2 hr at 37◦ C). Separate the dermis from the epidermis with forceps and transfer to microcentrifuge tubes. Separation of epidermis from dermis of back skin is more tedious, as the number of suprabasal layers is lower than in ear or tail skin.

19. Extract genomic DNA from the samples by adding 500 μl of proteinase K digestion buffer and 7.5 μl of proteinase K stock solution (20 mg/ml), and incubate overnight at 55◦ C. 20. Add an equal volume of phenol/chloroform (1:1), mix well, and microcentrifuge 5 min at 14,000 × g, room temperature. 21. Transfer the supernatant to a new tube. Add 1 ml EtOH to the tube, mix, and microcentrifuge 5 min at 14,000 × g, room temperature. Wash the DNA pellet with 70% ethanol, air dry, and dissolve in 100 μl sterile water. CAUTION: Phenol is an irritant and causes burns; wear gloves and safety glasses when handling. DNA can be stored up to several weeks at 4◦ C.

22. For each sample, set up two PCR amplification reactions in 0.2-ml thermal cycler tubes: one with primer pair P1/P2 and one with primer pair P1/P3 to identify GeneX L2 and L- alleles, respectively, according to steps 9 to 11. SpatioTemporally Controlled Targeted Somatic Mutations

23. Run the PCR products and DNA ladder gene ruler on an ethidium bromide–stained 2.0% agarose gel in TBE, and estimate the relative amount of amplified alleles. PCR amplification reactions of the L- alleles of a given gene with P1/P3 might require optimization. It is highly recommended to validate the detection of L- alleles on genomic

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DNA isolated from GeneXL-/+ mice, if available. PCR amplification of L2 and L- alleles with the three primers in one reaction is possible, but might require further standardization.

24. Analyze the phenotype of mice in which the target gene is ablated by macroscopic examination, functional tests, and histological analyses.

REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps.

Direct PCR lysis solution Add 30 μl of proteinase K stock solution (20 mg/ml; see recipe) to 1.5 ml direct PCR lysis reagent (Viagen Biotech, cat. no. 102-T). Prepare just before use. CAUTION: When handling, wear gloves and safety glasses.

Dispase solution, 4 mg/ml Dissolve 40 mg of dispase (Invitrogen, cat. no. 17105-041) in 10 ml phosphatebuffered saline (PBS; Sigma Aldrich, cat. no. D-5652) and mix well. Prepare just before use. CAUTION: When handling, wear gloves and safety glasses.

dNTP (10 mM each) For 1 ml: Mix 100 μl of 100 mM dATP (Amersham, cat. no. 27-2050-03), 100 μl of 100 mM dCTP (Amersham, cat. no. 27-2060-03), 100 μl of 100 mM dGTP (Amersham, cat. no. 27-2070-03), and 100 μl of 100 mM dTTP (Amersham, cat. no. 27-2080-03) in 600 μl of sterile distilled water. Store as 100-μl aliquots for several months at 4◦ C or −20◦ C.

Oligonucleotides Dissolve oligonucleotides (dATP, dCTP, dGTP, and dTTP; see recipe for dNTPs) in autoclaved deionized water at 100 mM. Store up to several months at –20◦ C.

PCR buffer, 10× 500 mM KCl (Sigma Aldrich, cat. no. 31248) 100 mM Tris·Cl, pH 8.8 15 mM MgCl2 (Sigma Aldrich, cat. no. 63068) Autoclave Divide into 0.5-ml aliquots Store up to several months at −20◦ C This buffer is more efficient for genomic DNA amplification from tail biopsy lysates than buffers supplied with commercial Taq polymerase.

Phenol:chloroform (1:1) solution Mix equal parts of stabilized phenol (Tris saturated, pH 7.5 to 8; Eurobio, cat. no. 018335) and chloroform (Carlo Erba, cat. no. 438603). Store in a light-tight bottle for several months at 4◦ C. CAUTION: When handling, wear gloves and safety glasses. Prepare the solution in a safety biohazard hood.

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Primer design Identify the GeneX wild-type (WT), L2, and L- alleles based on PCR amplification of 200- to 500-bp genomic DNA segments using 3 primers: a forward primer P1 and a reverse primer P2 located 5 and 3 of the 5 loxP site, respectively, and a reverse primer P3 located 3 of the 3 loxP site (see Fig. 2). Thus, the primer pair P1/P2 generates two PCR products of different length from the WT and L2 alleles, whereas the primer pair P1/P3 generates a PCR product from the L- allele (the larger distance between P1 and P3 primers on the WT and L2 alleles usually impairs efficient PCR amplification of these alleles). Such primer pairs are designed with the Primer 3 (v.0.4.0) software (http://frodo.wi.mit.edu/primer3/input.htm).

Proteinase K digestion buffer Combine 50 mM Tris·Cl (pH 8.0), 5 mM EDTA (Euromedex, cat. no. EDTA23), 1% SDS (Euromedex, cat. no. EU0660), and 0.2 M NaCl (Sigma Aldrich, cat. no. 31434). Autoclave, and store up to several months at room temperature (18◦ to 22◦ C).

Proteinase K stock solution at 20 mg/ml Dissolve 1 g of proteinase K (Sigma Aldrich, cat. no. P-6556) in 50 ml of autoclaved TE buffer (10 mM Tris·Cl, pH 8.0, 1 mM EDTA). Divide into 0.5-ml aliquots and store for several months at −20◦ C. CAUTION: Proteinase K is highly irritant for the eyes, airways, and skin. Wear gloves and safety glasses when handling. Prepare the solution in a safety biohazard hood.

Tamoxifen and vehicle solution, 1 mg/ml Dissolve 10 mg of tamoxifen (Sigma Aldrich, cat. no. T-5648) in 1 ml ethanol by vigorous vortexing for 2 min. Add 9 ml sunflower oil and vortex until a homogeneous solution is obtained. For the vehicle solution, add 9 ml sunflower oil to 1 ml ethanol and process similarly to the tamoxifen solution. Divide into 1-ml aliquots. Store aliquots for several months at −20◦ C. CAUTION: When handling, wear gloves and safety glasses. Prepare the solution in a safety biohazard hood.

TBE buffer 90 mM Tris-base (Euromedex, cat. no. 200923-A) 90 mM boric acid (Euromedex, cat no. 50765) 2 mM EDTA (Euromedex, cat. no. EDTA23) Store up to several months at room temperature COMMENTARY Background Information

SpatioTemporally Controlled Targeted Somatic Mutations

To generate somatic mutations in mouse epidermal keratinocytes, transgenic mice expressing the bacteriophage P1 site-specific Cre recombinase under the control of the K5 or K14 promoters have been established (Tarutani et al., 1997; Vasioukhin et al., 1999; Huelsken et al., 2001; Jonkers et al., 2001; Li et al., 2001; Berton et al., 2003; Mao et al., 2003; Ramirez et al., 2004). However, as these promoters become active during fetal skin formation, the mutation of loxP-flanked alleles of a given tar-

get gene (“floxed” gene) cannot be temporally controlled. Tetracycline-dependent regulatory systems have been used to achieve temporally controlled Cre recombinase expression upon doxycycline (a tetracycline analog) treatment of transgenic mice. Mice bearing transgenes encoding tetracycline-controlled transactivators under the control of basal keratinocyteselective promoters, as well as transgenic mice in which the expression of the Cre recombinase is under the control of a minimal promoter that contains operator sites of the tet operon, are available (Diamond et al., 2000; Perl et al.,

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2002; Schonig et al., 2002). However, producing mice bearing both transgenes and the two “floxed” alleles of a given target gene requires complex breedings, and further genetic engineering is required to eliminate basal levels of Cre expression (Branda and Dymecki, 2004; Feil, 2007; Sprengel and Hasan, 2007).

Critical Parameters The Cre-ERT2 transgenic line was successfully used to efficiently ablate various target genes in epidermal keratinocytes and to generate new mouse models of human diseases (e.g., see Li et al., 2000, 2005, 2009; McLean et al., 2004; Herrmann et al., 2005; Indra et al., 2005b, 2007; Calleja et al., 2006; Nenci et al., 2006; Stratis et al., 2006; Fadloun et al., 2007; Malanchi et al., 2008; Zhang et al., 2009). The Cre-ERT2 line is preferred to transgenic lines expressing other tamoxifendependent chimeric Cre recombinases in epidermal keratinocytes [e.g., K5-Cre-ERT (Indra et al., 1999) and K14-Cre-ERtam (Vasioukhin et al., 1999)], since ∼10-fold lower tamoxifen doses are required to efficiently induce Cremediated recombination. However, before generating large cohorts of mice for extensive phenotyping, it is recommended to verify floxed DNA excision specificity and its efficiency in epidermal keratinocytes. As epidermal suprabasal keratinocytes are renewed within about 2 weeks, mice should be sacrificed ∼2 weeks after the last tamoxifen administration, and excision analyzed in skin samples and other organs (e.g., salivary gland, tongue, esophagus, and stomach, in which the K14 promoter is also active). To ensure that excision of the floxed target gene is efficiently induced in epidermal stem cells, the excision rate should also be analyzed in skin biopsies several months after tamoxifen administration. To evaluate the efficiency of floxed DNA excision in epidermal keratinocytes, the epidermis can be separated from the dermis after dispase treatment of the skin. Even though genomic DNA PCR amplification from crude lysates of pup tail biopsies is very robust with the described procedure, it might be less efficient from other tissue lysates. Thus, genomic DNA extraction of proteinase K–treated samples with phenol is preferred for PCR-mediated DNA amplification in different tissues. Even though intraperitoneal injection of tamoxifen at 0.1 mg/day for 5 days induced efficient excision of most targeted floxed genes in epidermal keratinocytes of K14-Cre-ERT2

mice, higher doses may be required for poorly accessible loci. This problem could be circumvented by administration of tamoxifen for a longer time period, rather than by increasing the daily dose. Indeed, high levels of tamoxifen induce defects in the oral cavity and high saliva production in this transgenic line that may lead to lethality (D. Metzger and P. Chambon, unpub. observ.). As doses up to 5 mg/day can be given to most mouse lines expressing Cre-ER fusion proteins, including K5-Cre-ERT and K5-Cre-ERT2 lines, without major side effects, the observed toxicity might be caused by the higher Cre-ERT2 levels expressed in K14-Cre-ERT2 mice. It remains to be determined whether these defects result from illegitimate chromosomal rearrangements at cryptic/pseudo loxP sites, as previously seen in some Cre- and Cre-ERT2 expressing transgenic lines (Schmidt et al., 2000; Higashi et al., 2009), or from squelching of ERα coregulators. Even though we never observed any major side effect in epidermal keratinocytes of tamoxifen-treated K14-CreERT2 mice, such mice should be included in studies of K14-Cre-ERT2 /GeneXL2/L2 mice to avoid misinterpretation of the experimental results. To avoid lethality that might be induced by target gene ablation in all epidermal keratinocytes and/or pleiotrophic effects resulting from target gene ablation in other epithelia in which the K14 promoter is active, target gene ablation can be induced locally in epidermal keratinocytes, by topical application of 4-hydroxytamoxifen, the active metabolite of tamoxifen. Even though this ligand induces efficient recombination in keratinocytes of the topically treated skin, the required doses could also elicit some recombination in other epithelia, as it will enter the blood circulation (D. Metzger and P. Chambon, unpub. observ.). Although it is desirable in most studies to introduce targeted somatic mutations in all epidermal keratinocytes, in some cases (e.g., induction of oncogenic mutations), it is preferable to generate sporadic mutations in the epidermis. Such mutations can be generated by lowering the doses of tamoxifen administered. The extent of target gene recombination can be evaluated by quantitative PCR or preferably by immunohistochemical analyses after administration of tamoxifen intraperitoneally, orally, or topically at various concentrations and/or at different time points, to select the most appropriate treatment. As the efficiency of Cre-mediated excision of floxed DNA is gene-specific (Vooijs et al.,

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2001; D. Metzger and P. Chambon, unpub. observ.), recombination of some target gene may occur in the absence of tamoxifen treatment. This tamoxifen-independent excision has been observed in K14-Cre-ERT2 mice for few floxed genes, such as Nemo, I kappa B kinase 2, and RARγ (Nenci et al., 2006; Stratis et al., 2006; D. Metzger and P. Chambon, unpub. observ.). For loci that are very sensitive to Cre recombinase activity, transgenic lines expressing lower levels of ligand-dependent Cre recombinases (e.g., K5-Cre-ERT or K5-Cre-ERT2 ; Indra et al., 1999) can be used to circumvent this problem.

Troubleshooting Troubleshooting advice can be found in Table 4.

Anticipated Results

SpatioTemporally Controlled Targeted Somatic Mutations

The use of the present protocol has resulted in efficient ablation of most genes that have been targeted in epidermal keratinocytes. However, for each gene it is essential to verify that no background recombination occurred before tamoxifen administration, and to determine the efficiency and selectivity of gene ablation to interpret the outcome of the experiments. This is usually easily achieved by PCR amplification of the various alleles. As shown in Figure 5, RXRα ablation in K14-Cre-ERT2 /RXRαL2/L2 mice is selective and efficient in epidermal keratinocytes, as well as fully tamoxifen-dependent. Indeed, 2 weeks after tamoxifen treatment, most, if not all, RXRα L2 alleles were converted into RXRα L- alleles in the epidermis of K14-Cre-ERT2 /RXRαL2/L2 mice (called RXRαep-/-(i) mice), whereas no such alleles were detected in their dermal cells (compare Fig. 5 lanes 9 and 10). Moreover, no RXRα disruption occurred in vehicle-treated K14Cre-ERT2 /RXRαL2/L2 mice (Fig. 5, lanes 7 and 8). It is also highly recommended to verify the absence of the target gene product in epidermal keratinocytes at various time points after tamoxifen administration, by immunohistochemical or immunoblot analyses. If there is evidence of a truncated protein produced from the recombined allele, further characterization should be performed by analysis of the transcripts (e.g., by quantitative RT-PCR). If, as expected, the target gene is efficiently ablated in basal keratinocytes, including epidermal stem cells, the genetic modification will be present in epidermal keratinocytes for months.

Our phenotypic analyses of RXRαep-/-(i) mice revealed that selective disruption of RXRα in epidermal keratinocytes during adulthood results in alopecia, a hyperproliferation of interfollicular epidermis; abnormal differentiation of keratinocytes; and skin inflammatory reactions (Li et al., 2000). Temporal control of RXRα ablation in such mice also offered the possibility to study the role of this receptor in epidermal tumorigenesis induced by a single topical application of 7,12-dimethyl-benz[a]antracene (DMBA), followed by repeated topical application of a phorbol ester (e.g., 12-Otetradecanoylphorbol-13 acetate, TPA). The number and the size of epidermal tumors was increased about 2-fold in mice in which RXRα was ablated before or after tumor initiation, and such tumors had a much higher risk of malignant conversion than those of control mice (Indra et al., 2007). Thus, keratinocytic RXRα acts as a tumor modifier of both promotion and malignant progression of epidermal tumors induced by topical DMBA/TPA treatment. Interestingly, K14-Cre-ERT2 mice also allows us to efficiently ablate compound target genes in epidermal keratinocytes, as illustrated in our study of RXRαβep-/-(i) mice, in which both RXRα and RXRβ are efficiently and selectively ablated in epidermal keratinocytes by tamoxifen administration to adult K14-CreERT2(tg/0) /RXRαL2/L2 /RXRβL2/L2 mice (Li et al., 2005). These mice develop a phenotype similar to that of human atopic dermatitis, characterized by eczematous lesions with xerosis and pruritus, associated with a skin inflammatory infiltrate mostly composed of CD4+ T helper (Th) 2 cells, dendritic cells, eosinophils, and mast cells, and systemic abnormalities including elevated serum IgE and IgG levels, as well as blood and tissue eosinophilia. These studies show that RXRs are involved in the control of cutaneous inflammation, and strongly suggest that keratinocytes could play a key role in the pathogenesis of atopic dermatitis. Other examples of K14-Cre-ERT2 mediated gene ablation in epidermal keratinocytes of adult mice are reported in several references (McLean et al., 2004; Herrmann et al., 2005; Indra et al., 2005b, 2007; Calleja et al., 2006; Nenci et al., 2006; Stratis et al., 2006; Fadloun et al., 2007; Malanchi et al., 2008; Li et al., 2009; Zhang et al., 2009). Note that the K14-Cre-ERT2 transgenic line also offers the possibility to induce temporally

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Table 4 List of Common Problems and Solutions

Step Problem

Possible cause

Solution

23

The characteristics of the used K14-Cre-ERT2 transgenic line changed upon breeding [can be evaluated on previously used reporter mice; e.g., RosaR26 (Soriano, 1999)]; see reference Metzger et al. (2003).

Start a new colony of K14-Cre-ERT2 transgenic mice from frozen embryos.

Recombination in the absence of tamoxifen treatment [has been observed in a few cases; e.g., Nemo, I kappa B kinase 2 (Nenci et al., 2006; Stratis et al., 2006) and RARγ (D. Metzger and P. Chambon, unpub. observ.)]

It is gene-specific within a given The target locus is efficiently cell type (D. Metzger and P. recombined with low amounts Chambon, unpub. observ.). of Cre recombinase (minute amounts of active Cre recombinase might be produced from cleaved Cre-ERT2 fusion protein).

Tamoxifen-independent recombination of some floxed target genes can be circumvented by using other transgenic lines expressing lower levels ligand-dependent Cre recombinases in basal keratinocytes (e.g., K5-Cre-ERT or K5-Cre-ERT2 ) (Indra et al., 1999).

Low level of excision after tamoxifen-treatment

Administer tamoxifen for a longer time period (e.g., after 5 days of daily i.p. administration, give tamoxifen every second day for 2 weeks).

The target locus is poorly accessible to Cre recombinase.

It is not recommended to inject higher daily doses of tamoxifen to K14-Cre-ERT2 mice, even though doses up to 5 mg/day can be given to most Cre-ER transgenic lines. Indeed, for unclear reasons, high tamoxifen concentrations induce defects in the oral cavity and high saliva production in K14-Cre-ERT2 mice, and might lead to death of mice. 24

Ablation of the target gene in all Severe epidermal defects (e.g., loss epidermal keratinocytes is lethal of skin barrier) resulting from lack of target protein expression in epidermal keratinocyte of the whole skin.

Perform topical 4-hydroxytamoxifen applications. This ligand is preferred to tamoxifen, as the latter has to be converted into 4-hydroxytamoxifen in the liver to efficiently induce Cre-ERT2 recombinase activity. Adjust tamoxifen levels to induce recombination in a lower number of keratinocytes. Induce targeted ablation in cultured keratinocytes

Ablation of the target gene induces pleiotropic effects

Target gene ablation occurs in other Perform topical 4-hydroxytamoxifen tissues, in which K14 promoter is applications. active (Vassar et al., 1989). Decrease tamoxifen dosing to make recombination more keratinocyte specific.

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Figure 5 Analysis of K14-Cre-ERT2 -mediated recombination of “floxed” RXRα in genomic DNA of adult skin. L2 and L- RXRα alleles were identified by PCR analysis of genomic DNA extracted from epidermis “E” or dermis “D” isolated from the tail two weeks after administration of either tamoxifen (tam) or vehicle (veh) to K14Cre-ER T2(tg/0) /RXRα L2 /L2 and K14-Cre-ER T2(0/0) /RXRαL2/L2 mice (lanes 5 to 10), as indicated. Lanes 2 and 3, amplification products from RXRαL2/L2 and RXRαL-/+ mice, respectively. The PCR products corresponding to the RXRα WT (158 bp) and RXRα L2 (199 bp) alleles were amplified with primers BAA239 (P1) and BAA982 (P2) (top panel); the PCR products corresponding to RXRα L- alleles (133 bp) were amplified with primers BAA239 (P1) and BAA983 (P3) (bottom panel). Lane 1, DNA ladder (L). The size of the DNA segments is given in base pairs.

controlled targeted mutations in keratinocytes during fetal epidermis formation, by tamoxifen administration to gestating females (Indra et al., 2005a).

Time Considerations

SpatioTemporally Controlled Targeted Somatic Mutations

Steps 1 to 13: Generation of premutant K14-Cre-ERT2(tg/0) /GeneXL2/L2 mice for temporally controlled GeneX ablation in epidermal keratinocytes and control mice takes 6 months. Steps 14 to 15: Induction of loxPflanked DNA excision takes 5 days. Efficient floxed DNA excision in basal epidermal keratinocytes is usually obtained within 5 days of tamoxifen administration. In some cases, 1 to 2 days are sufficient. Steps 16 to 24: Characterization of floxed DNA excision takes 3 weeks. As renewal of all suprabasal keratinocytes takes ∼2 weeks, we recommend determining the efficiency of floxed DNA excision in epidermal keratinocytes at least 2 weeks after the last tamoxifen injection. PCR analysis of the target gene alleles requires ∼2 days.

Acknowledgements The authors would like to thank present and past members of the laboratory who have contributed to developing the foregoing protocol. This work was supported by funds from the Centre National de la Recherche Scientifique, the Institut National de la Sant´e et de la Recherche M´edicale, the Minist`ere de l’Enseignement Sup´erieur et de la Recherche, the Coll`ege de France, the Association pour la Recherche sur le Cancer, and the Fondation pour la Recherche M´edicale.

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Calleja, C., Messaddeq, N., Chapellier, B., Yang, H., Krezel, W., Li, M., Metzger, D., Mascrez, B., Ohta, K., Kagechika, H., Endo, Y., Mark, M., Ghyselinck, N.B., and Chambon, P. 2006. Genetic and pharmacological evidence that a retinoic acid cannot be the RXR-activating ligand in mouse epidermis keratinocytes. Genes Dev. 20:1525-1538. Diamond, I., Owolabi, T., Marco, M., Lam, C., and Glick, A. 2000. Conditional gene expression in the epidermis of transgenic mice using the tetracycline-regulated transactivators tTA and rTA linked to the keratin 5 promoter. J Invest. Dermatol. 115:788-794. el Marjou, F., Janssen, K.P., Chang, B.H., Li, M., Hindie, V., Chan, L., Louvard, D., Chambon, P., Metzger, D., and Robine, S. 2004. Tissuespecific and inducible Cre-mediated recombination in the gut epithelium. Genesis 39:186-193. Fadloun, A., Kobi, D., Pointud, J.C., Indra, A.K., Teletin, M., Bole-Feysot, C., Testoni, B., Mantovani, R., Metzger, D., Mengus, G., and Davidson, I. 2007. The TFIID subunit TAF4 regulates keratinocyte proliferation and has cellautonomous and non-cell-autonomous tumour suppressor activity in mouse epidermis. Development 134:2947-2958. Feil, R. 2007. Conditional somatic mutagenesis in the mouse using site-specific recombinases. Handb. Exp. Pharmacol. 178:3-28. Feil, R., Wagner, J., Metzger, D., and Chambon, P. 1997. Regulation of Cre recombinase activity by mutated estrogen receptor ligandbinding domains. Biochem. Biophys. Res. Commun. 237:752-757. Fuchs, E. 2007. Scratching the surface of skin development. Nature 445:834-842. Herrmann, T., Grone, H.J., Langbein, L., Kaiser, I., Gosch, I., Bennemann, U., Metzger, D., Chambon, P., Stewart, A.F., and Stremmel, W. 2005. Disturbed epidermal structure in mice with temporally controlled fatp4 deficiency. J. Invest. Dermatol. 125:1228-1235. Higashi, A.Y., Ikawa, T., Muramatsu, M., Economides, A.N., Niwa, A., Okuda, T., Murphy, A.J., Rojas, J., Heike, T., Nakahata, T., Kawamoto, H., Kita, T., and Yanagita, M. 2009. Direct hematological toxicity and illegitimate chromosomal recombination caused by the systemic activation of CreERT2. J. Immunol. 182:5633-5640. Huelsken, J., Vogel, R., Erdmann, B., Cotsarelis, G., and Birchmeier, W. 2001. Beta-Catenin controls hair follicle morphogenesis and stem cell differentiation in the skin. Cell 105:533-545.

ER(T2) recombinases. Nucleic Acids Res. 27:4324-4327. Indra, A.K., Li, M., Brocard, J., Warot, X., Bornert, J.M., Gerard, C., Messaddeq, N., Chambon, P., and Metzger, D. 2000. Targeted somatic mutagenesis in mouse epidermis. Horm. Res. 54:296-300. Indra, A.K., Dupe, V., Bornert, J.M., Messaddeq, N., Yaniv, M., Mark, M., Chambon, P., and Metzger, D. 2005a. Temporally controlled targeted somatic mutagenesis in embryonic surface ectoderm and fetal epidermal keratinocytes unveils two distinct developmental functions of BRG1 in limb morphogenesis and skin barrier formation. Development 132:4533-4544. Indra, A.K., Mohan, W.S., Frontini, M., Scheer, E., Messaddeq, N., Metzger, D., and Tora, L. 2005b. TAF10 is required for the establishment of skin barrier function in foetal, but not in adult mouse epidermis. Dev. Biol. 285:28-37. Indra, A.K., Castaneda, E., Antal, M.C., Jiang, M., Messaddeq, N., Meng, X., Loehr, C.V., Gariglio, P., Kato, S., Wahli, W., Desvergne, B., Metzger, D., and Chambon, P. 2007. Malignant transformation of DMBA/TPA-induced papillomas and nevi in the skin of mice selectively lacking retinoid-X-receptor alpha in epidermal keratinocytes. J. Invest. Dermatol. 127:1250-1260. Jonkers, J. and Berns, A. 2002. Conditional mouse models of sporadic cancer. Nature Rev. Cancer 2:251-265. Jonkers, J., Meuwissen, R., van der Gulden, H., Peterse, H., van der Valk, M., and Berns, A. 2001. Synergistic tumor suppressor activity of BRCA2 and p53 in a conditional mouse model for breast cancer. Nat. Genet. 29:418-425. Li, M., Indra, A.K., Warot, X., Brocard, J., Messaddeq, N., Kato, S., Metzger, D., and Chambon, P. 2000. Skin abnormalities generated by temporally controlled RXRalpha mutations in mouse epidermis. Nature 407:633-636. Li, M., Chiba, H., Warot, X., Messaddeq, N., Gerard, C., Chambon, P., and Metzger, D. 2001. RXR-alpha ablation in skin keratinocytes results in alopecia and epidermal alterations. Development 128:675-688. Li, M., Messaddeq, N., Teletin, M., Pasquali, J.L., Metzger, D., and Chambon, P. 2005. Retinoid X receptor ablation in adult mouse keratinocytes generates an atopic dermatitis triggered by thymic stromal lymphopoietin. Proc. Natl. Acad. Sci. U.S.A. 102:14795-14800.

Imayoshi, I., Ohtsuka, T., Metzger, D., Chambon, P., and Kageyama, R. 2006. Temporal regulation of Cre recombinase activity in neural stem cells. Genesis 44:233-238.

Li, M., Hener, P., Zhang, Z., Ganti, K.P., Metzger, D., and Chambon, P. 2009. Induction of thymic stromal lymphopoietin expression in keratinocytes is necessary for generating an atopic dermatitis upon application of the active vitamin D3 analogue MC903 on mouse skin. J. Invest. Dermatol. 129:498-502.

Indra, A.K., Warot, X., Brocard, J., Bornert, J.M., Xiao, J.H., Chambon, P., and Metzger, D. 1999. Temporally-controlled site-specific mutagenesis in the basal layer of the epidermis: Comparison of the recombinase activity of the tamoxifen-inducible Cre-ER(T) and Cre-

Malanchi, I., Peinado, H., Kassen, D., Hussenet, T., Metzger, D., Chambon, P., Huber, M., Hohl, D., Cano, A., Birchmeier, W., and Huelsken, J. 2008. Cutaneous cancer stem cell maintenance is dependent on beta-catenin signalling. Nature 452:650-653.

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Mao, C.M., Yang, X., Cheng, X., Lu, Y.X., Zhou, J., and Huang, C.F. 2003. Establishment of keratinocyte-specific Cre recombinase transgenic mice. Yi Chuan Xue Bao 30:407-413. McLean, G.W., Komiyama, N.H., Serrels, B., Asano, H., Reynolds, L., Conti, F., HodivalaDilke, K., Metzger, D., Chambon, P., Grant, S.G., and Frame, M.C. 2004. Specific deletion of focal adhesion kinase suppresses tumor formation and blocks malignant progression. Genes Dev. 18:2998-3003. Metzger, D. and Chambon, P. 2001. Site- and timespecific gene targeting in the mouse. Methods 24:71-80. Metzger, D., Indra, A.K., Li, M., Chapellier, B., Calleja, C., Ghyselinck, N.B., and Chambon, P. 2003. Targeted conditional somatic mutagenesis in the mouse: temporally-controlled knock out of retinoid receptors in epidermal keratinocytes. Methods Enzymol. 364:379-408. Nenci, A., Huth, M., Funteh, A., Schmidt-Supprian, M., Bloch, W., Metzger, D., Chambon, P., Rajewsky, K., Krieg, T., Haase, I., and Pasparakis, M. 2006. Skin lesion development in a mouse model of incontinentia pigmenti is triggered by NEMO deficiency in epidermal keratinocytes and requires TNF signaling. Hum. Mol. Genet. 15:531-542. Park, E.J., Sun, X., Nichol, P., Saijoh, Y., Martin, J.F., and Moon, A.M. 2008. System for tamoxifen-inducible expression of Crerecombinase from the Foxa2 locus in mice. Dev. Dyn. 237:447-453. Perl, A.K., Wert, S.E., Nagy, A., Lobe, C.G., and Whitsett, J.A. 2002. Early restriction of peripheral and proximal cell lineages during formation of the lung. Proc. Natl. Acad. Sci. U.S.A. 99:10482-10487. Rajewsky, K., Gu, H., Kuhn, R., Betz, U. A., Muller, W., Roes, J., and Schwenk, F. 1996. Conditional gene targeting. J. Clin. Invest. 98:600-603. Ramirez, A., Page, A., Gandarillas, A., Zanet, J., Pibre, S., Vidal, M., Tusell, L., Genesca, A., Whitaker, D.A., Melton, D.W., and Jorcano, J.L. 2004. A keratin K5Cre transgenic line appropriate for tissue-specific or generalized Cremediated recombination. Genesis 39:52-57.

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Ratnacaram, C.K., Teletin, M., Jiang, M., Meng, X., Chambon, P., and Metzger, D. 2008. Temporally controlled ablation of PTEN in adult mouse prostate epithelium generates a model of invasive prostatic adenocarcinoma. Proc. Natl. Acad. Sci. U.S.A. 105:2521-2526. Ruzankina, Y., Pinzon-Guzman, C., Asare, A., Ong, T., Pontano, L., Cotsarelis, G., Zediak, V.P., Velez, M., Bhandoola, A., and Brown, E. 2007. Deletion of the developmentally essential gene ATR in adult mice leads to age-related phenotypes and stem cell loss. Cell Stem Cell 1:113126. Schmidt, E.E., Taylor, D.S., Prigge, J.R., Barnett, S., and Capecchi, M.R. 2000. Illegitimate Cre-dependent chromosome rearrangements in

transgenic mouse spermatids. Proc. Natl. Acad. Sci. U.S.A. 97:13702-13707. Schonig, K., Schwenk, F., Rajewsky, K., and Bujard, H. 2002. Stringent doxycycline dependent control of CRE recombinase in vivo. Nucleic Acids Res. 30:e134. Schuler, M., Dierich, A., Chambon, P., and Metzger, D. 2004. Efficient temporally controlled targeted somatic mutagenesis in hepatocytes of the mouse. Genesis 39:167-172. Schuler, M., Ali, F., Metzger, E., Chambon, P., and Metzger, D. 2005. Temporally controlled targeted somatic mutagenesis in skeletal muscles of the mouse. Genesis 41:165-170. Slezak, M., Goritz, C., Niemiec, A., Frisen, J., Chambon, P., Metzger, D., and Pfrieger, F.W. 2007. Transgenic mice for conditional gene manipulation in astroglial cells. Glia 55:15651576. Soriano, P. 1999. Generalized lacZ expression with the ROSA26 Cre reporter strain. Nat. Genet. 21:70-71. Sprengel, R. and Hasan, M.T. 2007. Tetracyclinecontrolled genetic switches. Handb. Exp. Pharmacol. 178:49-72. Stratis, A., Pasparakis, M., Markur, D., Knaup, R., Pofahl, R., Metzger, D., Chambon, P., Krieg, T., and Haase, I. 2006. Localized inflammatory skin disease following inducible ablation of I kappa B kinase 2 in murine epidermis. J. Invest. Dermatol. 126:614-620. Tarutani, M., Itami, S., Okabe, M., Ikawa, M., Tezuka, T., Yoshikawa, K., Kinoshita, T., and Takeda, J. 1997. Tissue-specific knockout of the mouse Pig-a gene reveals important roles for GPI-anchored proteins in skin development. Proc. Natl. Acad. Sci. U.S.A. 94:7400-7405. Vasioukhin, V., Degenstein, L., Wise, B., and Fuchs, E. 1999. The magical touch: Genome targeting in epidermal stem cells induced by tamoxifen application to mouse skin. Proc. Natl. Acad. Sci. U.S.A. 96:8551-8556. Vassar, R., Rosenberg, M., Ross, S., Tyner, A., and Fuchs, E. 1989. Tissue-specific and differentiation-specific expression of a human K14 keratin gene in transgenic mice. Proc. Natl. Acad. Sci. U.S.A. 86:1563-1567. Vooijs, M., Jonkers, J., and Berns, A. 2001. A highly efficient ligand-regulated Cre recombinase mouse line shows that LoxP recombination is position dependent. EMBO Rep. 2:292297. Wendling, O., Bornert, J.M., Chambon, P., and Metzger, D. 2009. Efficient temporally-controlled targeted mutagenesis in smooth muscle cells of the adult mouse. Genesis 47:14-18. Zhang, Z., Hener, P., Frossard, N., Kato, S., Metzger, D., Li, M., and Chambon, P. 2009. Thymic stromal lymphopoietin overproduced by keratinocytes in mouse skin aggravates experimental asthma. Proc. Natl. Acad. Sci. U.S.A. 106:1536-1541.

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Echocardiography in Mice Shumin Gao,1 David Ho,1 Dorothy E. Vatner,1 and Stephen F. Vatner1 1

University of Medicine & Dentistry of New Jersey, New Jersey Medical School, Newark, New Jersey

ABSTRACT Murine models have been utilized with increasing frequency mainly due to availability of genetically engineered models. With advancement in high spatial and temporal resolution, echocardiography is used extensively for the evaluation of cardiovascular function in murine models of cardiovascular disease. This review summarizes the general applications and methods involved in echocardiography used to study mouse models for cardiovascular research, based on 20 years of experience in the authors’ laboratory. The goal of this article is to provide a practical guide to the use of echo techniques in mice to evaluate cardiac systolic and diastolic function. Curr. Protoc. Mouse Biol. C 2011 by John Wiley & Sons, Inc. 1:71-83  Keywords: echocardiography r systolic function r diastolic function r mouse

INTRODUCTION Murine models for cardiovascular disease have been utilized with increasing frequency mainly because of the expanding availability of genetic models. Echocardiography (echo) is a useful non-invasive method to visualize the cardiovascular structures and evaluate cardiac function in mice. Improved echocardiography instrumentation enhances the spatial and temporal resolution for imaging, resulting in more accurate assessment of left ventricular systolic, diastolic, regional, and vascular function. Echocardiography is an extremely versatile tool for cardiovascular research allowing the evaluation of left ventricular (LV) systolic function and diastolic function in mouse cardiomyopathy models (Iwase et al., 1996, 1997; Asai et al., 1999), the myocardial ischemia model (Odashima et al., 2007), and chronic pressure overload induced by transverse aortic constriction (TAC; Depre et al., 2006; Gelpi et al., 2009; Guellich et al., 2010). Also, coronary reserve in mice can be measured by echocardiography (Gao et al., 2008a,b). The applications and advances of echocardiography in mice have been recently summarized (Rottman et al., 2007; Stypmann, 2007; Scherrer-Crosbie and Thibault, 2008). In this article, focus will be on how to apply echocardiography for research in normal mice, genetically altered mice, and models of cardiovascular disease using examples from the general echocardiography protocols used in the authors’ laboratory.

ECHOCARDIOGRAPHY IN CONSCIOUS MICE Anesthesia depresses contraction, heart rate, and autonomic reflex control (Vatner and Braunwald, 1975; Vatner et al., 2002). Therefore, it is preferable to perform cardiovascular experiments in the conscious state rather than under anesthesia. This is applicable to experiments in mice as well as large animal models. However, there are also serious limitations to performing echocardiography in conscious mice; most notably, excitement in the animals can result in enhanced sympathetic tone and heart rate. Accordingly, training mice for studies in the conscious state is essential. 1. Before echocardiography can be performed effectively in conscious mice, train mice for two to three sessions over a 3-day period by holding the nape of the neck with the first two fingers and tail held tightly with the last two fingers, and placing the ultrasound Current Protocols in Mouse Biology 1: 71-83, March 2011 Published online March 2011 in Wiley Online Library (wileyonlinelibrary.com). DOI: 10.1002/9780470942390.mo100130 C 2011 John Wiley & Sons, Inc. Copyright 

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transducer on the chest for probe contact training. After a few days of training, the mouse will remain calm in this position. Just prior to the acquisition of echo images, remove the chest hair by shaving or by applying hair removal cream. Remove as small an area of hair as possible to prevent excessive heat loss. Further details of echocardiography methods are described below in the section Echo Machines and Transducers. 2. For echocardiography, the mouse is picked up in the palm of one hand. Pre-warmed echo transmission gel is applied to the hairless chest. While holding the mouse with the back of the mouse towards the palm, the hand is turned so the chest of the mouse faces the floor. The transducer is applied from under the mouse to avoid reflex effects induced by the pressure of the transducer on the chest. 3. Parasternal long-axis view, short-axis view at the papillary muscle level, and 2-D guided M-mode images are recorded as described in the mouse echo protocol detailed below. Baseline values for heart rate (HR) and fractional shortening (FS) in conscious and anesthetized mice are summarized and displayed in Table 1, showing much higher values in conscious mice than in those under anesthesia. The heart rate in conscious mice is usually 600 to 700 beats/min (Yang et al., 1999). The disadvantages of conscious echocardiography are: (1) mice must be trained for a few days to minimize the excitement induced by mice manipulation (Yang et al., 1999; Rottman et al., 2007); (2) a second person is needed to operate the ultrasound machine; and (3) the faster heart rates in conscious mice complicate Doppler recording, e.g., the Doppler waveforms fuse together because of the short diastolic time.

ANESTHESIA FOR MOUSE ECHOCARDIOGRAPHY Because of the limitations of echocardiography in conscious mice, anesthesia is frequently used in murine echocardiography. Various regimens of anesthesia, e.g., a continuously delivered gas inhalation agent isoflurane (1% to 3%) (Roth et al., 2002; Hartley et al., 2008; Wikstrom et al., 2008), avertin (tribromoethanol, 250 to 400 mg/kg i.p.) (Gardin et al., 1995; Hart et al., 2001; Roth et al., 2002; Schmidt et al., 2002; Tan et al., 2003; Luo et al., 2007), pentobarbital (50 mg/kg i.p.) (Yang et al., 1999; Rottman et al., 2003; Tan et al., 2003), ketamine (80 to 150 mg/kg i.p.) mixed with xylazine (5 to 20 mg/kg i.p.) (Tanaka et al., 1996; Yang et al., 1999; Hart et al., 2001; Tan et al., 2003; Schaefer et al., 2005), are employed in different laboratories. According to Roth et al. (2002), both isoflurane and avertin cause less depression of cardiac function and heart rate, are easy to administer, provide reproducible results, and are rapid in onset and recovery. It has been found that 2.5% avertin, 0.012 ml/g body weight (300 mg/kg) i.p. allows for the most predictable and reproducible level of cardiac suppression and maintenance of heart rate, leading to consistency in the quality of echo measurements. Using this anesthesia, the heart rates of normal mice are generally 400 to 500 beats/min, and typically LV FS is ∼35%. These values are comparable to the measurements with 2% isoflurane– anesthetized mice (Stypmann, 2007). The effects of different anesthesia on FS and HR are shown in Table 1. Conversely, a ketamine/xylazine mixture resulted in the greatest depression of heart rate. One potential adverse effect of too low a heart rate induced by anesthesia is cardiac dilatation with consequent functional valvular regurgitation. This has been observed during rat echos (Droogmans, 2008), and most likely will also occur in mice. Echocardiography in Mice

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Table 1 Fractional Shortening (FS) and Heart Rate (HR) in Conscious and Anesthetized Micea

Type of anesthesia None (conscious)

Avertin

Pentobarbital

Ketamine + xylazine

Isoflurane

FS (%)

59±5

35±0.4

33±2

35±1

39±1

HR (bpm)

683±63

411±17

377±11

293±19

457±17

Dose

None

300 mg/kg i.p.

50 mg/kg i.p.

150+15 mg/kg i.p.

2% inhalation

Data resource Rottman et al. (2003) Yan et al. (2007) Yang et al. (1999) Yang et al. (1999)

Stypmann (2007)

a All values are mean ± SE. The heart rate and systolic function in conscious mice were much higher than those seen in anesthetized mice. FS,

fractional shortening; HR, heart rate; i.p., intraperitoneal injection.

ECHO MACHINES AND TRANSDUCERS Echo transducers with frequencies >10 MHz are generally selected for mouse echocardiography. This is necessary because of the small size of the heart and its rapid rate of contraction. In the authors’ core echo laboratory, there is one Siemens Sequoia C256 with a 13-MHz linear transducer, one GE Vivid7 with an i13L probe (14 MHz), and one VisualSonics Vevo770 with 30- and 40-MHz probes for mouse cardiac and vascular examination.

Mouse echo protocol 1. The mouse is injected intraperitoneally with 2.5% avertin, 0.012 ml/g body weight (300 mg/kg). Heart rates are monitored and generally maintained at 400 to 500 beats/min. 2. When using the Siemens Sequoia C256 or GE Vivid7, the chest hair is shaved and EKG needle leads are connected to the limbs for electrocardiogram gating. The mouse is then placed on a warm pad to maintain the body temperature ∼37◦ C. A rectal thermometer is inserted for monitoring the body temperature. 3. Warmed echo gel is placed on the shaved chest. The mouse heart is imaged with a 13-MHz linear transducer (Siemens Sequoia C256) or 14-MHz probe (GE Vivid7) while the mouse lies on the warm pad at a shallow left-side position. 4. When using the VisualSonics Vevo 770, due to the much higher probe frequency and interference from remaining hair, a depilatory lotion is applied to the chest to help facilitate complete removal of hair. The platform temperature of the equipment is set at 40◦ to 42◦ C, which is higher than optimal animal core temperature to help maintain the mouse core temperature at 37◦ C. The mouse is placed onto the warm plate in the supine position and the limbs are taped onto the metal EKG leads. For cardiac imaging, the 30-MHz transducer is used, while the 40-MHz transducer is utilized for vascular imaging. 5. By placing the transducer along the long-axis of LV, and directing it to the right side of the neck of the mouse, two-dimensional (2-D) LV long-axis is obtained. Then the transducer is rotated clockwise by 90◦ , and the LV short-axis view is visualized. The diagrams showing the positions and directions of the transducer for basic mouse echo views are demonstrated in Figure 1. 2-D-guided LV M-mode at the papillary muscle level is recorded from either the short-axis view and/or the long-axis view. Transmitral inflow Doppler spectra are recorded in an apical four-chamber view by placing the sample volume at the tip of the mitral valves. Angle correction can be used for accurate flow velocity measurements. Doppler waveforms from other regions of the heart can be recorded as needed.

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A

AO LV

LA

B

LV

Figure 1 Diagrams for basic mouse echocardiography views. A shows the position and direction (small arrow) of probe (upper left) for LV long-axis view (upper right). B demonstrates the position and direction (small arrow) of probe (lower left) for LV short-axis view (lower right). LA, left atrium; AO, aorta.

6. After the scanning is finished, the residual echo gel is removed, and the mouse is returned to the cage for recovery. 7. Echo images are downloaded and analyzed offline using Scion images software or an echo work station. At least three beats need to be measured and averaged for the interpretation of any given measurement. From the authors’ experience, at least five mice are normally needed per experimental group to show statistically significant relevance.

CONSIDERATIONS IN MURINE ECHOCARDIOGRAPHY To obtain consistent, reproducible echocardiographic data, in addition to obtaining good images, the condition of the animals must be controlled. Therefore, adhere to the following parameters during echo scanning. 1. The mouse body temperature should be carefully monitored and maintained at 37◦ C during the entire procedure. 2. Since cardiac function is closely related to heart rate, the heart rate should be controlled at a similar level within each strain of mice. Therefore, the choice of anesthetic agent, dose, and dosing interval should be carefully reproduced and considered. From the authors’ experience, the variation of HR within 100 bpm for a strain/set of experiments should be acceptable. Echocardiography in Mice

3. Echo measurement time should be similar after anesthesia to minimize the effects of changes in anesthetic levels with time on echocardiography parameters.

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ECHO MEASUREMENTS LV systolic function LV interventricular septal thicknesses (IVS), LV internal dimensions (LVID), and posterior wall thicknesses (PW) at diastole and systole (IVSd, LVIDd, PWd, and IVSs, LVIDs, PWs, respectively) are measured from M-mode images at the level of the papillary muscles. An example of LV M-mode in mice is displayed in Figure 2A. LV ejection fraction (EF), LV fractional shortening (FS), and LV posterior wall thickening (PWT) are calculated by using the following formulas (Gardin et al., 1995; Tanaka et al., 1996; Syed et al., 2005; Tsujita et al., 2005): EF (%) = 100 × [(LVIDd3 – LVIDs)3 )/LVIDd3 ] FS (%) = 100 × [(LVIDd – LVIDs)/LVIDd] PWT (%) = 100 × [(PWs – PWd)/PWd] LV ejection fraction (EF) and LV fractional shortening (FS) are measured for evaluation of LV global systolic function. When the LV contracts without regional wall motion

A

B

Sa

IVSd

IVSs

LVIDd

LVIDs

PWd

C

PWs

Ea ET

Aa

IVCT IVRT

LV fractional shortening (FS%)

Figure 2 Images of echocardiographic measurements in mice. (A) LV M-mode, allows for assessment of LV systolic function. IVSd, LVIDd, PWd, and IVSs, LVIDs, PWs are LV interventricular septum thicknesses, LV internal dimensions and LV posterior wall thicknesses at diastole and systole, respectively. (B) Doppler of transmitral inflow most often used for evaluation of LV diastolic function. E and A are peak velocities at early and late filling, respectively. IVRT and IVCT are isovolumetric relaxation and contraction time. ET is LV ejection time. (C) Tissue Doppler waveform obtained in LV posterior wall, used for assessing regional wall motion abnormality. Ea and Aa were two waveforms at early and late diastolic phases. Sa is the peak wall motion velocity in systole.

40

FVB C57BL/6J

35

*

30 25

*

*

20

*

15 Baseline

1W-TAC

2W-TAC

3W-TAC

Figure 3 Comparing fractional shortening (FS) in two strains of mice (FVB, C57BL/6J) before and after 1, 2, and 3 weeks of pressure overload induced by transverse aortic constriction (TAC). FS was significantly decreased in C57BL mice (square) even 1 week after TAC. However, in FVB mice (triangle) FS was maintained at normal levels even after 2 weeks of TAC. *p <0.05 versus baseline. FS, fractional shortening.

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abnormalities, EF and FS are related. However, in ischemia or myocardial infarction models, because of the changes of LV geometry, EF calculated by the simple cubic assumption of LV volume may not be accurate and the calculated LVEF could be different from the actual LV ejection fraction. In these cases, FS is preferable to express LV global function. Serial echocardiography performed in FVB and C57BL/6J mice at baseline, 1, 2, and 3 weeks after chronic pressure overloading induced by transverse aortic constriction (TAC) is illustrated in Figure 3. The effects of this stress on the heart vary in different mouse strains. Echo techniques are useful to detect the differences. For example, echocardiography detected systolic dysfunction, i.e., reduced LVEF in C57BL/6J mice after 1 week of TAC, whereas LVEF was still maintained at baseline levels even 2 weeks after TAC in FVB mice, demonstrating a major mouse strain difference in response to chronic pressure overload. Furthermore, echo is useful for detecting and monitoring the progression of cardiac dysfunction such as in the case of cardiomyopathy; a sample of M-mode echo for cardiomyopathy is shown in Figure 4C. The progression of cardiomyopathy in transgenic mice (Tg) overexpressing beta 1–adrenergic receptors (β1 -AR), illustrated in Figure 4E, is seen as an initial increase in LV fractional shortening (LV FS) in the young Tg mice (gray bar) with a decrease in LV FS in the older Tg mice (black bar) compared to wild type (WT) mice (white bar) (Peter et al., 2007).

A

B

ES

ED

C

ED

ES

ED

E

D 50

Beta1 AR transgenic mice 60

* LV FS %

45 LV FS %

ES

40

40

*

20

35 30

0 Base

0.01

0.02

0.04

WT

Young

Old

Figure 4 These are representative images and echocardiography data displaying changes in LV fractional shortening (FS) with isoproterenol and cardiomyopathy. (A) Represents a baseline image (ES, end systole; ED, end diastole). (B) After infusion with isoproterenol 0.04 μg/kg/min, LV contraction was markedly increased. (C) In transgenic mice with cardiomyopathy a clear decrease in LV contraction is observed. (D) LV FS increases with increasing doses of isoproterenol. (E) LV FS is enhanced in young transgenic mice over-expressing β1-adrenergic receptors in the heart (β1-AR Tg, gray bar) as compared to the wild type (WT, white bar). However, as the mice develop cardiomyopathy with age (black bar), LV FS is found to be decreased. * p <0.05 versus WT (Peter et al., 2007). Reproduced from Peter et al. (2007) with permission from the American Society for Clinical Investigation. Echocardiography in Mice

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The velocity of circumferential fiber shortening (Vcf) is a pre-load-independent measurement for LV systolic function, which is calculated by Vcf = FS/ET. ET is the ejection time of the LV, which can be measured by PW Doppler. Since ET is heart rate dependent, correcting ET by dividing it by the square root of the R-R interval can make the corrected Vcf heart rate independent (Odley et al., 2004; Syed et al., 2005). This correction is relevant in both conscious and anesthetized mice. LV wall thickening is another method for the assessment of global LV systolic function in the absence of abnormal wall motion. However, when abnormal wall motion exists, the wall thickening represents only regional LV systolic function. Furthermore, LV mass (in diastole) can be obtained from M-Mode measurements by the cubed formula: LV mass = 1.05 × [(IVSd + LVIDd + PWd)3 – LVIDd3 ]

LV diastolic function Pulse-wave Doppler Transmitral inflow Doppler obtained in apical four-chamber view or LV long-axis view is used for evaluation of LV diastolic function in mice (Schmidt et al., 2002; Schaefer et al., 2003; Semeniuk et al., 2003; Du et al., 2008). A Doppler example of transmitral flow is displayed in Figure 2B. The Doppler indexes include the ratio of peak velocity of early to late filling of mitral inflow (E/A), deceleration time (DT) of early filling of mitral inflow, isovolumetric relaxation time (IVRT), and isovolumetric contraction time (IVCT). There are four basic Doppler patterns of transmitral inflow and these four patterns represent the progression from normal to severe diastolic dysfunction (Ohno et al., 1994; Du et al., 2008): (1) normal LV filling E>A; (2) abnormal LV relaxation EA; and (4) restrictive filling E>>A. Since diastolic dysfunction progresses rapidly in mice, multiple different Doppler patterns may exist in the same group of surgically modeled or genetically altered mice and this may lead to misinterpretation of the stage of diastolic dysfunction. Thus, confirming Doppler measurements by other methods such as tissue Doppler, color M-mode Doppler, or pressure measurements, is essential. The Doppler parameters at baseline and 2 weeks after TAC in FVB mice were measured; diastolic dysfunction was evident in the echocardiogram, as reflected by decreased A wave velocity, increased E/A ratio, and an increased index (IVRT + IVCT)/ET, implying increased stiffness of LV after TAC as seen in Table 2. However, systolic function in these same FVB mice was maintained 2 weeks after TAC (Figure 3). Tissue Doppler imaging Tissue Doppler imaging (TDI) is tissue motion velocity obtained from the mitral annulus or LV posterior wall from the myocardium, which normally consists of three basic waveforms: two in early and late diastole (Ea and Aa, respectively), and one in systole (Sa). Decreased Ea/Aa ratio indicates diastolic dysfunction. Importantly, these values are influenced to a lesser extent by loading conditions (Schaefer et al., 2003). A TDI example is demonstrated in Figure 2C. Color M-Mode Doppler Color M-Mode Doppler flow propagation of transmitral inflow (Vp) is obtained by placing the M-mode cursor through the center of the mitral inflow, which is guided by color Doppler. Decreased Vp implies impaired LV relaxation, as correlated to pulse wave Doppler parameters (Schmidt et al., 2002; Tsujita et al., 2005). Myocardial performance index Pulse wave Doppler or tissue Doppler–derived myocardial performance index (MPI) is a useful index for assessing cardiac systolic and diastolic function in mice. It can be calculated by using the ratio of isovolumetric contraction and relaxation time to ejection

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Table 2 LV Diastolic Function: Transmitral Doppler Parameters at Baseline and at 2 Weeks after TAC in FVB Micea

Baseline

2 weeks TAC

E wave velocity (cm/sec)

54.2±2.7

61.9±3.5

A wave velocity (cm/sec)

43.8±2.8

28.0±4.1b

E/A

1.26±0.06

2.5±0.4b

DT (msec)

18.6±1.7

21.9±2.3

IVRT (msec)

17.5±0.4

14.8±0.7b

R-R (msec)

133±3.4

136±5.8

ET of LVOT (msec)

53.3±1.0

48.9±1.3b

(IVRT + IVCT)/ET

0.43±0.03

0.59±0.05b

11

10

n

a All the values are mean ± SE. DT, deceleration time; IVRT, isovolumetric

relaxation time; R-R, interval between R waves in EKG; ET, LV ejection time; n, number of mice. b p < 0.05 versus baseline. After 2 weeks of TAC, there was clearly LV diastolic dysfunction as reflected by a decrease in A wave velocity, increase in E/A ratio as well as myocardial performance index (IVRT + IVCT)/ET.

time (IVRT + IVCT)/ET. Increased MPI indicates diastolic dysfunction. Since this index is based on the ratio of several portions within the same cardiac cycle, MPI is independent from heart rate and LV shape (Broberg et al., 2003; Schaefer et al., 2005).

LV regional function LV wall thickening As mentioned above, LV wall thickening is measured from several regions of the LV wall and is a basic index for evaluation of LV regional systolic function (Thibault et al., 2007). This can be a critical measurement in a heart with dysynchronous contraction, for example, after a myocardial infarction, where one LV wall might exhibit enhanced function, while the other wall may not contract at all, or even paradoxically. Tissue Doppler imaging and strain rate Systolic waveform (Sa) is a measurement of regional LV wall systolic motion velocity as obtained by tissue Doppler and represents regional wall contraction. Strain rate (SR) is the relative change of length of myocardial tissue over time, and as such it can be measured using TDI. TDI and SR have been demonstrated to be sensitive methods for the detection of LV regional wall contractile changes associated with aging, exercise, cardiac toxic drugs, or myocardial ischemia (Sebag et al., 2005; Thibault et al., 2007; Derumeaux et al., 2008; Jassal et al., 2009).

Echocardiography in Mice

Two-dimensional speckle tracking echocardiography Two-dimensional (2-D) speckle tracking echocardiography (STE), also known as realtime strain rate, is a novel method for the assessment of LV segmental function by tracking the speckle motion in a 2-D echocardiography imaging. Briefly, LV short axis view is acquired at a high frame rate, e.g., over 200 frames/sec, and specific software is needed to measure the radial and circumferential strain and strain rate for each segment of the LV wall. The feasibility of 2-D-STE in mice has been tested (Peng et al., 2009). Compared to the strain rate derived from TDI, which is Doppler angle dependent and can only be obtained in the anterior and posterior LV segments, 2-D speckle tracking has the ability to assess all segments in radial and circumferential strain components. However,

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due to the thin LV wall and very high heart rate in mice, the application of the (2-D) speckle tracking technique in mice needs to be improved.

VASCULAR ULTRASOUND IN MICE Coronary flow reserve in mice Coronary reserve (CR) is the ratio of maximal coronary flow under hyperemia to baseline coronary flow. Therefore, monitoring the coronary flow is essential for the measurement of CR. High-resolution echocardiography machines make it possible for the measurement of coronary reserve in mice (Wikstrom et al., 2005, 2008; Saraste et al., 2006; Hartley et al., 2008). Since CR derived from coronary flow velocity (CFVR) correlates very well with the CR derived from volumetric coronary flow (CFR) in mice, it is acceptable to simply use CFVR for determination of CR (Wikstrom et al., 2008). In the authors’ laboratory, the high-resolution ultrasound machine VisualSonics Vevo770, with a probe frequency of 30 MHz or 40 MHz, is used for this measurement. The proximal left coronary artery (LCA) is visualized in a modified parasternal LV long-axis view, and Doppler spectrum of LCA is recorded at baseline, and under hyperemic conditions induced by infusing adenosine (160 μg/kg/min) for at least 3 min. From the Doppler spectrum of the left coronary artery, mean diastolic velocity and peak diastolic velocity are measured at baseline (CFVbaseline ) and following maximal coronary vasodilation induced by adenosine infusion (CFVhyperemia ). Coronary reserve based on coronary flow velocity is calculated using the following formula: CFVR= CFVhyperemia /CFVbaseline . Simultaneously, left main coronary artery diameter (d) is measured in the modified LV short-axis view. Cross-sectional area (A) of LCA is calculated as A = π × d 2 /4. Velocity time integral (VTI) of LCA is obtained from Doppler. Blood flow of LCA (CF) = VTI × A × HR. HR is heart rate. Coronary reserve from blood flow is calculated as CFR = CFhyperemia /CFbaseline . CR measured in normal 129SVJ mice in the authors’ laboratory by maximum velocity and by volumetric blood flow are 2.28 ± 0.1 and 2.68 ± 0.15, respectively (Gao et al., 2008a,b). These are similar to those from previously reported studies (Wikstrom et al., 2005, 2008).

Other vessels in mice With the high frequency probe (30 to 40 MHz), mouse carotid arterial lumen, length, and Doppler waveform can also be studied. Williams et al. (2007) verified the feasibility by measuring the pulse wave velocity in mouse carotid artery. Using the same technique, the aortic arch and abdominal aorta can also be visualized in mice (Feintuch et al., 2007; Luo et al., 2007). In TAC mice, measuring the flow velocity through the banded site can help assess the pressure gradient between LV and aorta non-invasively. In general, the following steps can be used to scan the vessels: first, place the probe along the course of the vessel of interest to obtain the long-axis images for lumen, length, and wall thickness measurements, and then tilt the probe to direct the ultrasound beam along the direction of blood flow to record the Doppler signals.

MYOCARDIAL CONTRAST ECHOCARDIOGRAPHY Myocardial contrast echocardiography (MCE) is performed with the aid of intravenously injected contrast agents (micro-bubbles) to enhance the myocardial image for evaluation of myocardial perfusion and the perfusion defect in myocardial ischemia experiments. The feasibility of MCE in mice has been demonstrated by several groups (Mor-Avi et al., 1999; Scherrer-Crosbie et al., 1999; French et al., 2006; Kaufmann et al., 2007; Raher et al., 2007). The high resolution of the VisualSonics echocardiography machine makes

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it easier to perform this function in mice. One might predict that it will be more difficult with echo machines of lower resolution.

STRESS ECHOCARDIOGRAPHY IN MICE In mice, stress echocardiography is generally performed with administration of pharmacologic agents under anesthesia. In the authors’ echo laboratory, echocardiography is often performed for the purpose of monitoring cardiac response to sympathomimetic amines, e.g., isoproterenol or dobutamine. For example, the protocol for isoproterenol in the authors’ laboratory is as follows: 1. A jugular vein catheter is inserted in advance for drug infusion. 2. A Harvard infusion/withdrawal pump is used for drug infusion and set to deliver isoproterenol at 0.01, 0.02, and 0.04 μg/kg/min. 3. The isoproterenol solution is prepared to deliver a final concentration of 0.01 μg/kg/min using an infusion speed of 2 μl/min. When preparing the solutions, it is important to take into account the body weight for each mouse. 4. The mouse is anesthetized using 2.5% avertin, as described above, and LV 2-D and M-mode images are obtained at baseline. 5. The catheter is connected to a 100-μl syringe prefilled with the isoproterenol solution. The syringe diameter in the infusion pump is input and the infusion speed is set at 2 μl/min. The first dose is infused at 0.01 μg/kg/min for 5 min. The echo images are recorded at 5 min of infusion. 6. The next dose is switched to dose at 0.02 μg/kg/min by adjusting the infusion speed to 4 μl/min, and increased again to 0.04 μg/kg/min by increasing the infusion speed to 8 μl/min. Echo is recorded after 5 min of infusion for each of these dosages. 7. After completing all of the doses, the echo data are analyzed offline. LV M-mode images at baseline and after isoproterenol infusion are compared in Figure 4A-B. LV fractional shortening is increased with increasing isoproterenol dose in FVB mice as shown in Figure 4D.

COMMENTARY Background Information

Echocardiography in Mice

Multiple methods for cardiac imaging have been developed over the years for the visualization and assessment of cardiac function. Among these cardiac echocardiography, micro CT (Nahrendorf et al., 2007), PET scan (Kreissl et al., 2006), and contrast-enhanced cardiac MRI are included (Slawson et al., 1998; Wiesmann et al., 2001; Yang et al., 2004). However, due to the cost and frequent need for contrast material in cardiac MRI and CT and the ease of echocardiography, echo remains the most frequently used modality for the routine evaluation of cardiac function in mice. In performing echocardiography in mice, care must be taken to control the heart rate, body temperature, and the level of anesthesia. Once the animal has been properly prepared and good images are obtained, both sys-

tolic and diastolic cardiac function can be accurately measured and compared for the monitoring of cardiac pathophysiology, as well as the effectiveness of any intervention.

Acknowledgements This work has been supported in part by NIH grants HL033107, HL069020, AG027211, HL101420, HL093481, HL059139, HL095888, DK083826, and HL102472.

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Imaging Cancer in Mice by PET, CT, and Combined PET-CT Francisca Mulero,1 Luis E. Donate,1 and Manuel Serrano1 1

Spanish National Cancer Research Centre (CNIO), Madrid, Spain

ABSTRACT The possibility of imaging tumors in live mice has opened new opportunities for cancer research, particularly regarding the ability to perform longitudinal studies in combination with a therapeutic intervention. Here, we detail methods to optimize visualization of murine tumors by positron emission tomography (PET), computed tomography (CT), C 2011 by John Wiley & and combined PET-CT. Curr. Protoc. Mouse Biol. 1:85-103  Sons, Inc. Keywords: cancer r mouse models r positron electron tomography r computed tomography

INTRODUCTION Here we describe protocols for two imaging techniques, namely, positron emission tomography (PET) and computed tomography (CT), applied to cancer research in mouse models. PET detects the uptake of radiolabeled probes by tumors. Current PET technology for mice is of moderate spatial resolution (∼1 mm), but this is compensated for by its unparalleled sensitivity in detecting tumors (Wang et al., 2006). Standard PET technology currently exploits the high glucose avidity of cancer masses by the use of labeled analogs of glucose. The capacities of PET are rapidly expanding to measure other functional properties of tumors, such as cellular proliferation, hypoxia, or apoptosis (Massoud and Gambhir, 2003). On the other hand, CT allows the visualization of anatomical structures with high resolution (∼50 μm), but the ability to identify tumors depends on the differential absorption of radiation between the tumor and its surrounding tissue, and this is not always sufficient to guarantee high sensitivity. The combination of PET and CT overcomes the intrinsic limitations of each technology, combining the high sensitivity of PET with the high resolution of CT, thus offering an unprecedented ability to identify tumors, their functional status, and their dynamics (Massoud and Gambhir, 2003).

Positron emission tomography (PET) PET alone or in combination with CT (PET-CT) has become an important imaging technique for monitoring tumor dynamics in mice, particularly in response to anticancer drugs or other therapeutic interventions (Abbey et al., 2004; Dearling et al., 2004). PET devices detect high-energy gamma rays emitted from within the subject. Natural biological molecules can be labeled with a positron-emitting isotope. Positrons annihilate upon collision with nearby electrons emitting two gamma rays of high energy (511 keV) in exact opposite directions. The two gamma rays must be simultaneously detected at the crystals of the detector; this is what is called “a coincidence.” Positron-emitting isotopes frequently used include 15 O, 13 N, 11 C, and 18 F; the last is employed as a substitute for hydrogen. Other, less commonly used positron emitters are 14 O, 64 Cu, 62 Cu, 124 I, 76 Br, 82 Rb, and 68 Ga. Most of these isotopes are produced in cyclotrons (Strijckmans, 2001) and then incorporated through the appropriate chemical reactions into the desired molecule of biological interest.

Current Protocols in Mouse Biology 1: 85-103, March 2011 Published online March 2011 in Wiley Online Library (wileyonlinelibrary.com). DOI: 10.1002/9780470942390.mo100137 C 2011 John Wiley & Sons, Inc. Copyright 

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Labeled molecular probes are administered to the subject, and PET imaging monitors their distribution and concentration. The half-lives of many of the positron-emitting isotopes used for PET imaging are relatively short (e.g., 18 F has a half life of 110 min) and, therefore, the administration of the probe to the subject must take place relatively quickly (Gambhir, 2002). Commercial PET radiopharmacies are capable of ensuring the provision of commonly used PET tracers on a daily basis. The focus of this protocol will be on [18 F]FDG (fluorodeoxyglucose), which currently is the most widely used radiocompound for PET imaging. For human [18 F]FDG PET-CT studies, standard optimization protocols are well established (Schelbert, 1998). In the case of mouse PET-CT studies, previous researchers have pioneered efforts towards the optimization of mouse anesthesia and handling (Toyama et al., 2004; Fueger et al., 2006). The physiological uptake arising from organs with a high metabolic rate, mainly the brain and the myocardium, can mask the uptake of glucose by the tumors. Also, kidneys, urinary bladder, and to a lesser extent colon and gall bladder, are involved in the physiological elimination of [18 F]FDG and, therefore, may transiently produce [18 F]FDG signals. In addition, the dietary state, the ambient temperature, or the muscle activity can modify [18 F]FDG uptake by normal tissues and these, in turn, may affect tumor detection. The basal metabolic rate per unit body weight in mice is approximately 7-fold higher than that of humans and, therefore, the effect of dietary state and ambient temperature on [18 F]FDG biodistribution in mice is more pronounced than in humans (Fueger et al., 2006). [18 F]FDG uptake by muscle and brown adipose tissue (BAT) increases with the stress of the mice and with lower environmental temperature. Therefore, the stress inflicted on the mice during handling should be minimized as much as possible and temperature should be controlled. Finally, to increase the tumor/background rate, high glucose levels should be avoided because high glycemia can outcompete the [18 F]FDG probe.

Computed tomography (CT) Images in computed tomography (CT) are based on the differential absorption of X-rays by tissues of different composition, including tumors (Dilmanian et al., 1997; Dendy and Heaton, 1999). Volumetric data are acquired through a low-energy X-ray source of 30 to 50 kV, i.e., lower energy than in human CT scanners (115 to 120 kV), and a detector rotating around the animal, thus generating a three-dimensional rendering of the mouse that can be subsequently analyzed by sections along the three axes. This allows the identification of organs, anatomical structures, and tumors, as well as the acquisition of their volumetric data (Paulus et al., 2001; Berger et al., 2002; Holdsworth and Thornton, 2002). Unlike magnetic resonance imaging (MRI), CT has a relatively poor soft-tissue contrast, often making it necessary to administer an iodinated contrast agent to delineate organs or tumors. Mouse CT images are registered on high-resolution phosphor screen/CCD (coupledcharged device) detectors to optimize image quality. The completion of a scan of an entire mouse at a resolution of 200 μm takes ∼15 min. Higher-resolution (50-μm) images are achievable at longer scanning times. Three factors limit the spatial resolution of the system: the sampling rate of the pixel, the size of the X-ray source, and blurring on the phosphor screen. It is noteworthy that the radiation dose is not negligible (0.6 Gy per scan at 200-μm resolution; 5% of the lethal median dose, LD50 , for mice), which limits the repeated imaging of the same animal.

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Basic Protocol 1 describes the protocol to perform PET. Basic Protocol 2 details the protocol for CT. Finally, Basic Protocol 3 explains the combined, multimodal, PET-CT imaging of tumors.

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CAUTION: The premises hosting the equipment needed for PET and CT studies must have all the necessary permits according to the applicable regulations. All personnel (investigators and technicians) must be qualified as radioactivity operators and be in the possession of the corresponding diplomas. Radiation safety equipment needed will include syringe and vial radioactivity protectors, shielded scales, shielded waste bins, and shielded screens. NOTE: Prior to commencing with any procedures involving the use of rodents, investigators are required to be trained in the proper use and care of small experimental animals and to be in the possession of the corresponding “Ethics Committee” approvals. These protocols must conform to the applicable regulations regarding the humane use and care of laboratory animals.

IMAGING BY POSITRON EMISSION TOMOGRAPHY (PET) The imaging of tumors at inner locations is more challenging than the imaging of subcutaneous tumors. Our interest lies in studying spontaneous tumors in genetically modified mice and, thus, we have optimized the procedures outlined in the following protocol to the particulars of visualizing tumors at internal body locations, such as the lung or the pancreas. Briefly, the mice are fasted the night before, anesthetized prior to the administration of the [18 F]FDG dose, and kept under anesthesia during the whole period of probe uptake and imaging, ensuring at all times that the mice are warm. The standardization of mouse handling and of anesthesia usage is essential to ensure data reproducibility and comparability.

BASIC PROTOCOL 1

Materials Genetically modified mouse models bearing spontaneous tumors in any body location Special mouse diets as necessary Diazepam (5 mg/ml in flip-top vial; see recipe) Isoflurane Oxygen [18 F]FDG (0.01 to 0.1 μg/ mCi), delivered daily from a local cyclotron (e.g., 40 mCi of [18 F]FDG of 95% to 99% radiochemical purity in 1 ml of physiological saline solution buffered at pH 6.0, for ∼10 PET scans) Physiological saline: 0.9% (w/v) NaCl Lacryvisc Gel 10 G (3 mg/ml carbomere in benzalconium chloride, commercially available from Alcon, http://www.alcon.com) Infrared heating lamp Isoflurane/oxygen-based anesthesia system fitted with an induction chamber and inhalation masks for mice Dose calibrator (also known as activimeter): e.g., VDC-505 dose calibrator from Veenstra Instruments (http://www.dosecalibrator.com/) PET-CT imaging system: e.g., eXplore Vista PET-CT, GE Healthcare (Fig. 1A); Argus PET-CT, SEDECAL (http://www.sedecal.com/) 1-cc tuberculin syringes 30-G needles Heating pads: e.g., Gaymar Mul-T-Pads (http://www.gaymar.com/) Heating pump to maintain temperature of heating pads: e.g., Gaymar TP600 (http://www.gaymar.com/) eXplore Vista PET-CT MMWKS software for image acquisition, processing, and analysis Workstation (e.g., Dell PowerEdge) for image acquisition, processing, and analysis meeting the following specifications:

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PE1950 Xeon 5120 1.86 GHz/4 MB 1066 FSB processor PE1950 PCIX Riser (2 slots) PE1950 Bezel Assembly 2 GB FB 667 MHz Memory (2 × 1 GB dual rank DIMMs) Dell Studio XPS Desktop 435 MT PC (for 3DOSEM image reconstruction) meeting the following specifications: Processor: Intel Core i7 Quad CPU 940 4 × 2.93 GHz Memory: 6144 MB (6 × 1024) 1067 MHZ DDR3 Graphics: ATI Radeon HD 3450 256 Mb GDDR2 Fast mice 1. Fast mice overnight prior to the procedure (see exceptions below). The specific idiosyncrasy of each genetically modified mouse strain will have to be borne in mind when deciding whether fasting is appropriate and, if so, for how long. For example, a 4- to 6-hr fast may be more appropriate for some strains or disease conditions for which an overnight fast is too stressful.

a. For thoracic tumors, such as lung tumors or pulmonary metastases, mice can be fed a specifically formulated commercial high-fat diet (such as, for example, diet D12451 from Research Diets, with 45% of total calories from fat) or, alternatively, sunflower seeds, which are rich in vegetable fats, during the 24 hr prior to the day of analysis. In this case, mice will not be fasted during the night previous to the analysis. This will decrease glucose uptake by the myocardium (the so called “FDG-robbing effect”), eliminating, to a great extent, the interferences arising from the high uptake of the heart under standard feeding conditions. b. For abdominal tumors, such as those of the stomach, pancreas, colon, etc., it is absolutely essential that the mice be kept in the absence of food for at least 4 hr previous to PET exploration. Water will be always supplied. c. For brain tumors, there is no need for any special diet or fasting prior to the exploration. The feeding conditions of the mice in this case are the standard ones. d. Fasting prior to the exploration can be omitted in those mice with a compromised health status. This results in a slight decrease in the quality of the image but, in turn, it enhances the chances of mice surviving the procedure. Fasting increases the ratio of uptake between the tumor and the surrounding healthy tissue. In addition, it diminishes the uptake by the BAT by decreasing diet-induced thermogenesis. In case of diabetic mice, a glucose determination test will be done prior to the procedure and, if needed, insulin can be administered to reduce hyperglycemia (reducing uptake competition of the probe with circulating glucose). It must be borne in mind that this will result in an increase of [18 F]FDG uptake in the striated muscle, rendering a noisier image (showing, for instance, hotspots in the limbs).

Sedate and anesthetize mice 2. Transport mice to the imaging unit the day before the exploration, prepare the diet, and determine their weight in order to prepare the adequate doses of sedatives and anesthetics. 3. Administer an intraperitoneal (i.p.) injection of a muscle relaxant (diazepam) at a dose of 3 mg/kg, 20 to 60 min before commencing the procedures. This will decrease FDG uptake by muscles.

4. Place an infrared lamp above the cage to keep the mice warm. Imaging Cancer in Mice by PET, CT and PET-CT

Normal body temperature in mice should be 36◦ to 37◦ C, and this should be monitored closely with a rectal probe (Fig. 1B, C). Temperatures higher than 40◦ C imply risk of dehydration. Temperatures lower than 35◦ C may result in hypothermia.

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Figure 1 Equipment at a PET-CT facility for mice. (A) PET-CT machine (eXplore Vista, GE Healthcare). (B) Monitor of vital constants (VisionVet, RGB) registering ECG, temperature, and respiratory frequency. (C) Mouse fitted with electrodes and monitoring probes being introduced into a PET-CT machine.

5. Hold the mouse carefully in your hand and bring its tail closer to the infrared lamp (∼5 cm) during 3 to 4 min, taking care not to expose the rest of the body to excessive heating. This procedure results in vasodilation at the tail, thus facilitating injection of [18 F]FDG through the caudal veins.

6. Once the tail veins have dilated, introduce the mice into the anesthesia chamber where a deep anesthesia will be induced by inhalation of 2% isoflurane in 100% oxygen at a rate of 1 liter/min. In our hands, inhaled isoflurane is more effective and less harmful than other anesthetic agents, although noninhalant anesthetics can be used. In this last instance, our recommendation is i.p. injection of a mixture containing ketamine (200 mg/kg) and xylazine (10 mg/kg). Protocols for anesthesia may vary by institution, so consult your veterinary staff.

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Figure 2 Administration of the probe. Tail vein injection of the radioactive probe into a mouse with anesthetic mask and using a shielded syringe.

Administer the radiolabeled probe 7. Transfer anesthetized mice to the injection area and, using a tuberculin syringe and a 30-G needle, inject them with [18 F]FDG in the tail (Fig. 2) at a dose of 500 μCi in a volume of 0.2 ml physiological saline (0.9% NaCl), calibrating the dosage using an activimeter. The injection of 500 μCi per mouse corresponds to 5 to 50 ng of [18 F]FDG, which is very low compared to the normal glycemia and therefore without pharmaceutical effect. It is advisable to make the first attempt at injection in a distal location on the tail (Fig. 2). In case of failure, this will make it possible to repeat the injection in more proximal positions. In case of excessive radioactive decay, it may not be possible to administer a 500-μCi dose, in which case we advise injection of as much activity as possible in a maximum volume of 0.2 ml. It is not worthwhile to inject less than 100 μCi because it will result in poor-quality images. To ensure reproducibility, it is advisable to use the same dose and volume in all the mice, in order to be able to later compare the results of the quantification of [18 F]FDG uptake. All the procedures involving handling of the [18 F]FDG must be carried out with double-gloved hands and using radioprotection devices to maximize the safety of the operator.

8. Once injected, maintain mice under anesthesia during the duration of the [18 F]FDG uptake time, which is 45 min. Imaging Cancer in Mice by PET, CT and PET-CT

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During this period, [18 F]FDG distributes all over the body. For that reason, it is paramount to keep the mice under the best possible conditions, which include minimal stress achieved by anesthesia and sedation (with the added benefit of minimizing muscle uptake), and body temperature around 36◦ to 37◦ C, aided by the infrared lamp (to minimize BAT uptake). Current Protocols in Mouse Biology

IMPORTANT NOTE: In case of brain tumors, or if the health of the mice is severely compromised, perform the injection and uptake in the absence of anesthesia. In this case, mice will be kept in their cage with appropriate shield protection and visible warning signs. For PET-CT imaging, the CT image acquisition (15 min) is carried out during the [18 F]FDG uptake period. In this manner, wait for 30 min after [18 F]FDG administration and then carry out the CT study; the PET study will be performed immediately after the CT one is finished.

Perform image acquisition 9. Before placing mice on the exploration table, administer eye lubricant (Lacryvisc Gel 10G) to mice to avoid lesions in the cornea while the mice are anesthetized, since under these conditions the blinking reflex is lost. During image acquisition, mice must be anesthetized with a mask providing 2% isoflurane in 100% oxygen at 1 liter/min. 10. Set the heating pads on the exploration table of the PET machine at a constant temperature of 37◦ C. Place a blotter on top of the heating pad to collect possible urine from the mouse. When studying nude mice or mice of compromised health, cover the mice with a blanket of plastic bubbles to avoid heat dissipation as much as possible.

11. Place mice on the exploration table as stretched out as possible to minimize organ superposition. Mice should be fixed to the exploration table with adhesive tape even if they are anesthetized, to minimize involuntary movements (Fig. 1C). The size of the bed depends on the dimensions of the detector and on the number of crystals (in the case of the eXplore Vista PET-CT, one bed position corresponds to 47 mm). Always make sure that the exploration table is free of possible residues of urine from a previous mouse, since urine will contain radioactive probe and will produce false zones of probe hyper-uptake. A new blotter will always be used for each exploration.

12. Perform image acquisition as described below. The duration of the PET exploration will be ∼20 to 30 min. The standard setup values are as follows: 20 to 30 min for whole acquisition in 1 bed position, and 45 min for whole acquisition in 2 bed positions (bed positions are units of measurement in imaging studies that refer to the length of the static field of view). In mice with compromised health, PET acquisition can be reduced to 10 min in 1 bed position. PET acquisition times below 10 min will result in poor-quality images. If only 1 bed is registered, focus must be on the anatomical area of interest. If the likely location of the primary tumor is unknown or extension studies are needed because of possible metastasis, register 2 beds, ensuring that the whole body of the mouse is explored.

a. Perform a blank test. This test consists in performing an empty acquisition, without any radiation. It must be done daily, before any real work is carried out, to assess the correct functioning of the glass detectors of the PET equipment. If the system is working properly, as determined by the blank test, then the user can start the normal operation of the PET. The software program MMWKS VISTA CT (Pascau et al., 2006) is launched, the option “PET ACQ” is ticked, and a static study of 5 min duration is selected, without any isotope.

b. Decide in advance the anatomical area of interest because, if possible, it is preferable to perform 1 bed position, which should take between 20 and 30 min. If it is necessary to explore the entire body, then perform 2 bed positions (sufficient for a mouse of standard size), and PET acquisition time in this case should not exceed 45 min.

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c. Divide the acquisition into two parts. Select or create the folder where the study will be saved and create what is known as a scout (the scout delimitates the area to be studied), or use a scout previously created (if a CT of the same mouse has already been performed, a scout for this given mouse will already be available). Once the area to be studied has been determined, select the kind of PET study to be carried out, define the duration of the procedure, the isotope to be used, and the energy window of the isotope of choice. In the authors’ instance the isotope is 18 F and the window is fluorine.

d. There are some predefined PET acquisition protocols currently available. Static (1 bed): This protocol allows for one bed position; it requires previous knowledge of the anatomical region of interest (i.e., abdominal or thoracic). This is the protocol of choice for a standard tumor study. While acquisition is in progress, take notice of the number of coincidences; this value should be between 200 and 500. This serves as a quality control for the injection of the radiolabeled compound. The number of coincidences is equivalent to the number of detected photons arising from radioactive disintegrations. If that number of coincidences is not reached, it will not be possible to perform PET acquisitions. In this last instance, the user should inject another dose of the probe again, so as to have coincidence events to be detected. Whole-body (≥2 beds): This protocol allows the acquisition of two or more bed positions. Usually, two bed positions suffice for the complete exploration of the entire body of a mouse of average size. With the eXplore Vista PET-CT, it is possible to study rats, since it is prepared to admit settings of more than two bed positions. This would be the protocol of choice when studying adult mice for which the anatomic location of the tumor is not known or when the user wants to investigate the possible existence of distant metastases. In this instance, the user can either set the same duration time for each one of the two beds or set a larger time for one of the beds, depending on the area of interest to be studied or the area for which he/she would like to have higher counting statistical values, but never exceeding a total exploration time of 45 min. Dynamic studies: This protocol may cover one or more beds. These are the protocols of choice for studies of probe kinetics and biodistribution, and also for gating studies. Dynamic studies need to predefine the periodicity and duration of each frame. In the case of gating, images are acquired concurrently with respect to an external signal, such as breathing or electrocardiogram (ECG; Fig. 1B, C). After acquisition, during image processing, frames are grouped according to the external signal. ECG and respiratory signals are obtained from an external monitoring device (Fig. 1B, C). Respiratory gating registers inhaling and exhaling, and this is of relevance in CT studies of lung tumors (see Basic Protocol 2). The feasibility of separating systole from diastole by means of the ECG-gated trigger signal is of relevance in PET studies, since this makes it possible to carry out studies of cardiac functionality (imaging systole and diastole) and to calculate the ejection fraction.

e. Perform the PET exploration setting the type of isotope to 18 F and set the lower energy threshold to 150 KeV. 13. Once PET acquisition is finished, retrieve mice out of the equipment, disconnect the anesthesia, and take mice to a warm cage where they will wake up on their own. Ensure that the mice are placed in a way that it facilitates their recovery and that their breathing path is free of any obstacles (pellets of food, shreds of litter, etc.). Keep the cage containing the mice in a shielded isolation rack. If it is not required that the mice be returned to their housing cages, they may be kept overnight at the imaging facility to allow for a complete radioactivity decay and returned to their housing cells the next day. Imaging Cancer in Mice by PET, CT and PET-CT

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Reconstruct PET image 14. Choose an option for image reconstruction. Three options for image reconstruction are available in the eXplore Vista PET-CT equipment: 2D-FBP (filtered back projections), 2D-OSEM (ordered-subsets expectation maximization), and 3D-OSEM. The authors’ preference is 3D-OSEM with the number of iterations set to eight and employing random and scatter correction. In this manner, we achieve a good imaging quality with reasonable computing resources and usage and an acceptable calculation running time (it must be borne in mind that an independent PC is needed to run 3D-OSEM). The average running time for a 3D-OSEM calculation (eight iterations, random and scatter correction) in a Dell Studio XPS Desktop 435MT PC is <2 min.

Quantify the data 15. Once the total scan volume is reconstructed, inspect the images until a positive [18 F]FDG signal is identified in an anatomical position consistent with a tumor, and distinct from the normal uptake organs (e.g., heart, urinary bladder, kidneys, and to a lesser extent BAT). 16. Manually draw a region of interest (ROI) around the perimeter of the tumor. Subsequently, inspect adjacent sections where the software has automatically drawn the same ROI; if adjustment is needed, it can be done manually. When reconstructing a volume, different types of geometric voxels, such as spheres, cubes, etc., can be used. The 3D-OSEM reconstruction algorithm employs cubic voxels. The software predetermines the size of the reconstructed cubic voxel; in our equipment, its value is of 0.7 × 0.7 × 0.7 mm. Therefore, tumor volumes are built in unit volumes of cubic voxels. It has to be verified that all the tumor activity has been included. The quantification of [18 F]FDG uptake contained in the ROI is performed automatically, expressed in MBq/cc.

17. From the quantification described under the previous step, calculate the Standardized Uptake Value (SUV). Two types of SUV measurements can be calculated using the average SUV (SUVave ) and the maximum SUV (SUVmax ) formulas below:

SUVave = [uptake (MBq)/volume of the ROI (cc)]/[mouse weight (g) × injected [18 F]FDG dose (MBq) × calibration factor] SUVmax = [uptake in the voxel with maximal activity (MBq)/volume of the voxel (cc)]/[mouse weight (g) × injected [18 F]FDG dose (MBq) × calibration factor] SUVave reflects the amount of radioactivity per unit of volume across the entire ROI volume. SUVmax reflects the radioactivity per unit of volume at the voxel, within the ROI, with the maximal value of uptake. In the particular case of mice, due to the great variability in radiolabeled compound distribution, it is advisable to normalize the results of the ROI of interest to a reference ROI in an organ of stable uptake, such as the liver. Data, for example, can be expressed as the ratio of tumor SUVave versus liver SUVave .

IMAGING BY COMPUTED TOMOGRAPHY (CT) CT studies are simpler than PET since CT requires little or no preparation. However, the mice must be immobilized under anesthetics to avoid artifacts arising from movement and to maintain a constant body temperature to prevent anesthesia-induced hypothermia. Exogenous contrast agents are often used to improve the signal ratio between the tumor and the surrounding healthy tissue. Contrast agents enhance the density of the tissue to be imaged. Contrast is provided by the presence of a heavy atom. The presence of heavy

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atoms in the tissues modifies the intensity of the X-ray traversing the organ, and this is used to generate an image of reduced noise. In the case of soft-tissue tumors, detection requires a contrast agent. In humans, water-soluble contrast agents are frequently used; however, this does not apply well to mice. One reason is that the CT technology for small animals is not as rapid in capturing images as it is for humans, with acquisition times between 10 and 20 min in animals versus seconds in humans. Thus, the watersoluble contrast agents used in human studies (such as Iopamiro, see below) are rapidly cleared through the kidney (within minutes in humans and seconds in mice), not allowing sufficient time to acquire CT images in mice. To circumvent this problem, some other contrast agents specific for small animals, including mice, have been developed. They are discussed in the protocol below.

Materials Computed tomography system (e.g., CT Locus from GE Healthcare or CT eXplore Vista from GE Healthcare) eXplore Vista PET-CT MMWKS software for image acquisition, processing and analysis (incorporating a modified version of the FDK algorithm for CT reconstruction); or Microview for Locus CT for image analysis Additional reagents and equipment for imaging by PET (Basic Protocol 1) To perform CT without contrast 1a. Anesthetize and warm mice; apply some eye lubricant (Lacryvisc Gel 10G) to the cornea for protection as specified in Basic Protocol 1 before placing mouse inside the scanner chamber. For CT acquisition the mice do not require prior preparation (fasting and sedation are not required).

2a. Before starting the CT, select the area of interest and properly set the different parameters. The parameters for CT acquisition are usually set within the following ranges of values: i. Intensity of the power supply: from 140 to 1000 mA. ii. Number of shots: from 1 to 32. The number of shots refers to the number of times that X-rays are emitted from the source. Although a higher number of shots results in a higher signal-to-noise ratio in the image, it also results in larger acquisition times and greater radiation damage inflicted on mice.

iii. Resolution: standard (200 μm), high (100 μm), or maximum (50 μm). iv. Number of projections: from 360◦ to 720◦ . The number of projections refers to the number of rotations of the X-ray beam around the mice, expressed in sexagesimal degrees. Double full rotations (720◦ ) versus single full rotation (360◦ ) results in an image of higher signal-to-noise ratio, but at a cost of larger acquisition times and, consequently, greater exposure of the mice to a hazardous energy source.

v. Number of bed positions: 1 to 3. When more than one bed position is selected, they superimpose on top of each other, resulting in an overlap in that area. Try to minimize the overlap as much as possible so that it does not coincide with the anatomic area to be studied, since the overlapping of two beds is not perfect and may give rise to aberrant images.

vi. Voltage of the power supply: from 30 to 80 kV. Imaging Cancer in Mice by PET, CT and PET-CT

When studying soft tissues, it is advisable to set the voltage at a value of 30 kV; for in vivo bone studies the recommended value is 50 kV; for isolated ex vivo bones, the recommended value is 80 kV.

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3a. Fix mice to the exploration table with adhesive tape even if they are anesthetized, to minimize involuntary movements. Once the mice are ready on the exploration table, lower down the protective shielded screen of the Vista-PET-CT and lock in the safety key to perform data acquisition.

Standard CT data acquisition 4a. Set the power supply to an intensity of 150 mA and a voltage of 45 kV for a standard resolution of (200 μM), 360◦ , and 16 shots for 1 bed position, or 8 shots if performing 2 bed positions. If performing single-bone studies, select the maximum resolution values as long as the CT scan does not exceed a total duration of 30 min. As an example, the standard acquisition to visualize pulmonary tumors at a standard resolution of (200 μM) is 1 bed position and 16 shots, a voltage of 45 kV, and an intensity of 150 mA. Once all the parameters have been selected and set, the system will inform on the duration of the acquisition, which in CT studies is about 15 min. The above acquisition parameters correspond to a radiation dose in the mouse of 0.6 Gy.

5a. Observe the scout image that the CT machine will generate. This scout gives a projection where the user can see how the subject is placed.

6a. Select, with the aid of a laser, the beginning and the end of the ROI that the user would like to acquire (e.g., the whole mouse). Scouts have the extension .plan; if a study is aborted, the scout has to be repeated, since it cannot be used again. If the mouse moves excessively during image acquisition, the complete CT must be repeated including a new scout. We can also select a previous scout from a study already finished as long as it corresponds to the same mouse and we have not altered its disposition on the exploration table.

Respiratory gating for thoracic CT 7a. Obtain respiratory signals from an external monitoring device simultaneously with the CT scan (Fig. 1B, C; respiratory movement artifacts are important in CT acquisition, especially in the case of thoracic studies, including lung tumors). After acquisition, during image processing, group frames according to the external signal, generating two independent scans, one corresponding to the inhalation and the other to the exhalation (grouping into intermediate stages is also possible). 8a. Stop the anesthetic delivery once CT acquisition is over, retrieve the mice out of the equipment, and take them to a warm cage where they will wake up on their own.

CT image reconstruction 9a. Reconstruct images using a modified version of the cone-beam (CB) algorithm of Feldkamp, Davis, and Kress (FDK; Vaquero et al., 2008) included in the software of the eXplore Vista PET-CT. FDK is a widely used filtered-back projection algorithm for three-dimensional image reconstruction from cone-beam projections measured with a circular orbit of the X-ray source. The modified version by Vaquero et al. (2008) makes use of a certain property of this approximate algorithm (the integral of the reconstructed image along any axial line orthogonal to the plane of the orbit is exact when the cone-beam projections are not truncated) to speed up volume calculation times without compromising the quality of CT reconstruction. Reconstruction time when the algorithm is run on a Dell PowerEdge workstation is <5 sec. Imaging Cancer in Mice by PET, CT and PET-CT

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Figure 3 Example of ROI selection in a CT scan. (A) The example shows three serial CT transversal sections separated by 0.6 mm. The operator manually outlines the perimeter of the tumor in a section where the tumor is clearly detectable (such as in the middle section). The software automatically draws the same outline in adjacent sections separated every 50 μm. The operator can modify the outline to adapt it to the changing dimensions of the tumor (such as in the upper and lower sections). The software integrates all the sections into the volume corresponding to the ROI (region of interest). (B) From top to bottom, sagittal, coronal, and transverse sections of the CT volume. The ROI, visible in all three sections, is shown in solid green. Imaging Cancer in Mice by PET, CT and PET-CT

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Quantification using a ROI 10a. Identify tumors by visual inspection of the images. 11a. Draw manually a region of interest (ROI) around the perimeter of the tumor. Subsequently, inspect adjacent sections where the software draws the same ROI; if adjustment is needed, it can be done manually (Fig. 3). When reconstructing a volume, different types of geometric voxels, such as spheres, cubes, etc., can be used. The software predetermines the size of the voxels. Our reconstruction software employs cubic voxels of 50 × 50 × 50 μm to define the ROI. The volume of the selected ROI, expressed in cubic centimeters, is automatically calculated by the system.

Perform CT with contrast 1b. Follow step 1a as above. Administer the contrast agent prior to introducing the mice into the X-ray tube. This is done either through tail vein injection, oral, or i.p. administration. Several contrast agents are available: i. Lopamiro 300 Iopamidol (Bracco). Lopamiro is a nonionic water-soluble contrast agent. This type of agent has a reduced general toxicity, even in delicate structures such as vascular endothelia and the central nervous system. The product is available as pre-constituted solution in four different concentrations. The authors use 50-ml aliquots of Iopamiro 300 containing 30.62 g of Iopamidol, and employ a dose of 0.18 ml (which corresponds to 0.11 mg) per 30 g of body weight. ii. Fenestra VC and Fenestra LC (Alerion Biomedical). The Fenestra technology is based on emulsions of iodinated lipids. Fenestra LC (LC, liver contrast) provides visualization of the entire hepatobiliary system. Fenestra VC (VC, vascular contrast) is a variant of Fenestra LC that provides superior contrast enhancement of the entire body vasculature for up to several hours after injection. This product is usually distributed at a concentration of 50 mg/ml; the recommended dose for mice is of 0.3 ml/30 g. iii. Exia 160XL (Binitio Biomedical, Inc; http://www.binitio.com). Exia 160XL is an aqueous colloidal polydisperse contrast with prolonged blood-pool effect. This product is usually distributed at a concentration of 160 mg/ml; the recommended dose for mice is of 0.10 ml of the stock solution (which corresponds to 16 mg) per 30 g of body weight. Fenestra and Exia remain in the organism for long periods of time, and they are useful in vascular studies. In our workplace, we routinely use Iopamiro. For liver studies, we administer Iopamiro intravenously; for pancreas and colon, we use oral or intraperitoneal administration. Theoretically, Fenestra and Exia are small-animal-dedicated contrast agents with slow kidney clearance, but, in our experience they do not lend great additional advantages over Iopamiro, and they are much more costly.

2b. The rest of the acquisition parameters are the same as those for CT without contrast studies. Perform steps 2a to 11a, as described under CT without contrast.

IMAGING BY MULTIMODALITY (PET-CT) The immobilization of the mice during the exploration is essential for image coregistration in multimodality imaging studies. If the positions of the mice during PET and CT studies are different, the matching (coregistration) of the images obtained by PET and CT is not possible.

BASIC PROTOCOL 3

Materials PET-CT imaging system (e.g., eXplore Vista PET-CT, GE Healthcare; Fig. 1A) Additional reagents and equipment for PET (Basic Protocol 1) and imaging by computed tomography (Basic Protocol 2)

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Prepare mice 1. Prepare the mice prior to the multimodality exploration as described for the PET exploration in Basic Protocol 1, steps 1 to 6. Multimodality exploration requires extreme attention to the care of the mice as well as constant monitoring of the temperature, since the study times are longer than in CT or PET alone.

Acquire CT image 2. Perform the acquisition of the CT image during the [18 F]FDG uptake period (lasting 45 min). See Basic Protocol 1, step 8, and Basic Protocol 2. Acquire PET image 3. Perform PET image acquisition following the same procedure as described in Basic Protocol 1, steps 7 to 13. Reconstruct images 4. Reconstruct both images separately and use the MMWKS software to obtain the co-register (Fig. 4).

A

B

Iv

h

C

Imaging Cancer in Mice by PET, CT and PET-CT

D

Figure 4 PET-CT imaging of a lung tumor. (A) Transverse CT section. From top to bottom, liver (lv) and heart (h). The orange arrow points to the ROI, a lung tumor. (B) Transverse PET section of the same anatomic area as in A. (C) Histological section of the same lung imaged at the optical microscope by Hematoxylin & Eosin staining. Two tumors of different dimensions were detected: the one already observed by PET-CT (orange arrow) and a smaller one (blue arrow) that was not detected by PET-CT probably due to its intrabronchial location. (D) 3-D rendering of the PET-CT image merged with the lung. The tumor is shown as a pinkish-colored small ball.

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vehicle

therapeutic compound

month

0

⫹2

⫹4

⫹8

Figure 5 Example of a longitudinal study of lung cancer. 3-D lung renderings of PET-CT studies at the indicated time-points of two individual mice carrying genetically induced lung tumors, either treated with vehicle (upper mouse) or with a therapeutic compound (lower mouse). Mouse model courtesy of Dr. Mariano Barbacid (CNIO).

Quantify images 5. Once the tumors have been identified in the organs of interest, quantify the CT image and the PET image separately, as described in Basic Protocols 1 and 2. Rendering of volumes 6. Generate 3-D renderings of the data to obtain information on tumor location and volume in a more graphical way (Fig. 5; Videos 1 to 3). REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps.

Diazepam, 5 mg/ml 5 mg/ml diazepam (Valium, Roche Pharmaceuticals) 40% (v/v) propylene glycol 10% (v/v) ethanol 5% (w/v) sodium benzoate/benzoic acid buffer, pH 6.6 (6.2 to 6.9) 1.5% (v/v) benzyl alcohol (as preservative) Store up to 1 year at room temperature COMMENTARY Background Information Until recently, the study of cancer in mice has been limited by the inability to use imaging techniques of sufficient sensitivity and resolution. In the absence of appropriate imaging technology, each individual mouse can only provide one measuring time point. Also, if the interest is to study a therapeutic treatment, in the absence of imaging technology, there is always a certain degree of uncertainty about whether the treated mice carry tumors or not. In general, endogenous cancers in in-

ternal organs, such as liver, lung, or pancreas, are asymptomatic during early stages and appear asynchronously in each individual mouse. These limitations end up requiring prohibitive numbers of mice, unnecessary use of therapeutic drugs that can be very expensive, long tests, and laborious analyses before obtaining meaningful data. The application of imaging technology circumvents these problems by allowing for the acquisition of multiple measuring time points from each mouse. This allows application of the therapeutic treatments

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at the right time for each mouse. In addition, the current imaging techniques provide an accurate measurement of volumes and also of function (e.g., glucose uptake) of individual tumors. The combined application of PET and CT requires a significant investment, but its costs are compensated by the decrease in the number of mice used, the total experimentation time involved, and the quality and speed of the data obtained.

Critical Parameters

Imaging Cancer in Mice by PET, CT and PET-CT

When applying imaging techniques to murine tumors, it is of key importance to adhere to a very careful handling procedure with respect to the mice. It has to be taken into account that, to carry out the study, the mice will be placed in unusual situations, which may be fatal for them. Therefore, it is vital that each study be customized to the specific mouse strain or genetically modified mouse cohort being analyzed, adapting fasting times and doses of anesthetics to the particular needs of the mice. The maintenance of body temperature and vital constants is fundamental to the study. As it is the case for any procedure that involves administration of anesthesia, monitoring the mice during the recovery time is essential for their well-being, and for a successful outcome. If hypothermia is observed, increase the temperature of the cage. It is well demonstrated that the handling of the mice has a profound impact on the biodistribution of [18 F]FDG and that it significantly influences the visualization of tumors. Varying fasting state, body temperature, and mode of anesthesia may affect the [18 F]FDG uptake in normal organs by a factor of 10-fold, and in tumors by a factor of 3.7-fold (Fueger et al., 2006). The influence of blood glucose and insulin levels on [18 F]FDG biodistribution is also well known (Wahl et al., 1992). Because [18 F]FDG competes with glucose for intracellular transport and phosphorylation, tumor [18 F]FDG uptake decreases with increasing blood glucose levels. Therefore, tumor [18 F]FDG uptake and image contrast are lower in the nonfasted state (high insulin and glucose levels) than in the fasted state (low insulin and glucose levels). The effect of ambient temperature on [18 F]FDG uptake in BAT has been described in human patients (Cohade et al., 2003). However, these effects are even more pronounced in mice. At room temperature (21◦ to 22◦ C), mice need to generate heat by the activation of

BAT and muscle to maintain a stable body temperature. Mice that were kept at 30◦ C showed a markedly lower [18 F]FDG uptake by BAT and muscle (Fueger et al., 2006). [18 F]FDG uptake by BAT is also reduced after overnight fasting of the subject mice. Feeding increases the metabolic activity of BAT in rodents. This is considered to represent a mechanism for the stabilization of body weight: an excessive caloric intake is converted to heat by BAT. The standardization of mouse handling, anesthesia procedures, and scanning protocols will be essential to ensure that data obtained from [18 F]FDG PET-CT imaging studies of transgenic mouse models performed at different institutions are reproducible and comparable. It has to be borne in mind that as longitudinal studies progress in time so does tumor development, and the health status of the mice deteriorates progressively. For this reason, at the beginning of longitudinal studies, the user has to assess the optimization of all the acquisition parameters so the results obtained from the quantification of the tumors do not alter. Similar conditions must be maintained over the study period, since the quality of life of the mice will worsen as the longitudinal study progresses.

Troubleshooting Table 1 lists the potential problems that may be encountered when performing PET, and Table 2 lists those related to CT or PET-CT.

Anticipated Results In the case of lung and liver tumors, where it is possible to identify and follow individual tumors within individual mice, the application of PET-CT imaging technology allows for longitudinal studies in cohorts of about 10 mice per treatment condition. In therapeutic studies, mice are treated only when the presence of a sufficient number of tumors has been confirmed by PET-CT. In this manner, in a cohort of 10 mice, it is feasible to obtain data from around 20 to 50 tumors, depending on the tumor multiplicity. With these numbers, it is possible to obtain statistically significant data with a minimum expenditure of mice and time. Moreover, PET provides information about the malignancy of each individual tumor. Very often, mice may carry multiple tumors, but only a minority have acquired the malignant stage characterized by high glucose uptake. This is very valuable information

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Table 1 Troubleshooting Guide for PET Studies

Problem

Possible cause

Solution

No [18 F]FDG uptake even with long exposures (30 min)

Check the mouse tail for extravasation of [18 F]FDG was not injected properly Diabetic mouse with high glucose levels [18 F]FDG Check glucose levels in blood competing with the [18 F]FDG probe

Mice die during imaging

Inappropriate anesthesia

Check that the levels of isoflurane and O2 are correct Check temperature of heating pads; if vital constants are abnormal then increase temperature

Hypothermia

High muscle uptake

Mice are stressed

Keep mice anesthetized during uptake; administer muscle relaxant

High BAT uptake

Mice are cold

Keep constant temperature with the aid of infrared lamp and heating pads during the whole duration of the study

High myocardium uptake during thorax exploration

Myocardium glucose uptake

High-fat diet instead of fasting to decrease myocardium [18 F]FDG uptake

[18 F]FDG uptake in unexpected location (not corresponding to a tumor)

Urine contamination on the exploration table or on the mouse skin/coat

Verify that a new blotter has been fitted to the exploration table. In case of skin contamination, wait 2 hr for decay and start again.

Table 2 Troubleshooting Guide for CT Studies

Problem

Possible cause

Solution

Artifacts in CT image

Breathing movements

Perform respiratory gating

Poor CT image quality

Improper setting of CT parameters

Check settings of intensity and voltage in power supply

Artifacts in PET image merging

Movement of mice during the study

Verify mouse immobilization

No contrast signal

Improper contrast administration or clearance of contrast before CT scan

Verify correct contrast administration and acquisition times after contrast injection

because human cancers correspond precisely to this category of tumors. The response to a therapeutic regime may differ in pre-malignant tumors (PET-negative) compared to malignant tumors (PET-positive). These types of studies are progressively becoming the standard of research for mouse models of cancer therapy.

Time Considerations The preparation of the mice for the exploration requires knowing first the routine of the protocol to be carried out for sedation, anesthesia, heating, and injection of either a radioactive probe (for PET) or contrast agent, if necessary (for CT). After training, these procedures take ∼10 min. For PET, the [18 F]FDG uptake time after the intravenous administration is about 45 min

and the exploration time ranges from 10 to 30 min per bed. Longer times do not improve the quality of the study and may be harmful for mice. The CT studies without contrast do not need previous preparation other than anesthetics for the immobilization of the mice and warming to avoid hypothermia while the mice are asleep. The total time to complete a scan will depend on the desired level of resolution. For tumor location and follow-up, 10 min will suffice. The analysis and reconstruction of the images is quite dependent on the study to be carried out. In the case of longitudinal studies where one is comparing with previous studies on the same mouse, prior PET and CT images are required to be displayed on the screen to confirm measurements on the same tumor.

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For studies aimed at assessing the effectiveness of drugs or other therapeutic modalities, the experiments should be planned in consultation with a statistician to ensure that the minimum number of treated and control mice are used while still allowing the acquisition of statistically significant data. The use of in vivo imaging for longitudinal quantification of tumor burden is advantageous and should significantly reduce the number of mice needed.

Acknowledgements We are very grateful to Coral Velasco, Elena Andr´es, and Juan Antonio C´amara for their expertise and key contributions to the optimization of the protocols described here. We are also indebted to Isabel Blanco, Manuel Desco, and Juanjo Vaquero for their continuous help and support. This study is partially funded by CDTI under the CENIT Programme (AMIT Project) and supported by the Spanish Ministry of Science and Innovation.

Literature Cited Abbey, C.K., Borowsky, A.D., McGoldrick, E.T., Gregg, J.P., Maglione, J.E., Cardiff, R.D., and Cherry, S.R. 2004. In vivo positron-emission tomography imaging of progression and transformation in a mouse model of mammary neoplasia. Proc. Natl. Acad. Sci. U.S.A. 101:1143811443. Berger, F., Lee, Y.-P., Loening, A.M., Chatziioannou, A., Freedland, S.J., Leahy, R., Lieberman, J.R., Belldegrun, A.S., Sawyers, C.L., and Gambhir, S.S. 2002. Whole-body skeletal imaging in mice utilizing microPET: Optimization of reproducibility and applications in animal models of bone disease. Eur. J. Nucl. Med. Mol. Imaging 29:1225-1236. Cohade, C., Mourtzidos, K.A., and Wahl, R.L. 2003. “USA fat”: Prevalence is related to ambient outdoor temperature evaluation with 18 FFDG PET/CT. J. Nucl. Med. 44:1267-1270. Dearling, J.L., Flynn, A.A., Sutcliffe-Goulden, J., Petrie, I.A., Boden, R., Green, A.J., Boxer, G.M., Begent, R.H., and Pedley, R.B. 2004. Analysis of the regional uptake of radiolabeled deoxyglucose analogs in human tumor xenografts. J. Nucl. Med. 45:101-107. Dendy, P. and Heaton, B. 1999. Tomographic imaging. In Physics for Diagnostic Radiology (P. Dendy and B. Heaton, eds.) pp. 249-278. Institute of Physics, Bristol, U.K.

Imaging Cancer in Mice by PET, CT and PET-CT

Dilmanian, F.A., Wu, X.Y., Parsons, E.C., Ren, B., Kress, J., Button, T.M., Chapman, L.D., Coderre, J.A., Giron, F., Greenberg, D., Krus, D.J., Liang, Z., Marcovici, S,, Petersen, M.J., Roque, C.T., Shleifer, M., Slatkin, D.N., Thomlinson, W.C., Yamamoto, K., and Zhong, Z. 1997. Single- and dual-energy CT with monochromatic synchrotron X-rays. Phys. Med. Biol. 42:371-387.

Fueger, B.J., Czernin, J., Hildebrandt, I., Tran, C., Halpern, B., Stout, D., Phelps, M.E., and Weber, W.A. 2006. Impact of animal handling on the results of 18F-FDG PET studies in mice. J. Nucl. Med. 47:999-1006. Gambhir, S.S. 2002. Molecular imaging of cancer with positron emission tomography. Nat. Rev. Cancer 2:683-693. Holdsworth, D.W. and Thornton, M.M. 2002. Micro-CT in small animal and specimen imaging. Trends Biotechnol. 20:S34-S39. Massoud, T.F. and Gambhir, S.S. 2003. Molecular imaging in living subjects: Seeing fundamental biological processes in a new light. Genes Dev. 17:545-580 Pascau J., Vaquero, J.J., Abella, M., Cacho, R., Lage, E., and Desco, M. 2006. Multimodality workstation for small animal image visualization and analysis. Mol. Imaging Biol. 8:9798. Paulus, M.J., Gleason, S.S., Easterly, M.E., and Foltz, C.J. 2001. A review of high-resolution X-ray computed tomography and other imaging modalities for small animal research. Lab. Anim. (NY) 30:36-45. Schelbert, H.R. 1998. The usefulness of positron emission tomography. Curr. Probl. Cardiol. 23:69-120. Strijckmans, K. 2001. The isochronous cyclotron: Principles and recent developments. Comput. Med. Imaging Graph. 25:69-78 Toyama, H., Ichise, M, Liow, J.S., Vines, D.C., Seneca, N.M., Modell, K.J., Seidel, J., Green, M.V., and Innis, R.B. 2004. Evaluation of anaesthesia effects on 18 F-FDG uptake in mouse brain and heart using small animal PET. Nucl. Med. Biol. 31:251-256. Vaquero, J.J., Redondo, S., Lage, E., Abella, M., Sisniega, A., Tapias, G., and Desco M. 2008. Assessment of a new high-performance smallanimal X-ray tomography. IEEE Trans. Nucl. Sci. 55:898-905. Wahl, R., Henry, C.A., and Ethier, S.P. 1992. Serum glucose: Effects on tumor and normal tissue accumulation of 2-[F-18)]-fluoro-2-deoxyD-glucose in rodents with mammary carcinoma. Radiology 183:643-647. Wang, Y., Seidel, J., Tsui, W., Vaquero, J.J., and Pomper, M.G. 2006. Performance evaluation of the GE Healthcare eXplore VISTA dualring small-animal PET Scanner. J. Nucl. Med. 47:1891-1900.

Internet Resources https://www2.gehealthcare.com/portal/site/usen/ menuitem.e8b305b80b84c1b4d6354a1074 c84130/?vgnextoid=ea21351f9b7e0210Vgn VCM10000024dd1403RCRD&productid= da21351f9b7e0210VgnVCM10000024dd1403 General Electric Triumph Tri-modality PET/ SPECT/CT. http://www.sedecal.com/en/divisiones/division prod.php?p=44&c=5&i=2 SEDECAL Argus PET-CT.

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http://www.medical.siemens.com/webapp/wcs/stores/ servlet/CategoryDisplay∼q catalogId∼e -11∼ a categoryId∼e 1029715∼a catTree∼e 100010, 1007660,1011525,1029715∼a langId∼e 11∼a storeId∼e 10001.htm Siemens Inveon PET & Inveon CT. http://www.healthcare.philips.com/main/products/ preclinical/ Philips Mosaic HP PET.

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Arterial Pressure Monitoring in Mice Xin Zhao,1 David Ho,1 Shumin Gao,1 Chull Hong,1 Dorothy E. Vatner,1 and Stephen F. Vatner1 1

The University of Medicine & Dentistry of New Jersey, New Jersey Medical School, Newark, New Jersey

ABSTRACT The use of mice for the evaluation and study of cardiovascular pathophysiology is growing rapidly, primarily due to the relative ease of developing genetically engineered mouse models. Arterial pressure monitoring is central to the evaluation of the phenotypic changes associated with cardiovascular pathology and interventions in these transgenic and knockout models. There are four major techniques for measuring arterial pressure in the mouse: tail-cuff system, implanted fluid-filled catheters, Millar catheters, and implanted telemetry systems. Here we provide protocols for their use and discuss the advantages C 2011 and limitations of each of these techniques. Curr. Protoc. Mouse Biol. 1:105-122  by John Wiley & Sons, Inc. Keywords: arterial pressure monitoring r mice r methods

INTRODUCTION Since the mouse is most suited to the development of different transgenic and knockout models, potential biological and molecular targets are further explored in these genetically altered mice. It is essential to characterize and evaluate the phenotypic changes that are present within these different models. One key variable that must be monitored and considered for cardiovascular pathophysiology research is arterial pressure. The accurate determination of arterial pressure is important not only for pathological conditions such as hypertension or determining the effects of pharmacological intervention, but is also important for the assessment of the condition of the animal at the time of the experiment. Excessively elevated or depressed arterial pressure may affect the experimental design and values. Indeed, it is not possible to interpret left ventricular (LV) function and ejection fraction (EF) in the absence of afterload, which is most commonly assessed by systolic arterial pressure. The major techniques for measuring arterial pressure in the mouse are described in this unit: tail cuff (Basic Protocol 1), implanted fluid-filled catheters (Basic Protocol 2), Millar solid-state catheters (Basic Protocol 3), and implanted telemetry systems (Basic Protocol 4). Since the implantation of fluid-filled catheters, Millar solidstate catheters, and telemetry systems requires anesthesia and surgery, the authors first compare conscious versus sedated arterial pressure monitoring. Comparing conscious versus sedated arterial pressure monitoring Anesthesia affects the heart, blood vessels, and reflex control of the circulation (Vatner and Braunwald, 1975; Vatner et al., 2002). Accordingly, values obtained under anesthesia for arterial pressure are depressed (Table 1). Because of the hemodynamic effects of the anesthetic agents, studies in conscious animals are preferred when possible (Kurtz et al., 2005). However, it is difficult to train mice to remain still in the conscious state, and movement and excitement all affect arterial pressure. Even though training will help minimize variability, it will not eliminate the normal physiologic response to stress. This is particularly important for experiments requiring a stable baseline of pressure for a prolonged period of time. Furthermore, catheter migration and bleeding are less of an issue in anesthetized animals compared to those studied in the conscious state. For studies

Current Protocols in Mouse Biology 1: 105-122, March 2011 Published online March 2011 in Wiley Online Library (wileyonlinelibrary.com). DOI: 10.1002/9780470942390.mo100149 C 2011 John Wiley & Sons, Inc. Copyright 

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Table 1 Mean Arterial Pressure (MAP) and Heart Rate (HR) in Conscious and Anesthetized Micea Type of anesthesia None (conscious)

Avertin

Pentobarbital

K+X

Isoflurane

Strain

C57/B6SJL

C57BL/6

Webster 4

C57/B6JSL

Swiss, C57BL/6

MAP (mmHg)

103 ± 1

75 ± 2

89 ± 2

64 ± 3

79 ± 3

HR (bpm)

588 ± 14

431 ± 15

514 ± 17

297 ± 15

544 ± 31

Dose

None

300 mg/kg i.p.

60 mg/kg i.p.

65+13 mg/kg, i.p.

2% inhalation

Data resource

Uechi et al. (1998)

Odashima et al. (2007) Ma et al. (2002)

Vatner et al. (2000) Janssen et al. (2004)

a All values are mean ± SE. The heart rate and arterial pressure in conscious mice were much higher than those seen in anesthetized mice.

(K = ketamine, X = xylazine; HR = heart rate; IP = intraperitoneal injection, IM = intramuscular injection, SC = subcutaneous injection).

in tranquilized mice, many choices of injectable or inhalant anesthetics are available. The mixture ketamine/xylazine decreases cardiac contractility and output with the advantage of rapid anesthesia induction and recovery, providing an adequate duration of anesthesia (∼30 min) to allow for the completion of most surgical procedures, while Avertin lowers cardiac function at a more modest level without affecting cardiac output (Hart et al., 2001; Kiatchoosakun et al., 2001), and is thus ideal for hemodynamic monitoring under anesthesia; however, with the relatively short anesthetic duration (10 to 15 min), Avertin is more suited for surgery of relatively shorter duration. Isoflurane results in the lowest number of complications after anesthesia (Pena and Wolska, 2005; Szczesny et al., 2004); however, it may lead to an initial tachycardia with gradual decrease to a heart rate similar to that seen with Avertin (Roth et al., 2002). The major advantage of isoflurane anesthesia is that it allows for titratable continuous anesthesia with a relatively quick recovery time. However, the use of isoflurane requires an anesthetic chamber in a well ventilated operating room with continuous monitoring of oxygen saturation. The hemodynamic effects of some of the common anesthetic agents used in mice are listed in Table 1. BASIC PROTOCOL 1

Arterial Pressure Monitoring in Mice

TAIL-CUFF SYSTEM Tail-cuff systems are routinely used for the monitoring of arterial pressure in rats (van Nimwegen et al., 1973; Krege et al., 1995), and over time this method has been adapted for the monitoring of arterial pressure in mice (Feng et al., 2009). In brief, the system involves a period of training for the mice with repeated arterial pressure measurements taken via the placement of a computer-controlled arterial pressure cuff around the tail of the mouse. The tail-cuff system uses a sphygmomanometer coupled to a method for measuring blood flow in the tail artery, e.g., photoplethysmography or piezoplethysmography. Photoplethysmography utilizes a light source (incandescent or LED) to sense and record the pulse-wave signal, while piezoplethysmography uses piezoelectric crystals to sense the pulse. In general, the tail cuff is inflated to occlude the blood flow, and the disappearance of arterial pressure during inflation or the first appearance of the pressure wave during deflation is taken as the systolic pressure. Since flow cannot be quantified by either of these methods, in both cases diastolic pressure is determined by mathematical calculation. Recently, a new technique has been developed where tail volume as a measure of blood flow into the tail is monitored and used to determine the diastolic pressure as the pressure at which the blood flow into and out of the tail are equalized. This technique provides the most accurate of the three methods for tail-cuff arterial pressure measurements (Feng et al., 2008). Advantages and disadvantages of the tail-cuff system The advantages of this system are that it provides a less costly, simple, noninvasive method for measurements of arterial pressure, mostly in conscious mice. However, there

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Systolic arterial pressure

IA catheter (mmHg)

160 140 120 100

R 2 ⫽ 0.9048 80

B

Mean arterial pressure

160 IA catheter (mmHg)

A

140 120 100

R 2 ⫽ 0.9881 80 60

60 60

80

100 120 Tail cuff (mmHg)

140

60

80

100 120 Tail cuff (mmHg)

140

Figure 1 A comparison of arterial pressure obtained by tail cuff and intra-arterial catheter (IA) using the same male C57BL/6J mouse. A good correlation between systolic, R2 =0.9048 (A), and mean arterial pressure, R2 =0.9881 (B), is observed with the two methods.

is nothing to preclude the use of the tail-cuff system with anesthetized animals. It can also be used for regular monitoring of large groups of mice. A disadvantage is that conscious mice almost always continue to experience some degree of anxiety and stress during balloon inflation on the tail, as evidenced by elevated heart rate (Lorenz, 2002). However, arterial pressure seems to be affected to a lesser extent by this stress (Krege et al., 1995; Ma et al., 2002; Janssen et al., 2004; Whitesall et al., 2004; Odashima et al., 2007), most likely due to the powerful arterial baroreflexes. The degree of correlation between the arterial pressure obtained via tail cuff versus invasive methods is operator dependent (Krege et al., 1995; Whitesall et al., 2004), but in our hands there is a good correlation (Fig. 1). One additional problem is that tail-cuff arterial pressure does not always reflect central arterial pressure (Lorenz, 2002). The inherent properties of peripheral arteries, environmental temperature, increased sympathetic tone, hypotension, or vasoactive substances could all lead to vasoconstriction of the tail artery and affect the accuracy of the measurements. Tail pressure is sensitive to temperature, blood-volume status, and vascular tone; therefore, control of volume status and temperature is essential. Thus, all these factors should be considered when using tail cuff for arterial pressure measurements. Finally, there is a decrease in sensitivity with a narrowing of the pulse pressure (Doevendans et al., 1998), and due to the time required for the inflation and deflation of the tail cuff, continuous monitoring of arterial pressure is not possible. This method is best suited for single-point, periodic monitoring of arterial pressure in mice, where the overall arterial pressure trend may be more important than the instantaneous arterial pressure.

Materials Mice Visitech BP-2000 series II arterial pressure analysis system: four channel mouse platform, control unit, arterial pressure analysis software 1. Train the mice continuously for 5 days prior to the scheduled experiment. During training, the animals are subjected to the entire recording process and are placed in the restrainer before performing steps 2 to 7c, below. A period of 5 days of training prior to the actual experimental recording is recommended by the manufacturer to obtain the most reliable data, as stress from balloon inflation and being in the restrainer can all lead to excitation of the mice and result in falsely elevated heart rate and arterial pressure. With an increasing number of training sessions, the mice are able to adapt more quickly to the procedures and provide more reproducible results. Investigators should record the arterial pressures from each of the daily training sessions (between 1 and 30 measurements will

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be taken for each of these training sessions; a similar number of measurements should be used in the actual experiment) and perform the actual experimental recording on day 6. By comparing the arterial pressures from each of the training days, the investigators can then determine when the heart rate and arterial pressure measurements have reached a stable level. Usually, the results from the last 3 days of recording are relatively consistent, and all of these can be used in the final data analysis. If excessively high variation in blood pressure, heart rate, or movement is noted, additional training sessions will be needed. The actual number of training sessions required for acclimation will vary depending on the strain, gender, and age of the mice. IMPORTANT NOTE: Take care not to place mice of different gender near one another during training and recording period, as proximity to mice of the opposite sex could lead to excitation in some of the animals.

2. Adjust the heating mechanism on the platform by clicking on the “Adjust Temperature” option under the “Configuration” menu. Check the temperature of the platform to ensure a stable temperature of 38◦ to 40◦ C depending on the level of anesthesia (if used) and condition of the animal. 3. Prepare the tail cuff by inspecting, cleaning with alcohol, and calibrating the tail-cuff apparatus. The tail-cuff material may degrade and become brittle over time. To replace the balloon, as per manufacturer’s instructions, first remove the old balloon, cut 4 pieces of 1-1/3-in. mouse tail-cuff material (Visitech, BP-CE-M-38-100), thread the balloon through the tail cuff, and fold back the ends to wrap around the end of tail cuff. Use the tail-cuff balloon cap to hold the balloon in place, to ensure proper inflation of the balloon (Fig. 2).

Figure 2 Illustration of the tail-cuff system (A). To replace the tail-cuff balloon, the tail-cuff material is threaded through the tail-cuff device (B), the ends are then everted over the ends (C), and the balloon caps are then placed on to each of the two ends (D,E). The mice are restrained individually during the recording process while the tail-cuff system is connected to a computer for data acquisition (F).

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4. Connect the pressurizing tubing to the platform, then check the balloons for leakage with the “Pressure Leak Wizard,” found under the “Configuration” menu. 5. Load the animals onto the platform for acclimation while the system is being calibrated. a. Place the mouse inside one of the magnetic restrainers. b. Pass the tail through the cuff and pulse sensor. Make sure the pulse sensor holds the tail within the groove. Taping the distal part of the tail onto the platform will help avoid excessive movements.

6. Calibrate the system. Connect the platform to the sphygmomanometer, then select “Calibrate.” Pressure” from the “Configuration” menu, and follow the on screen directions. Ideally, four pressure inputs should be performed for this calibration setup. The system will ask to verify the pressure calibration by re-pressurizing.

7. Obtain arterial pressure measurements as follows: a. Re-connect the pressurizing tubing to the platform. b. Using the “Specimen Registration” option from the “Analysis” menu, enter the experimental groups and animal IDs used for data acquisition and analysis. c. Start recording. A user-entered number of measurements (1 to 30) will be used to allow acclimation of the animal to the balloon inflation and deflation. These acclimation measurements are done automatically by the tail-cuff apparatus, and are performed during each of the training sessions as well as prior to the experimental arterial pressure recording. IMPORTANT NOTE: The recording system should be set up in a quiet room with minimal through traffic to ensure the reproducibility of the obtained arterial pressure measurements.

d. Record and analyze an additional, user-determined number of measurements. This number should not be less than 20.

e. After the data acquisition is finished, to view the results, look under the “Report” option within the “Data” menu. According to the individual needs, “Measurement Set Statistics” or “Individual Measurements” is selected and exported to an Excel or text file for further analysis.

FLUID-FILLED CATHETER SYSTEM The use of indwelling fluid-filled catheters implanted into the aorta through either the femoral or carotid artery results in direct and continuous measurements of arterial pressure. The catheter is connected to a strain-gauge manometer for transduction of the pressure signal to an electrical signal, whereupon it is observed and stored using analog or digital recorders. When these catheters are exteriorized and secured, they can be tethered to a swivel device on the cage to allow free movement of the animals. In this way, the tethered system is used for monitoring of arterial pressure in a conscious animal (Fig. 3). Advantages and disadvantages of the fluid-filled catheters The advantage of this system is that it provides direct and continuous measurements of arterial pressure. In addition, it allows not only for monitoring of arterial pressure, but also for vascular access, thus permitting more experimental flexibility (Mattson, 1998). One example is the delivery of phenylephrine or sodium nitroprusside in a conscious

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Figure 3 Schematic diagram of the tether system. After the arterial line has been placed in the mouse, the line is inserted into the catheter harness (1), threaded up the spring casing (2) for the catheter, and then attached to the swivel device (3) on the swivel arm (4). The catheter tubing (5) exits from the spring casing and is attached to the pressure transducer (6) set at the level of the heart.

mouse, leading to a corresponding increase or decrease in arterial pressure (Fig. 4). The disadvantages under such a system, aside from morbidity and mortality associated with surgery, are that the catheters require a heparin lock and daily flushing to prevent clotting of the catheter. This system requires single housing of the animal and is not suited for continuous prolonged recording, as catheter patency diminishes and signal dampening occurs with time. Although flushing of the catheter will restore signal intensity, due to the low total blood volume of a mouse, frequent flushing may result in volume overload. Finally, when the heart rate of the mouse is elevated, the arterial signal may be dampened (Mattson, 1998). No training period is required for this protocol.

Materials Mice Anesthetic of choice (see Table 1 and step 2, below) Heparinized saline solution: dilute heparin to 1 U/ml in 0.9% NaCl

Arterial Pressure Monitoring in Mice

Catheters PE-05 tubing (0.28 mm i.d × 0.61 mm o.d) for the carotid artery PE-08 tubing (0.20 mm i.d. × 0.36 mm o.d.) for the femoral artery Connect either catheter onto RPT-040 tubing (0.64 mm i.d. × 1.02 mm o.d.) and glue at the connection part with silicon glue Adapter: 23-G Intramedic luer stub adapter (Becton Dickinson, cat. no. 427565) Animal clippers Surgical instruments Suture: 7-0 Silk for vessel ligation; 6-0 nylon for skin closure Customized needle-wire plug: cut off the plastic part of a 23-G adapter, using a file to collapse one end completely Solid-state pressure transducer (Becton-Dickinson, cat. no. 682018) Signal amplifier (Triton Technology, System 6, model 200; http://www. physiology.com/)

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Figure 4 Arterial pressure recording with an implanted fluid-filled catheter showing an example of blood pressure change in response to vasoactive compounds in a conscious C57BL/6J mouse. An increase in both (A), the aortic pressure (AOP) and (B), the mean arterial pressure (MAP), and a decrease in (C), the heart rate (HR) are noted with the administration of phenylephrine (PE). (D) A decrease in AOP and (E) MAP with (F) an increased HR are noted with the administration of sodium nitroprusside (SNP).

Recording system: e.g., Power Lab (ADInstruments, http://www.adinstruments.com), Dataquest (Data Sciences International, http://www.datasci.com), NOTOCORD-hem (Notocord, http://www.notocord.com), EMKA IOX (EMKA Technologies) Sphygmomanometer (blood pressure cuff of appropriate size to fit transducer; available at pharmacies) 1-ml and 50-ml syringe Tether (Harvard Apparatus; counter-balanced lever arm, PY861-0023; swivel, PY 856-1324) PE-05 tubing for catheter extension on tether Prepare and insert catheters 1. Calibrate the fluid-filled catheters as follows. Connect a 10-ml syringe filled with saline to the bottom of the transducer and flush the air out. Connect the transducer to the pressure amplifier. Connect the amplifier to a stripchart and to a computerized data acquisition system. Connect a sphygmomanometer to the top of the transducer. Run both the stripchart and the acquisition system at slow speed. With the pressure reading at 0, adjust the amplifier offset so the reading on the stripchart is at the bottom of the grid, then set the acquisition system in calibration mode to 0. Using a 50-ml syringe, set the pressure on the sphygmomanometer to the desired high point. Adjust the amplifier gain to set the stripchart to the top of the grid and set the acquisition system to the pressure reading on the sphygmomanometer. 2. Anesthetize mouse, shave, and place in supine position. Depending on experimental design and the experience of each individual laboratory, the choice of anesthesia will vary; please refer to Table 1 for reference on the hemodynamic

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distal suture (1st suture) middle suture (3rd suture) proximal suture (2nd suture)

Figure 5 Illustration of the suture placements for the three suture technique used during carotid artery catheter insertion. The sutures are placed around the right carotid artery. The numbers indicate the sequence of suture placement.

effects of each of the anesthetic agents. Different anesthetic agents may be used for the day of the surgery and the day of the experiment. In our laboratory, we generally use a mixture of ketamine and xylazine for catheter implantation. This mixture has a shorter induction time and a more rapid recovery time when compared to pentobarbital; however, any anesthetic agent that the individual surgeon is comfortable with could be used. As for recording, the conscious condition is preferred. If needed, Avertin anesthesia is used as the agent of choice for hemodynamic monitoring under sedation because of its lower suppressive effects on cardiac function, heart rate, and temperature when compared to pentobarbital or ketamine/xylazine mixture.

3. Make a 1- to 2-cm midline neck incision from just below the mandible to the thoracic inlet. Under a dissecting microscope, expose the right carotid artery and carefully separate from neighboring structures, including the vagus nerve. 4. Once the carotid artery has been isolated, place a silk suture (7-0 or 6-0) distally (closer to the head) for the complete ligation of the vessel. Place a second silk suture proximally (closer to the heart) to allow temporary obstruction of blood flow. Finally, place a third silk suture loosely between the first two ligatures and make a small incision (arteriotomy) distal to the middle ligature (i.e., between the first and the third suture) These procedures are illustrated in Figure 5.

Arterial Pressure Monitoring in Mice

Optional: Due to the smaller diameter of the femoral artery as compared to the carotid artery, a smaller-caliber catheter (PE-08 tubing) is needed for the femoral artery. Due to the anatomical relationship between the femoral artery and the abdominal aorta, it is technically more difficult to advance the femoral catheter beyond the bifurcation point into the abdominal aorta. Once the femoral catheter is successfully inserted into the abdominal aorta, the caliber of the catheter remains a point of consideration. With the smaller catheter used for femoral artery, there is a decrease in signal wave form quality and intensity of pressure. Furthermore, because of the location and associated potential movement of the legs, the femoral catheter is more prone to signal loss during movement. For similar reasons, the insertion of the femoral artery catheter requires a more secure fixation of the surrounding musculature compared to what is needed for the insertion of carotid artery catheters. Although not preferred, either the left or the right femoral artery

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can still be used in place of the carotid. Using the three-suture technique in a similar fashion to the one described above for the carotid artery, the femoral vessels are exposed and the artery isolated for cannulation through a 1- to 2-cm inguinal skin incision.

5. Insert the tip of a catheter that has been pre-filled with 10% to 30% heparinized saline into the carotid artery via the arteriotomy in the direction of the heart and secure it in place by tying the mid (i.e., the third) suture once that catheter has been advanced past the ligature. 6. Release the proximal ligature and re-ligate after the catheter has been advanced about 11 to 12 mm into the ascending aorta or 18 to 20 mm into the left ventricular chamber. When advancing or withdrawing the catheter, hold the catheter between the thumb and index finger; use a gentle twisting motion in conjunction with the slow advancement of the catheter to help avoid vascular injury. Pay special attention to the resulting waveform to ensure the proper placement of the catheter. If using the femoral artery: Upon isolation of a section of the femoral artery, apply a couple of drops of 2% lidocaine to the area to help dilate the vessel. Insert the catheter and advance it past the bifurcation point into the abdominal aorta. If resistance is felt, do not force the catheter; re-position the mouse to line up the operated leg with the axis of the body. Using a twisting motion, move the catheter forward to advance the catheter beyond the bifurcation.

7. Verify cannula patency by blood return via withdraw as described below, then flush with a little more heparinized saline and cap the distal end of the catheter with a customized needle-wire plug. To verify catheter patency, once the heparinized-saline-prefilled catheter has been advanced to the proper position, the open end of the catheter is attached to a 1-ml syringe filled with 0.3 to 0.6 ml heparinized saline. Next, check blood return by withdrawing the plunger; blood should be seen entering the catheter. Once blood is seen, the plunger is pushed back to clear blood from the catheter. If no blood return is seen, the catheter needs to be repositioned prior to rechecking for blood return. With the low total blood volume of a mouse (80 ml/kg body weight), caution should be used when flushing the catheter, to avoid excessive volume loading.

8. Place the mouse in the right lateral recumbent position (i.e., right side down), and subcutaneously tunnel and externalize the catheter through a mid-scapular skin incision on the back. If using the femoral artery, secure the catheter in place through the application of sutures, again tunnel subcutaneously, and externalize in the midscapular region on the back.

9. Close the neck incision with 6-0 nylon sutures and fix the external portion of the catheters in the back to the underlying muscle. For both the carotid and femoral catheters, once they have been exteriorized in the midscapular region, protect the catheters from being chewed by the animal by housing the catheter within a plastic cap transfixed to the midscapular region of the dorsal skin.

10. Unless immediate post-anesthesia measurements are to be performed, allow the animals to recover inside a pre-warmed (31◦ C) rodent cage after the surgery.

Record arterial pressure 11. Connect the catheter to a solid-state pressure transducer placed at the level of the mouse heart. 12. Connect the pressure transducer to a signal amplifier that is connected to the recording system of choice.

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Additional steps for tether The exteriorized catheter can be attached to the tether device for conscious recording of freely mobile animals. 13. First, thread the catheter through the harness and spring casing of the tether. The spring casing is connected to a swivel device which is connected to the swivel arm attached to the side of the cage.

14. Once the catheter exits from the spring, attach the casing to a section of PE-05 tubing, which is in turn attached to the pressure transducer placed at the level of the mouse heart. 15. Attach the pressure transducer to a signal amplifier connected to the recording system (Fig. 3). BASIC PROTOCOL 3

MILLAR SOLID-STATE MICRO-PRESSURE TRANSDUCER TIPPED CATHETER Solid-state Millar catheters provide the frequency response needed for assessing alterations in the phasic arterial pressure waveform. In brief, the Millar solid-state catheter contains a micro-pressure transducer mounted at its distal end, allowing the conversion of the pressure wave into an electrical signal. The procedure used for implantation of a Millar catheter is similar to that used for a fluid-filled catheter. Advantages and disadvantages of the Millar solid-state catheters The most important advantage of the Millar solid-state catheter system is its frequency response, making it the most sensitive and precise method for the monitoring of arterial pressure and for analysis of changes in the phasic arterial pressure waveform. In addition, unlike the fluid-filled catheters, Millar solid-state catheters do not require flushing and do not have the problem of decreasing signal intensity with increasing catheter length. One main disadvantage is the cost, and there are other drawbacks including surgical morbidity and mortality. Furthermore, due to its extreme sensitivity, the recordings are affected by ambient conditions such as temperature and blood viscosity or electrolyte change (Lorenz, 2002), so multiple parameters must be closely monitored and controlled. Additionally, we have noted electrical interference from other surgical equipment during hemodynamic measurements when utilizing this system. Finally, at present, Millar solid-state catheters are not convenient for conscious monitoring of arterial pressure in mice, although a mouse telemetry system utilizing the Millar solid-state catheter is in development (ADInstruments, Inc.). Therefore, Millar solid-state catheters are best suited for acute studies under anesthesia where the monitoring of arterial pressure needs to be coupled to left ventricular pressure measurements. No training period is required for this protocol.

Materials Micromanometer catheter: 1.4F (Millar Instruments, http://www.millarinstruments.com/) Suture: Silk 7-0 or 6-0 for vessel ligation Additional reagents and equipment for implantation of fluid-filled catheter (Basic Protocol 2) Arterial Pressure Monitoring in Mice

1. Implant the Millar solid-state catheter following the procedure described in detail for the implantation of fluid-filled catheters (see Basic Protocol 2). Either the carotid or the femoral artery can be employed as vascular access.

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In smaller mice, catheter advancement into either the carotid or the femoral artery can be difficult. Application of an acoustic gel around the catheter entry site will provide lubrication. IMPORTANT NOTE: Throughout the implantation process, when the Millar solid-state catheter is not inside the animal, the tip of the catheter needs to be kept in warm saline (37◦ C) at all times.

2. Subject the Millar solid-state catheter to a cleaning process as described in the instructional manual. Keep the tip of the catheter protected at all times. 3. Turn on the recording instruments prior to connecting the catheter for calibration (warm-up time, 20 to 30 min). This is important because the Millar catheter is sensitive to sudden electrical surge.

4. Keep the vessel(s) around the catheter wet before and during implantation. If a shift in baseline or abnormal wave form, such as a spike noted on the top of the pressure wave, is observed, try repositioning the body of the animal or the catheter tip, as the shift may be the result of catheter position and the change in waveform may be the result of the catheter sensor coming into contact with the vessel or ventricular wall.

IMPLANTED TELEMETRY SYSTEM Radiotelemetry units have been accepted as the gold standard for the monitoring of arterial pressure in intact, conscious mice. These devices contain a short segment of catheter attached to a signal transmitter. The catheters function to sense pressure waves in a manner similar to the fluid-filled catheter. The catheter is inserted either into the carotid artery with subcutaneous placement of the transmitter or into the abdominal aorta with intraperitoneal placement of the transmitter. Prior to insertion into the animal, the catheters are pre-filled with a gel material (AD instruments) to improve signal conduction and help prevent clotting of the catheter. The unit functions similarly to a fluid-filled catheter, the difference being that the electrical signal is transmitted wirelessly to a receiving device. The units are activated via a magnet and the transmitted signals are then picked up by a receiver placed on the outside of the animal cage, and the data recorded by a connected computer.

BASIC PROTOCOL 4

Advantages and disadvantages of the implanted telemetry device The major advantages of implanted telemetry systems include remote monitoring of arterial pressure in conscious mice living in their natural environment while providing sensitive, accurate, long-term data (Fig. 6; Kramer et al., 2000; Desjardins et al., 2008). On the other hand, disadvantages include the cost and the surgical skill needed for implantation, with consequent surgically related morbidity and mortality. Furthermore, a 5- to 7-day recovery period is often needed after implantation (Lorenz, 2002). In the event of an acute study where intravenous (i.v.) drug delivery is needed, the insertion of an i.v. catheter is still required. When the gel-filled sensing catheters are implanted into the abdominal aorta, cases of vascular occlusion leading to decreased blood flow to the hindquarters have been reported (Lorenz, 2002). This is avoided by using the carotid artery, where ischemia is much less of an issue. With the use of newer techniques and models, these devices are implanted into mice as small as 17 g with a reported success rate of 90% (Carlson and Wyss, 2000). Telemetry systems are best suited for long-term continuous monitoring of mice either within their natural living environment or with exercise. No training period is required for this protocol.

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Figure 6 An example of telemetry blood pressure recording over a 24-hr time period. The circadian variation in arterial pressure and heart rate are clearly observed in this recording. (A) Arterial systolic pressure (SBP) and (B) arterial diastolic pressure (DBP) variations are seen in both the wild-type (WT, C57BL/6) and caveolin-1 knockout (KO) mice. The shaded and nonshaded areas on the x axis represent the dark and light cycles, respectively (Desjardin et al., 2008). Figure reproduced and modified with permission.

Materials Telemetry transmitter device (PA-C20 or PA-C10) (Data Sciences International, http://www.datasci.com/; also see Fig. 7) Recording system: e.g., Power Lab (ADInstruments, http://www. adinstruments.com), Dataquest (Data Sciences International, http://www.datasci.com), NOTOCORD-hem (Notocord, http://www.notocord.com), EMKA IOX (EMKA Technologies) Suture: 7-0 Silk for vessel ligation, 6-0 nylon for skin closure, 5-0 nylon for telemetry device fixation Additional reagents and equipment for implantation of fluid-filled catheter (Basic Protocol 2) 1. Anesthetize, shave, and place mouse in the supine position. Depending on experimental design and the experience of each individual laboratory, the choice of anesthesia will vary; please refer to Table 1 for reference on the hemodynamic effects of each of the anesthetic agents. Different anesthetic agents may be used for the day of the surgery and the day of the experiment. In our laboratory, we generally use a mixture of ketamine and xylazine for surgery; however, any anesthetic agent that the surgeon is comfortable with could be used for telemetry implantation. Recording is generally done under conscious conditions; however, if anesthesia is needed, Avertin is used for hemodynamic monitoring under sedation.

2. Make a 2-cm mid-cervical incision in the neck of the mouse and create a subcutaneous space by undermining the skin on the right chest to create a pocket large enough to accommodate the device. 3. Insert the catheter tip via an arteriotomy in the left carotid artery and advance it into the transverse aorta for about 13 mm, following the three-suture placement technique described in Basic Protocol 2, step 4, for fluid-filled catheter implantation. 4. Perform extra fixation of the catheter onto muscle and soft tissue of the mouse to prevent the dislodgement of the catheter tip from the arterial lumen, which could lead to inaccurate readings. Arterial Pressure Monitoring in Mice

Perform extra fixation by placing a suture through the muscle and soft tissue and tying this suture around the catheter to prevent the dislodgement of the catheter tip from the arterial lumen, which could lead to inaccurate readings.

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preformed holes for anchoring

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gel-filled pressure catheter

anchoring suture (blue)

Figure 7 Illustration of the telemetry unit with the anchoring sutures in place. In general, two to three anchoring sutures are used to fix the telemetry unit to the chest wall of the mice.

recording computer connected to data acquisition system

implanted telemetry unit receiver plate

connection to data acquisition system

Figure 8 Illustration of a telemetric recording of mouse arterial pressure. The mice are housed individually during the recording process, with the recording receiver placed below each cage and connected to a computer for data acquisition.

5. Insert the transmitter device into the pocket in the right chest with the side containing the predrilled suture holes facing up, and transfix the device to the overlying skin using a few 5-0 nylon sutures to prevent migration (Fig. 7). Due to the potential irritation to the mouse associated with the implanted telemetry device, the body of the device should be secured to prevent migration of the device and keep the mouse from scratching and biting the site of surgery or device.

Recording 6. After inserting the telemetry unit, allow the mice to recover for at least 5 to 7 days. 7. Once the animals have recovered from the insertion of the telemetry unit, house them individually in a regular mouse cage placed on top of the telemetry receiver plate (Fig. 8). Current Protocols in Mouse Biology

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a. If the recording of multiple animals is needed, each plate should be placed at least 17 inches from each other. When fully charged, the battery will allow for 1.5 months of continuous recording. If the arterial pressure waveform becomes dampened or the signal is lost, the device should be removed.

b. If telemetric recording is desired during exercise, the receiver plate should be placed above the treadmill. c. The device can be turned on/off with magnetic activation and can be used in combination with drug administration should the experimental need arise. d. After the device is removed from the animal, it should be sent back to the manufacturer for cleaning, reconditioning (insertion of new battery and catheter, and recalibration), and sterilization.

COMMENTARY Background Information In cardiovascular research and the development of new pharmacological agents, arterial pressure monitoring is of particular importance for evaluating cardiovascular function and response, such as change in arterial pressure in response to vasoactive drugs and change in vessel stiffness in aged and hypertensive animal models. Mice are an important animal model, and are widely used in laboratory and clinical research. Arterial pressure monitoring in mice is challenging due to the size of the animal and the rapid heart rate. This article reviews the four major methods for arterial pressure monitoring in mice: tail cuff, intravascular fluid-filled catheter (including the tether system), solid-state Millar catheter, and telemetry. Tail-cuff system This method (Basic Protocol 1) utilizes a sphygmomanometer coupled to blood-flow measurement in the tail artery via photoplethysmography or piezoplethysmography. In general, the tail cuff is inflated to occlude the blood flow, and the disappearance of arterial pressure during inflation or the first appearance of the pressure wave during deflation is taken as the systolic pressure; the diastolic pressure is determined by mathematical calculation. Since flow cannot be quantified by either of these methods, diastolic pressure is determined by mathematical calculation. However, employing a method that uses tail volume as a measure of blood flow into the tail, diastolic pressure is determined as the pressure upon which the blood flow into and out of the tail are equalized. Arterial Pressure Monitoring in Mice

Fluid-filled catheter system This method (Basic Protocol 2) uses an implanted indwelling fluid-filled catheter, con-

nected to a strain-gauge manometer, in either the aorta, the femoral artery, or carotid artery for direct and continuous measurements of arterial pressure. The pressure signal is then converted to an electrical signal, whereupon it is observed and stored using analog or digital recorders. Millar solid-state micro-pressure-transducer-tipped catheter This instrument, used in Basic Protocol 3, Consists of a micro-pressure transducer mounted at the distal end of a wire catheter converting the measured pressure wave into an electrical signal, which is then recorded in a manner similar to the signal obtained via the fluid-filled catheter. Implanted telemetry systems Implanted telemetry systems (Basic Protocol 4) are accepted as the gold standard for monitoring of arterial pressure in intact, conscious mice. These devices consist of a short segment of catheter attached to a signal transmitter; catheters are inserted into either the carotid artery or the abdominal aorta, after which the pressure-wave signals are then converted to radio waves to be received by a signal receiver. For detailed background on each of the individual methods please refer to Basic Protocols 1 to 4. Advantages and disadvantages The tail-cuff system is the simplest method for measuring arterial pressure within multiple animals; there is no need for anesthesia or invasive procedures. However, the obtained arterial pressure measurement may be influenced by the stress-associated balloon inflation and deflation and the peripheral arterial vasoconstriction. Fluid-filled catheters were widely used for measurement of central arterial pressure

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prior to the development of solid-state Millar catheters. This method is used for continuous monitoring of central arterial pressure in both conscious and anesthetized animals. Based on the skill of the surgeon, the less the blood loss during catheter insertion and the better the catheter patency, the more physiologic the results. The problem most often encountered concerns catheter patency; when the catheter is partially clogged, the pressure waveform may be dampened and the pulse pressure will be decreased. An additional drawback, aside from the surgically associated morbidity and mortality, is that this system requires a heparin lock and daily flushing of the catheter to prevent clotting. Furthermore, due to the need for continuous monitoring of catheter patency and signal quality, this system is not suited for continuous prolonged recording. Compared to the fluid-filled catheter, the Millar solid-state catheter provides increased sensitivity and responsiveness. The Millar catheter requires a financial commitment, and the recording may be affected by physiologic as well as ambient conditions, so much care must be taken to minimize these influences. Finally, the telemetry system allows for the recording of arterial pressure in conscious animals within their normal living environment, providing the most physiologic of the arterial pressure readings. However, since it is an implanted intravascular catheter, careful calibration and sterilizing of the device prior to implantation is of particular importance. For details of the advantages and disadvantages of each individual method, please refer to Basic Protocols 1 to 4.

Critical Parameters Invasive versus noninvasive Due to the stress, morbidity, and mortality associated with invasive methods of arterialpressure monitoring, noninvasive methods are preferred whenever possible. However, the tail-cuff arterial pressure monitoring system is not stress free, and there is some level of stress associated with the use of a restrainer. The noninvasive tail-cuff arterial pressure monitoring method allows for rapid, periodic monitoring of arterial pressure of experimental animals, thus allowing for the monitoring of arterial pressure change over time. However, it is important to remember that the tail-cuff system provides information only on peripheral arterial pressure, as compared to the central arterial pressure obtained by other invasive techniques, such as the fluid-filled catheter, telemetry, and the Millar catheter. If poor data

fidelity or an unexpected result is noticed, either due to the physical condition of the animal or lack of acclimation of the animal to the tail-cuff device, resulting in missing data points during the recording, the investigator should consider invasive methods for monitoring of arterial pressure. It is generally recommended that indirect methods of arterial pressure monitoring (e.g., tail cuff) not be used to measure blood pressure variability, diastolic blood pressure, or pulse pressure in conscious mice. In addition, this technique is not suited for studies in nonstressed, unrestrained mice, nor is it well suited for drug studies or the detection of mild or intermittent hypertension (Kurtz, 2005). Direct versus indirect Comparing the results of direct invasive arterial pressure monitoring techniques to those of indirect arterial pressure measurements in the peripheral artery may result in different results due to the condition of the animal, a low peripheral temperature, or difference in regional effects of vasoconstrictive compounds (e.g., phenylephrine). In drug dose-response experiments, a direct measurement is preferred over indirect methods. Direct methods allow for continuous monitoring of arterial pressure change, whereas tail cuff provides only intermittent arterial pressure readings, such that if the duration of the response is short, the drug response could be missed completely. Furthermore, the observed central arterial pressure effect could be very different from that of the peripheral arterial pressure. Thus, depending on the area of interest, the desire for peripheral versus central pressure monitoring or continuous versus intermittent arterial pressure monitoring should be considered in the experimental design. Need for telemetry The implanted telemetry system is ideal for long-term continuous monitoring of conscious arterial pressure, and is the gold standard for this purpose. For chronic studies, such as monitoring chronic vasoactive drug infusion or circadian arterial pressure change, heart-failure studies, or comparison of phenotypic arterial blood pressure difference between genetically modified animals, telemetry is preferred over other methods. Although the manipulation of the carotid artery during catheter insertion may injure the ipsilateral carotid baroreflex center, this is well compensated by the baroreflex center on the contralateral carotid and the aortic arch.

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Need for high-fidelity waveform The solid-state Millar catheter system provides the highest fidelity of the arterial pressure waveform with the most rapid response time of all the methods. In mice under stimulation, heart rate could be as high as 700 beats per minute (bpm), while under resting conditions, mice generally have a heart rate of 500 to 600 bpm. Therefore, to achieve a sufficient frequency-response rate for mouse LV blood pressure monitoring with dP/dt, a system capable of sampling at a rate of 1000 to 2000 Hz may be needed (Lorenz and Robbins, 1997). In order to achieve this goal, the Millar catheter is the most suitable method. For anesthetized drug-response experiments, the Millar solidstate catheters provide the most accurate measurement among all techniques. The experimental needs and design of a particular study will dictate the optimal method required for monitoring arterial pressure for that setting. The material discussed in this article will aid in selecting the technique that is best suited for any distinct application.

Troubleshooting Tail-cuff system Even after multiple training sessions, mice may still be agitated during the process of pressurizing the tail cuff. This will in turn affect the accuracy of the pressure readings. Taping the distal part of the tail on the platform may help avoid excessive movement from the animal. Although the restrainer is made of metal, there are predrilled holes through which ambient light will leak in, and the animals will be able to sense nearby movements. Using a black cover on the outside of the platform and the restrainer will help reduce light. In addition, a quiet room and a calm recording operator are indispensable to reduce noise level and to decrease the agitation experienced by the animal.

Arterial Pressure Monitoring in Mice

Fluid-filled catheter system Due to the rapid blood flow at the entrance of the ascending aorta, at times it is difficult to advance the catheter into the left ventricle. Moreover, forcefully advancing the catheter may result in aortic valve damage. While advancing the catheter, place the catheter between the thumb and index fingers, using a gentle twisting motion in conjunction with the slow advancement of the catheter to place the catheter into the left ventricle. The entrance of the catheter into the left ventricle is often felt as a sudden disappearance of resistance, and

to-and-fro movement of blood flow inside the catheter will be visibly intensified. During femoral artery catheter implantation, it may be difficult to advance the catheter beyond the aortic bifurcation of the iliac artery. If resistance is felt, reposition the mouse to line up the operated leg with the longitudinal axis of the body and twist the catheter forward; this body position will help to advance the catheter beyond the bifurcation point. Millar solid-state micro-pressuretransducer-tipped catheter After the Millar catheter is implanted into the left ventricle, sometimes spike artifacts are observed on the top of the pressure wave, as seen on the monitor or paper strip chart; this is at times associated with arrhythmia. One possible explanation for this observation is left ventricular irritation secondary to contact between the catheter sensor and the ventricular wall. This is remedied by repositioning the catheter tip via a change in direction or depth. Alternatively, the contact between the catheter and LV wall may be secondary to pharmacological stimulation, resulting in severely contracting ventricles. In this case, the spike artifact may persist until the drug effect has worn off. Occasionally, an upward or downward drift of the waveform may be observed, and this could lead to inaccurate pressure readings. This could be secondary to the sudden repositioning of the animal or catheter. Therefore, by repositioning the body of the animal or the catheter tip, stabilization of the waveforms may be observed. Due to tight fitting of the catheter in the vessel, catheter advancement in either the carotid or the femoral artery can be difficult. Application of an acoustic gel around the catheter entry site may serve as helpful lubricant. Application of 2% lidocaine is almost always necessary in femoral arterial catheter insertion. Although the catheter may be pushed out under the conditions of normal arterial pressure within the carotid artery, the chance of this occurring increases with increasing arterial pressure. In mice with transverse aortic banding, the right carotid artery is usually bulging and dilated, making catheter insertion and advancement easier; however, care should be taken to avoid having the catheter pushed out due to the high pressure (hypertensive) present in the carotid artery. Implanted telemetry system Due to the chronic nature of the implanted telemetry system, the mice may chew on

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Table 2 Mean Arterial Pressure (MAP) and Heart Rate (HR) in Three Strains of Conscious Micea

Strain

C57/B6SJL

129Sv/J

FVB

MAP (mmHg)

103 ± 1

102 ± 8

99 ± 2

HR (bpm)

588 ± 14

522 ± 11

617 ± 26

Data source

Uechi et al. (1998)

Gross and Luft (2003)

Lin et al. (2010)

a All values are mean ± SE. (MAP=mean arterial pressure; HR=heart rate).

the surgical sites due to associated irritation. Therefore, the body of the device should be secured properly in the pocket to prevent migration of the device. Movement may lead to the migration of the implanted catheter, resulting in the dislodgement of the catheter tip from the arterial lumen, thus leading to massive lethal bleeding. Extra fixation of the catheter onto the musculature and soft tissue of the mouse is necessary to help decrease the possibility of such an event. The battery in the telemetry device lasts for 1.5 months with continuous use. However, if a damped arterial pressure waveform or a loss of signal is noticed, the device may have lost its efficacy. At this point, the device should be removed and cleaned, then sent back to the manufacturer for reconditioning.

Anticipated Results The anticipated result for the tail-cuff system is a digital readout of a series of numbers representing each individual reading and the average arterial pressures, while the fluid-filled catheters, Millar solid-state catheters, and the telemetry system will provide arterial pressure wave forms in addition to the digital readout of the measured arterial pressures. Multiple variables are known to affect the measured arterial pressure, including the experimental and physiologic conditions, the level of sedation, volume status, age, and gender, as well as many environmental factors. As a general reference, the average conscious arterial pressures found in some common strains of mice used for cardiovascular research can be found in Table 2. In general, hypertension can be defined as blood pressure greater than two standard deviations above the mean.

Time Considerations The time required for the tail-cuff system of arterial pressure monitoring depends upon the number of acclimation and actual readings set by the investigator, but, in general, expect 1 to 2 hr. As for the other methods, the actual time

required for the completion of the method will depend on the skill of the surgeon. The time required for the completion of each of these methods is found below. Tail-cuff system: Depends on the number of acclimations and actual readings desired for each animal. On average, expect 1 to 2 hr to complete the process, allow 5 min for preparation, and allow 5 min for calibration of the device. This process is repeated for multiple days to reduce variation on different days. Fluid-filled catheters: On average, depending upon the skill of the surgeon, without tether, allow 30 to 40 min to complete the process. If tether is needed, add an additional 5 to 10 min for tether placement. Prior to recording, the time required for calibration and preparation is ∼5 min for each. Millar solid-state catheters: On average, depending upon the skill of the surgeon, allow 25 to 35 min to complete the process. Similar to the fluid-filled catheter and tail-cuff systems, prior to recording, the time required for calibration and preparation is ∼5 min for each. Telemetry system: On average, depending upon the skill of the surgeon, expect 30 to 40 min to complete the process. The recording process can be initiated with ∼5 min of preparation time.

Acknowledgements This work was supported in part by NIH grants HL033107, HL069020, AG027211, HL101420, HL093481, HL059139, HL095888, DK083826, and HL102472.

Literature Cited Carlson, S.H. and Wyss, J.M. 2000. Long-term telemetric recording of arterial pressure and heart rate in mice fed basal and high NaCl diets. Hypertension 35:E1-E5. Desjardins, F., Lobysheva, I., Pelat, M., Gallez, B., Feron, O., Dessy, C., and Balligand, J.L. 2008. Control of blood pressure variability in caveolin1-deficient mice: Role of nitric oxide identified in vivo through spectral analysis. Cardiovasc. Res. 79:527-536.

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Doevendans, P.A., Daemen, M.J., de Muinck, E.D., and Smits, J.F. 1998. Cardiovascular phenotyping in mice. Cardiovasc. Res. 39:34-49.

ponents of the baroreceptor reflex in mice. Am. J. Physiol. Regul. Integr. Comp. Physiol. 283:R1033-R1040.

Feng, M., Whitesall, S., Zhang, Y., Beibel, M., D’Alecy, L., and DiPetrillo, K. 2008. Validation of volume-pressure recording tail-cuff blood pressure measurements. Am. J. Hypertens. 21:1288-1291.

Mattson, D.L. 1998. Long-term measurement of arterial blood pressure in conscious mice. Am. J. Physiol. 274:R564-R570.

Feng, M., Deerhake, M.E., Keating, R., Thaisz, J., Xu, L., Tsaih, S.W., Smith, R., Ishige, T., Sugiyama, F., Churchill, G.A., and DiPetrillo, K. 2009. Genetic analysis of blood pressure in 8 mouse intercross populations. Hypertension 54:802-809. Gross, V. and Luft, F.C. 2003. Exercising restraint in measuring blood pressure in conscious mice. Hypertension 41:879-881. Hart, C.Y., Burnett, J.C. Jr., and Redfield, M.M. 2001. Effects of avertin versus xylazineketamine anesthesia on cardiac function in normal mice. Am. J. Physiol. Heart Circ. Physiol. 281:H1938- H1945. Janssen, B.J., De Celle, T., Debets, J.J., Brouns, A.E., Callahan, M.F., and Smith, T.L. 2004. Effects of anesthetics on systemic hemodynamics in mice. Am. J. Physiol Heart Circ Physiol 287:H1618-H1624. Kiatchoosakun, S., Kirkpatrick, D., and Hoit, B.D. 2001. Effects of tribromoethanol anesthesia on echocardiographic assessment of left ventricular function in mice. Comp. Med. 51:26-29. Kramer, K., Voss, H.P., Grimbergen, J.A., Mills, P.A., Huetteman, D., Zwiers, L., and Brockway, B. 2000. Telemetric monitoring of blood pressure in freely moving mice: A preliminary study. Lab. Anim. 34:272-280. Krege, J.H., Hodgin, J.B., Hagaman, J.R., and Smithies, O. 1995. A noninvasive computerized tail-cuff system for measuring blood pressure in mice. Hypertension 25:1111-1115. Kurtz, T.W., Griffin, K.A., Bidani, A.K., Davisson, R.L., and Hall, J.E. 2005. Recommendations for blood pressure measurement in humans and experimental animals. Part 2: Blood pressure measurements in experimental animals. Hypertension 45:299-310. Lin, M., Harden, S.W., Li, L., Wurster, R.D., and Chen, Z. 2010. Impairment of baroreflex control of heart rate in conscious transgenic mice of type 1 diabetes (OVE26). Auton. Neurosci. 152:6774. Lorenz, J.N. 2002. A practical guide to evaluating cardiovascular, renal, and pulmonary function in mice. Am. J. Physiol. Regul. Integr. Comp. Physiol. 282:R1565-R1582. Lorenz, J.N. and Robbins, J. 1997. Measurement of intraventricular pressure and cardiac performance in the intact closed-chest anesthetized mouse. Am. J. Physiol. 272:H1137-H1146. Arterial Pressure Monitoring in Mice

Ma, X., Abboud, F.M., and Chapleau, M.W. 2002. Analysis of afferent, central, and efferent com-

Odashima, M., Usui, S., Takagi, H., Hong, C., Liu, J., Yokota, M., and Sadoshima, J. 2007. Inhibition of endogenous Mst1 prevents apoptosis and cardiac dysfunction without affecting cardiac hypertrophy after myocardial infarction. Circ. Res. 100:13441352. Pena, J.R. and Wolska, B.M. 2005. Differential effects of isoflurane and ketamine/inactin anesthesia on cAMP and cardiac function in FVB/N mice during basal state and beta-adrenergic stimulation. Basic Res. Cardiol. 100:147153. Roth, D.M., Swaney, J.S., Dalton, N.D., Gilpin, E.A., and Ross, J. Jr. 2002. Impact of anesthesia on cardiac function during echocardiography in mice. Am. J. Physiol. Heart Circ. Physiol. 282:H2134-H2140. Szczesny, G., Veihelmann, A., Massberg, S., Nolte, D., and Messmer, K. 2004. Long-term anaesthesia using inhalatory isoflurane in different strains of mice-the haemodynamic effects. Lab. Anim. 38:64-69. Uechi, M., Asai, K., Osaka, M., Smith, A., Sato, N., Wagner, T.E., Ishikawa, Y., Hayakawa, H., Vatner, D.E., Shannon, R.P., Homcy, C.J., and Vatner, S.F. 1998. Depressed heart rate variability and arterial baroreflex in conscious transgenic mice with overexpression of cardiac Gsα. Circ. Res. 82:416-423. van Nimwegen, C., van Eijnsbergen, B., Boter, J., and Mullink, J.W. 1973. A simple device for indirect measurement of blood pressure in mice. Lab. Anim. 7:73-84. Vatner, D.E., Yan, G.P., Geng, Y.J., Asai, K., Yun, J.S., Wagner, T.E., Ishikawa, Y., Bishop, S.P., Homcy, C.J., and Vatner, S.F. 2000. Determinants of the cardiomyopathic phenotype in chimeric mice overexpressing cardiac Gsα. Circ. Res. 86:802-806. Vatner, S.F. and Braunwald, E. 1975. Cardiovascular control mechanisms in the conscious state. N. Engl. J. Med. 293:970-976. Vatner, SF., Takagi, G., Asai, K., and Shannon, R.P. 2002. Cardiovascular physiology in mice: Conscious measurements and effects of anesthesia. In Cardiovascular Physiology in the Genetically Engineered Mouse (B.D. Hoit, ed.) pp. 257-275. Kluwer Academic Publishers, New York. Whitesall, S.E., Hoff, J.B., Vollmer, A.P., and D’Alecy, L.G. 2004. Comparison of simultaneous measurement of mouse systolic arterial blood pressure by radiotelemetry and tail-cuff methods. Am. J. Physiol. Heart Circ. Physiol. 286:H2408-H2415.

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Heart Rate and Electrocardiography Monitoring in Mice David Ho,1 Xin Zhao,1 Shumin Gao,1 Chull Hong,1 Dorothy E. Vatner,1 and Stephen F. Vatner1 1

The University of Medicine & Dentistry of New Jersey, New Jersey Medical School, Newark, New Jersey

ABSTRACT The majority of current cardiovascular research involves studies in genetically engineered mouse models. The measurement of heart rate is central to understanding cardiovascular control under normal conditions, with altered autonomic tone and with superimposed stress, or in disease states, both in wild-type mice and in those with altered genes. Electrocardiography (ECG) is the “gold standard,” using either hard-wired or telemetry transmission. In addition, heart rate is measured or monitored from the frequency of the arterial pressure pulse or cardiac contraction, or by pulse oximetry. For each of these techniques, discussions of materials and methods, as well as advantages and limitations, are covered. However, direct ECG monitoring alone will determine not only the precise heart rates but also whether the cardiac rhythm is normal or not. Curr. Protoc. Mouse C 2011 by John Wiley & Sons, Inc. Biol. 1:123-139  Keywords: heart-rate monitoring r mice r electrocardiography ECG

INTRODUCTION With increasing numbers of transgenic mouse strains, the treatment and evaluation of cardiovascular pathophysiology have been explored in more ways than previously possible. The monitoring and controlling of heart rate is central to meaningful interpretation of the results. In the conscious state the normal murine heart rate ranges from 500 to 700 beats per minute (bpm) depending on the time of the day, environmental factors, and activity levels (Kramer et al., 1993; Uechi et al., 1998a; D’Angelo et al., 1997). The monitoring of heart rate is of particular interest since it often reflects the dynamic changes of cardiac function in response to environmental and hemodynamic changes (Appel et al., 1989). We have previously discussed imaging methods such as echocardiography (Gao et al., 2011) and arterial pressure monitoring methods including tail cuff system, implanted fluid filled catheter, Millar solid state micro-pressure transducer tipped catheter, and implanted telemetry systems (Zhao et al., 2011). All of these methods allow for the indirect measurement of heart rates once the number of cycles of either arterial pressure waves or cardiac contractions in a given time is calculated (Fig. 1). However, since these methods only detect changes in cycle length of successful cardiac contractions and cannot differentiate ectopic beats from sinus beats, the electrocardiogram (ECG) is considered to be the “gold standard” for the monitoring of heart rate. Additionally, as with arterial pressure and cardiac function, one must consider the effects of anesthesia on heart rate (Table 1). The major techniques that focus on the monitoring of conscious heart rate and that are able to determine the source of the impulse from the heart all revolve around the use of ECG. In this article, we will describe the three primary methods that utilize ECG—e.g., noninvasive ECG system, tethered ECG, and implanted telemetric ECG monitoring—as well as secondary methods involving the implementation of pulse oximetry and the previously discussed techniques of echocardiography and

Current Protocols in Mouse Biology 1: 123-139, March 2011 Published online March 2011 in Wiley Online Library (wileyonlinelibrary.com). DOI: 10.1002/9780470942390.mo100159 C 2011 John Wiley & Sons, Inc. Copyright 

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A

B

AOP

200

200

150

150 mmHg

mmHg

AOP

100

50

50 PE

0

C

D

HR 1000

800

800

600

600

bpm

1000

400 PE

200

SNP

0

HR

bpm

100

400 200

SNP

0

0

Figure 1 Calculation of heart rate (HR) (panels C and D) by counting the number of aortic pressure waves (AOP) (panels A and B) in a sedated mouse. The arrows represent the injections of phenylephrine (PE) to increase pressure and sodium nitroprusside (SNP) to decrease pressure. The arterial baroreceptor reflex reduces heart rate with pressure elevation and decreases heart rate when pressure falls. Table 1 Heart Rate (HR) in Conscious and Anesthetized Mice

Anesthesia

None

Avertin

Ketamine/xylazine

Pentobarbital Isoflurane

HR (bpm)

580±18

411±17

159±11.8

377±11

457±17

Data resource

Uechi et al., (1998a)

Yan et al. (2007)

Erhardt et al. (1984)

Yang et al. (1999)

Stypmann (2007)

measurement of arterial pressure, which also provide an indirect assessment of heart rate. All the methods of ECG monitoring allow for the assessment of all electrical activities in the heart in addition to providing information on heart rate, whereas measuring the arterial pressure only assesses those impulses that result in contraction. Through the use of ECG, the origin of the electrical activities in the heart can be determined, thus allowing for the monitoring of cardiac rhythm and the determination of heart-rate variability, as well as cardiac conduction abnormalities. Materials and methods needed for the monitoring, as well as the advantages and limitations for each of these methods, are discussed.

Heart Rate and ECG Monitoring in Mice

Heart-rate variability This observed time variation from beat to beat is a normal physiological phenomenon, and is the result of changes in sinoatrial (SA) node stimulation. This beat-to-beat variation in heart rate not only reflects a change in physiologic condition, volume status, and cardiac function, but also in the animal’s ability to respond to both environmental and physiologic stress, such as changes in volume status, arterial pressure, and autonomic tone. This beat-to-beat variation is best described as a change in RR interval (time interval between consecutive R waves in an ECG tracing; see Fig. 2) seen on an ECG. Some of the main regulators of this beat-to-beat variation include both the sympathetic and parasympathetic efferent arms and baroreflex afferent arm of the autonomic reflex system (Uechi et al., 1998a; Kovoor et al., 2001). Body temperature, stress, sleep-wake cycle, and animal activity all affect the autonomic tone of the animal, and thus affect beat-to-beat variability. Furthermore, a decrease in heart-rate variability indicates a decrease in the

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RR mouse 1 P

S

R

P

T

mouse 2

T

S

RR

mouse 3

T

P

PR S

Figure 2 Sample of telemetry ECG simultaneous recording of three mice. A typical mouse ECG contains a P wave, followed by the QRS complex and the T wave. The distance between two consecutive R waves is termed RR interval and represents the time it takes for one beat, and the PR interval represents the time for electrical conduction from the atria to the ventricle. The plus (+) on the ECG represents the computer-identified R wave.

ability of an individual to adapt to stress, and has been associated with increased mortality for both myocardial infarction (Kleiger et al., 1987; Bigger et al., 1992) and congestive heart failure patients (Sandercock and Brodie, 2006). As an example, an increase in heart rate with a decrease in heart-rate variability is seen in transgenic mice overexpressing cardiac Gsα, and is felt to be a risk factor for the development of cardiomyopathy in these transgenic mice (Uechi et al., 1998a).

Comparing conscious versus sedated heart-rate monitoring Various inhalant or injectable agents are available for anesthesia or sedation in mice, either as pre- and perioperative anesthetic agents or as sedatives during a specific experiment. As described in the articles on echocardiography and arterial pressure monitoring (Gao et al., 2011; Zhao et al., 2011), the effects of anesthesia on cardiac function, blood vessels, and reflex control of the circulation have been well established (Vatner and Braunwald, 1975; Vatner et al., 2002). Therefore, studies in conscious animals are preferred when possible (Kurtz et al., 2005a,b), and if anesthesia must be used, the choice of anesthetic agents is of particular importance since different agents may lead to different degrees of autonomic and heart-rate suppression in mice (Table 1; Erhardt et al., 1984; Yang et al., 1999; Odashima et al., 2007; Stypmann, 2007; Yan et al., 2007). The combination of ketamine/xylazine (50 to 100 mg/kg/2.5 mg/kg) via intraperitoneal (i.p.) or intramuscular (i.m.) injection provides ∼30 min of satisfactory anesthesia. By increasing xylazine from 2.5 mg/kg to 10 mg/kg, the duration of the anesthesia is prolonged to 40 min (Xu et al., 2007). One should exercise caution with the use of xylazine, since it is a potent respiratory depressant. Therefore, if re-dosing is necessary, ketamine alone (one-half the original

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dose) should be used. However, due to the more pronounced suppression of heart rate, the ketamine/xylazine mixture is less desirable for acute experiments. Tribromoethanol (Avertin, 2.5%, 290 mg/kg i.p.) is the preferred agent for acute experiments due to its ability to maintain a more physiologic blood pressure (BP) and heart rate when compared to the ketamine/xylazine cocktail. However, Avertin has a relatively shorter half life and requires frequent re-dosing. Alternatively, pentobarbital (40 to 85 mg/kg i.p.) provides a more stable anesthetic duration of 20 to 60 min, and heart rates are higher under this anesthesia.

Three primary ECG methods for conscious heart-rate monitoring in mice ECG is the surface recording of the electrical currents passing through the heart and represents the electrical activities associated with each heart beat. ECG allows for the visualization of the electrical conditions of the heart. A typical ECG contains P, QRS, and T waves representing atrial contraction (P), ventricular contraction (QRS), and ventricular relaxation (T), with the time between each of these waves reflecting the time between each of these components (Fig. 2). Since, in general, R waves are not only the most distinct waveform with the highest voltage differential from baseline, but also the easiest for an automated system to detect with high fidelity; typically, the time between two R waves is considered the time for one beat. The digital heart-rate readout is generated by passing the recorded ECG signals through an amplifier, and the amplified signal is then analyzed by a cardiotachometer, where a timing device is started by the first pulse wave and stopped by the next, thus allowing for the calculation of heart rate. Alternatively, the signal is processed by data-analysis software. In either case the heart rates are displayed as digital readouts in beats per minute. It is important to note that although the measured electrical activities usually correspond to cardiomyocyte contractions, less electrical activity is seen in cases of extreme shock with poor perfusion pressure. BASIC PROTOCOL 1

Heart Rate and ECG Monitoring in Mice

NONINVASIVE ECG SYSTEM Due to the size, rapid heart rates, and difficulty associated with keeping the mice calm during experiments, the use of surface ECG for conscious monitoring of small animals remains an arduous task. Thus, to facilitate the recording of noninvasive ECG in conscious mice, a restraint device is often needed. In general, an ECG records the voltage between a pair of electrodes; the size and direction of deflection obtained reflects the size and direction of the electrical activity in the heart relative to the pair of electrodes. By analyzing the ECG tracings, information about the heart rates and cardiac rhythm is obtained. In mice, this is accomplished by using a narrow platform where three pawsized electrodes are embedded and then connected to an amplifier. The mouse is placed gently on the electrodes, where three of the paws are in contact with the electrodes for continuous recording. The electrical signal among these three electrodes is recorded, and the resulting ECG is then analyzed on a computer. Advantages and disadvantages of noninvasive ECG system This system has the advantages of noninvasively obtaining information not only on heart rates but also heart-rate variability, arrhythmia analysis, and RR, QT, and PR intervals, as well as conduction abnormalities in conscious animals. Thus, it provides information about the phenotype and/or the effects of pharmacologic intervention on both heart rate and arrhythmia risk. Furthermore, the system can be employed to screen a large number of mice for ECG changes related to genetic, pharmacologic, or pathologic alterations. The main disadvantage is that such a system is not suited for continuous long-term monitoring; it is better adopted for studies where periodic short-term ECG monitoring or screening is needed.

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Materials Mice AnonyMouse ECG screening system (Mouse Specifics, Inc., http://www.mousespecifics.com/) Cotton-tipped sticks Gel-coated ECG electrodes Amplifier (HP78901A, Hewlett-Packard) Shielded 3 electrode lead set (M1605A Snap, Hewlett-Packard) Recording computer eMouse Internet-based wave-analysis portal (http://www.mousespecifics.com/) 1. Gently place the mice onto the ECG platform and allow them to acclimate for at least 10 min. The platform is capable of testing one mouse at a time, while having two additional spots for the acclimation of two mice.

2. Using a cotton-tipped stick, gently push the mouse into position where the two front paws and the left rear paw are in contact with the electrodes. 3. Turn on the system and perform the recording as per the manufacturer’s manual. 4. While the recording is being completed on the first mouse, place the next mouse on the side platform to allow for acclimation. 5. Analyze the data once the recording is completed using the eMouse Internet-based wave analysis portal. 6. Calculate the heart rate by either counting number of R waves over time or averaging several RR intervals. 7. Calculate heart-rate variability by using the average beat to beat variability, i.e., RR interval variability within the recorded period. 8. Correct the QT by using the following formula (Mitchell et al., 1998):

QTc =

QT RR 100

Equation 1

Since all phases of the ECG are affected by HR, QTc is the heart-rate-corrected QT, allowing for comparison of QTc from beat to beat and animal to animal.

TETHERED ECG SYSTEM Similar to noninvasive ECG monitoring of mouse heart rate, the tethered system provides information via ECG tracing. Heart rates, conduction abnormalities, and arrhythmias can all be recorded and analyzed. Unlike the noninvasive method of ECG recording, four electrodes are embedded on the back of the mouse, where the wires are then tunneled under the skin and exit via a mid-scapular incision. The wires are attached to a swivel device via a spring tether and connected to an amplifier where the signal is continuously recorded using an analog or digital recorder. Advantages and disadvantages of tethered ECG system The advantage of such a tethered system is that it allows the mouse to have increased freedom of movement and access to food and water without the stress associated with

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the use of a small restraining cage. Therefore, this system allows for longer duration of recording. Furthermore, since the electrodes are attached to the animal and embedded under the skin, the signal is affected to a lesser extent by mouse movement when compared with the noninvasive method. However, the disadvantages of this system are the need for surgically implanting the electrodes under general anesthesia, as well as the associated surgical morbidity and mortality. Even though the tether is out of the reach of the mouse, the animal should not be left alone during recording. Therefore, such a system is most suited for periodic recording of intermediate duration.

Materials Mice Anesthesia (see Table 1 and annotation to step 2, below) Insulated wire leads (AWG size 36) Surgical instruments Suture: 6-0 nylon for ECG lead fixation and skin closure Tether system including a harness (or jacket) (Instech, http://www.instechlabs.com/) Swivel system (Instech) ECG system including: Amplifier box (Gould Instrument Systems, item no. 11-4123-09) Amplifier (Gould Instrument Systems, item no. Amp6600) Tektronix TDS 1002 Oscilloscope (Tektronix, part no. 93K5765) Small protective plastic tube (made by cutting a 4-mm section from the mid-portion of a plastic transfer pipet; VWR, cat. no. 3-711-7) Restraint box with ceiling hole (Braintree Scientific, cat. no. 500M-C) 1. Prepare the implantable ECG wires: a. Four strands of insulated wire are tied together as a bundle about 5 cm from the end (make sure the length behind the tie is longer than the length of spring tether) and make the two wires for the forelimbs shorter (2.5 cm) (Fig. 3).

LA

ECG

RA RA

RL LA

LL

EKG limb lead wires

LL

RL

Figure 3 An illustration of tethered ECG insertion and recording of mouse heart rate. The ECG leads are labeled and subcutaneously tunneled onto the back of the mouse in positions of the four limbs. Mice are housed individually during the recording process with the leads connected to the tether system for data acquisition.

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b. Each tip of the wire is then made bare by removing 3 mm of insulation from the wire to increase the contact surface. Make sure to mark each wire to indicate which tip is for left arm (LA), right arm (RA), left leg (LL), and right leg (RL) leads, and do not forget to distinguish and mark which one of the longer-length wires corresponds to the forelimb leads so that proper recording of the signal is achieved once the whole system is connected to the recording machine.

2. Anesthetize the animal and place the mouse in the prone position. The choice of anesthetic depends upon the experimental design and the experience of the individual laboratory. Please refer to Table 1 for the effects of each of the anesthetic agents on the heart rate. Different agents may be used for surgery and experiments. In our laboratory, we generally use a mixture of ketamine and xylazine for surgery and perform the experiments under conscious conditions.

3. Make a 1-cm transverse incision in the mid-scapular region of the mouse. 4. Transfix the tied portion of the ECG wires to the underlying muscle with 6-0 nylon suture. 5. Tunnel the four shorter-length wires subcutaneously in the direction of the four limbs, respectively, so that the bare tips are in good contact with the underlying muscle (Fig. 3). 6. Bring out the four longer-length portions of the wires through the previously made skin incision and close with nylon sutures.

For tether system 7a. Pass the wires through the harness and connect to the swivel assembly via a spring tether. 8a. Connect the wires to an ECG signal amplifier and a recording system.

Alternative method: Handling the exteriorized ECG wires Since the tether system is generally employed to house the fluid line and wires together, and is more suited to house the fluid line for protection, if subcutaneous implantation of ECG wires is the only procedure needed for monitoring, the tether system can be avoided by following the simpler steps below. 7b. Coil the exteriorized wires to be housed in a small protective plastic tube which is suture-fixed to the skin around the closed mid-scapular incision site. 8b. Whenever ECG recording is needed, place the mouse in a restraint box which has a ceiling hole. 9b. Deliver the housing tube through the ceiling hole and uncoil the wires after bringing them out of the housing tube, then properly connect the four designated limb leads to the recording system. 10b. Once the recording is complete, re-coil the wires and return them to the housing tube for protection until the next recording.

IMPLANTED ECG TELEMETRY SYSTEM The ECG radiotelemetry units are the gold standard for monitoring the heart rates in conscious mice. A typical mouse ECG telemetry system contains two electrical ECG leads connected to a radio transmitter. The radio transmitter is placed into a subcutaneous pocket on the back of the mouse, and the leads are implanted with one lead toward the right upper chest and the other near the left lower chest. Upon activation of the transmitter by a magnet, the electrical signals are transmitted wirelessly to a nearby receiver attached to an amplifier and computer system for data acquisition, storage, and analysis. Through Current Protocols in Mouse Biology

BASIC PROTOCOL 3

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Heart rate (beats/min)

900

800

600 wild-type TG 400 0

10

20

30

40

50

60

Heart rate (beats/min)

Time (min)

600

wild-type TG

300

day 0 06:00

night 18:00

day 06:00

night 18:00

06:00

Time

Figure 4 Circadian variation of heart rate in transgenic mice with cardiac specific overexpression of Gsα (TG) and wild-type mice. Heart rates were obtained from ECGs recorded using a telemetric system in conscious unrestrained mice. The inset shows the difference in heart rate variability over a 1-hr period in one TG and one wild-type control mouse. Figure and legend used and modified with permission (Uechi et al., 1998a).

such a system, the heart-rate variability in different transgenic animals is recorded and analyzed, as exemplified by the decreased heart-rate variability seen in transgenic mice overexpressing Gsα (Fig. 4; Uechi et al., 1998a). Advantages and disadvantages of implanted telemetry system With the use of an implanted telemetry device, a continuous conscious recording of the mouse heart rate is possible. This provides a method for long-term monitoring of the mice in their natural living environment, which is often needed to detect circadian variations, as well as to monitor arrhythmia frequencies and to determine if arrhythmias are the cause of death in transgenic mice. Recently, the telemetry systems that allow simultaneous monitoring of ECG and arterial pressures in mice are in development. The disadvantages include high cost as well as surgically related morbidity and mortality. Since only two electrodes are present, only a single lead of ECG recording is obtainable. Furthermore, a 5- to 7-day recovery period is often needed after implantation, for stabilization of the heart rates (Lorenz, 2002). Therefore, the implanted telemetry system is most suited for studies where long-term monitoring of mouse heart rate and cardiac rhythm in a natural living environment is needed.

Materials Mice Anesthesia (see Table 1 and annotation to step 2, below)

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Surgical instruments Suture: 5-0 nylon for transmitter fixation to skin; 7-0 silk for telemetry wire fixation Telemetry transmitter device (ETA F-10 or EA-F20; Data Sciences International) Magnet (any commercially available magnet can be used) Anesthesia (see Table 1 and step 1) Current Protocols in Mouse Biology

ECG wire

telemetry unit

preformed holes for anchoring

anchoring suture (blue)

Figure 5 An illustration of the telemetry unit with the anchoring sutures in place. In general two to three anchoring sutures are used to fix the telemetry unit to the chest wall of the mice.

1. Anesthetize and place the mouse in the supine position, then make a 1-cm midscapular transverse incision on the back of the mouse. The choice of anesthetic depends upon the experimental design and the experience of the individual laboratory. Please refer to Table 1 for the effects of each of the anesthetic agents on heart rate. Different agents may be used for surgery and experiment. In our laboratory we generally use a mixture of ketamine and xylazine for surgery and perform the experiments under conscious condition.

2. Create a subcutaneous pocket by using a pair of round-tip scissors undermining the skin on the left lower back to create a pocket large enough to accommodate the transmitter device. 3. Insert the transmitter device into the pocket with the side of the device containing the predrilled suture holes facing up and transfix it to the overlying skin with 5-0 nylon sutures (Fig. 5). 4. Remove 5 mm of silicon insulation from the tips of both wires to increase the contact surface. 5. Make an extra small skin cut in the right shoulder area and tunnel the negative wire (white) subcutaneously toward the skin cut, and, through the cut opening, transfix the bare wire tip to the underlying muscle with a 7-0 silk suture. 6. Turn over the animal, tunnel the positive wire (red) subcutaneously, and transfix the tip to the abdominal muscle through a small skin cut just caudal to the left rib cage in a similar fashion. The correct lead length is managed by coiling the wires around the device.

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telemetered ECG

receiver

transgenic

wild-type

0.1 sec

Figure 6 Sample of ECG recording using a telemetric system for a free-ranging transgenic mouse overexpressing Gsα (top) and a wild-type control mouse (bottom). The telemetric implant is shown in the abdominal cavity of the mouse. The recorded heart rates were higher in the transgenic mice. The darker lines at the right of the ECG tracing shown are the compressed ECG tracing printed at a slow paper speed. Figure and legend used and modified with permission (Uechi et al., 1998a).

7. Close all incisions and cut openings with nylon sutures, with the knots buried beneath the skin. 8. Power on the battery in the transmitter using a magnet. Verify the functioning of the transmitter by powering up an AM radio, tuning to 88 MHz, and bringing it close to the device, then listening for a high-pitched tone corresponding to the cardiac cycle.

9. Return the animal to the cage and allow to recover. 10. Place the signal-reception plate under the animal cage and connect the cables to the data acquisition system for recording, storage, and analysis (Fig. 6). BASIC PROTOCOL 4

Heart Rate and ECG Monitoring in Mice

PULSE OXIMETRY The use of pulse oximetry has been a long-accepted method for the monitoring of mammalian heart rates and blood oxygen saturation. Iyriboz et al. (1991) validated the accuracy of pulse oximetry for the monitoring of heart rates both at rest and during exercise. Since that time, many different versions of the pulse oximeter have been developed for use in small laboratory animals, including mice. In general, pulse oximetry uses the inherent differences in the level of absorption of red light (600 to 750 nm) and infrared light (850 to 1000 nm) between deoxy- and oxyhemoglobin. By transmitting these two different wavelengths of light through a translucent part of the animal and monitoring the received light by a photodiode, the ratio of oxy- to deoxyhemoglobin is determined. Furthermore, since the signal intensity increases with arterial vessel expansion associated with the heart beat, the pulse rate is obtained simultaneously and recorded on a computer for data analysis. Advantages and disadvantages of pulse oximetry The advantage of such a system is that it is noninvasive and allows for the conscious monitoring of mouse heart rate while simultaneously monitoring the blood oxygen saturation. By using the neck clip, the mouse has limited amount of freedom of movement, yet the

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heart rate is monitored in a relatively non-stressed state. However, the disadvantage of such a system is that it does not allow for long-term monitoring and free movement, with the latter usually leading to distortion of the transmitting signal. Furthermore, the measured hemoglobin oxygen saturation may not always reflect accurately the respiratory sufficiency, as the measured oxygen saturation reflects only the percent hemoglobin saturation and not necessarily the amount of delivered oxygen. In cases of severe anemia, the actual amount of oxygen delivered is low despite a high percent saturation. Additionally, carboxyhemoglobin and methemoglobin all lead to falsely elevated oxygen saturation.

Materials Mice MouseOx pulse oximeter with CollarClips, ThroatClips, and Thigh Sensors (Starr Life Sciences Corp., http://www.starrlifesciences.com/) Hair clippers MouseOx software (Starr Life Sciences Corp., http://www.starrlifesciences.com) STARR-Link analog output module (Starr Life Sciences Corp.) 1. Prepare the animals by training them with desired clip for at least 1 hr prior to the scheduled experiment (3 to 6 hr are preferred). 2. Select the correct sensor. Extra-small-sized sensors are used on an unshaven mouse with body weight lower than 15 g, or a shaved mouse with body weight lower than 20 g. For mice of greater body weight, use a small-sized sensor.

3. Remove hair from the clip sensor site to allow for improved signal acquisition (preferably remove hair several hours prior to the start of the experiment). With the pulse oximeter, the experiments are conducted in either the conscious or sedated condition. Usually a white CollarClip is used in both conditions with animals in a prone position, while a black ThroatClip and Thigh Sensor are used only on anesthetized mice in the supine position.

4. Place the CollarClip right behind the ears and center the clip handle. For neonates, the clip is placed on the head across the ears. If the mouse struggles, hold the tail and let its front legs hold on to the cage, then attach the sensor. If necessary, animals may be lightly anesthetized to help with placement of the sensor. It is important to make sure that the clip is in the correct position with the tines squeezed together, to ensure consistent signal quality.

5. Send the mouse back to the cage with just enough length of wire so that the animal will not chew on the loose cable. 6. Place the animal on a nonreflective surface and use a light-blocking cloth covering on the cage to avoid light reflection or artificial light. 7. Connect the other end of the wires to the MouseOx pulse oximeter before starting the software. Then follow the menu to start recording. 8. Go to the MouseOx software main menu, and select “Monitor Subject,” which will ask whether the animal is anesthetized or awake. 9. Push the “Start/Reset” button to initiate data recording after the animal is stable for at least 15 min. If the signal does not come up quickly, check if the sensor is in the “ON” position. If the sensor is removed from the animal during recording, the “Start/Reset” button has to be clicked again when the sensor is put back on the animal.

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10. Perform the recordings. The recording window displays heart-rate waveforms on the left panel and Pulse Pleth on the right upper panel. Below the Pulse Pleth figure, there is a digital display showing the real-time reading and cursor-pointed reading. If pulse distention shows low values (<5 μm), it may be due to vessel contraction or low blood volume, etc. If the experiment requires controlling a relatively stable heart rate, a “Set alarm” button (on the right panel) is used to remind the investigator if the heart rate is above or below the limit set for the experiment.

11. Save the MouseOx data results as a .txt file accessible by Microsoft Excel for further analysis. In the last column, the listed “error codes” provide information on unstable pulse or lost pulse/signal. The manufacturer’s manual shows the listed description of all the codes.

COMMENTARY Background Information The monitoring of mouse heart rate is central to mouse phenotyping and determination of mouse cardiovascular pathophysiology. Different genetically altered mouse models have been shown to have either markedly higher or lower baseline heart rates (Table 2; D’Angelo et al., 1997; Uechi et al., 1998b; Peter et al., 2007), making the determination of baseline heart rate an important variable for phenotyping. An added dimension to differences in heart rates is manifested by the differences in heart-rate variability. Heart-rate variation reflects not only cardiac function but also the animal’s ability to respond to physiologic and environmental stress such as change in volume status, arterial pressure, and autonomic tone. For the monitoring of cardiovascular function and physiology, various methods have been developed and described. Below we discuss three major ECG methods, as well as other secondary methods such as pulse oximetry, echocardiography, and arterial pressure wave-form analysis for determination of the heart rate in mice.

It has been over a century since William Einthoven first described the use of surface electrodes to record cardiac electrical activity and generating the first ECG (Rivera-Ruiz et al., 2008). Since that time, ECG has been used to monitor the cardiac electrical activity within different mammalian models. Since R waves are the most distinct wave forms, they are the easiest for an automated system to detect with high fidelity. The amplified ECG signals are analyzed by either cardiotachometer or commercially available software where heart rate is determined by calculating the time between two consecutive R waves. In general, all three primary methods involve the analysis of ECG function by way of using the surface electrodes to monitor the electrical activity in the heart. By observing this electrical activity, the heart rate is calculated with the assumption that each of the observed electrical pulses results in a cardiac contraction cycle. However, in cases of extreme shock and hypo-perfusion pulse, less electrical activity is observed. The major advantage of ECG is its ability to assess all

Table 2 Baseline Heart Rates of Transgenic Micea

Overexpressed gene Transgenic Wild-type Data source β1 -AR

507±18

423±20

Peter et al. (2007)

β2 -AR

551±51

423±20

Peter et al. (2007)

Gsα

736±15

580±18

Uechi et al. (1998a)

Gαq-25

427±9

690±13

D’Angelo et al. (1997)

a Baseline heart rate in beats per minute in transgenic mice over-expressing β1-β2-

Heart Rate and ECG Monitoring in Mice

adrenergic receptors (AR) (conducted under Avertin anesthesia) or Gsα or Gαq (conducted in conscious mice) are compared to wild-type littermates.

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electrical activities within the heart, whereas measuring arterial pressure only assesses those impulses that result in contraction. ECG monitoring allows for the identification of arrhythmia and arrhythmogenic electrical activities. By determining the origin of cardiac electrical activity, those heart beats that originate in the SA node can be identified, and this information is needed for the determination of heart-rate variability. Noninvasive ECG system This method (Basic Protocol 1) uses a narrow platform embedded with three paw-sized electrodes where contact is maintained between three paws of the mouse and the electrodes. The electrical signal among these three electrodes is recorded and the resulting ECG is used for the analysis of heart rates. Tethered ECG system This system (Basic Protocol 2) embeds four electrodes on the back of the mice that are tunneled under the skin toward four limbs and exited via a mid-scapular incision. These wires are connected to an amplifier, and the obtained ECG signals are then used for heart-rate analysis. Radiotelemetry ECG system This system (Basic Protocol 3) is the gold standard for conscious monitoring of heart rates. A typical mouse radiotelemetry system for ECG monitoring contains two electrical ECG leads connected to a radio transmitter. Once they are implanted, the ECG signal is transmitted wirelessly to receiving and recording units for heart-rate analysis via commercially available software. Secondary methods for the monitoring of heart rate in mice Echocardiography, tail-cuff arterial pressure monitoring system, implanted fluid-filled catheter, Millar solid-state micro-pressuretransducer-tipped-catheter, and the implanted telemetry system can all be used for the determination of heart rate through either computer analysis software or a cardiotachometer, whereby the heart rate is determined by measuring the time required for each of the cycles of either arterial pressure waves or cardiac contractions; in this way, a heart-rate tracing is generated (Fig. 1). Echocardiography In terms of echocardiography, the heart rate is determined either by analyzing the M-mode images or using the simultaneous ECG recordings. Since distance per time is a known vari-

able, by marking either the systolic or diastolic peak of the M-mode images, or the R wave peak on the ECG, distance between the peaks is calculated by the image-analysis software. These data points are then imported into data-analysis software such as Microsoft Excel, whereby the time between the beats is calculated. Then, by averaging 3 to 4 consecutive beats, average heart rates are determined. Determination of heart rate along with arterial-pressure monitoring In the case of implanted fluid-filled catheters, Millar solid-state-micro-pressuretransducer-tipped catheter, and the implanted telemetry system, the heart rate is calculated via analysis of the pressure-wave signals by the recording system and displayed as an instantaneous readouts on the computer screen. In brief, the recorded pressure-wave signals are first passed through an amplifier, and the amplified signals are then processed through a cardiotachometer where a timer is started by the first pulse wave and stopped by the next wave, after which the heart rate is calculated and displayed in beats per minute. An additional method for heart-rate monitoring is the tail-cuff system, in which the heart rate is calculated via the attached pulse oximetry detailed below. Since these methods are only able to detect changes in cycle length and are unable to exclude ectopic heart beats that do not originate from the SA node, the ECG is considered to be superior for the monitoring of heart-rate variability.

Critical Parameters and Troubleshooting Noninvasive ECG system 1. It is a necessary step to acclimate the animals by keeping them on the platform for at least 10 min before experiments. 2. It is important to use the correctly sized electrode pads for the size of the mouse. Place the electrode pads with a spacing of 2.5 to 3 cm. Spacing that is too large or too small may cause a loss of signal with animal movement. 3. It is best to avoid bright lights in the room and reduce nearby motion and noise during the experiments, as any of these factors may lead to agitation of the mouse, thus leading to poor data quality. Tethered ECG system 1. For the implanted ECG tether, embedding the leads within the muscles (thus keeping the leads in good contact position) results

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in a more stable signal. A suitable way is to loop the bare wire tips and bury them between the muscle fibers. Fixation sutures are used to securely hold the wire tips in place. 2. Prior to recording it is important to establish connection of the properly designated lead wires to the recording system. Therefore, confusion will be minimized by marking each of the wires with any available means (e.g., knotting, painting, etc.) to indicate which tip is for left arm (LA), right arm (RA), left leg (LL), and right leg (RL) leads during implantation. Implanted ECG telemetry system The critical parameters are the same as those of the tether system. Proper fixation of the ECG leads will help to obtain a stable signal as well as lessen the pain associated with ECG wire tip movement within the muscles. Pulse oximetry The most common problem encountered with the system is poor signal acquisition. The following will help minimize this potential complication: 1. It is important to use the right size sensor and make sure the clip is well attached. 2. Shaving the sensor site will help improve signal quality, especially on mice with dark fur, in which case shaving is required. 3. For the thigh clip, it should be applied to the upper thigh, as close to the pelvis as possible, to ensure good signal acquisition. 4. It is important to ensure that the animals maintain a physiological temperature during recording. Utilize an external heating device (i.e., heating pad or lamp) under or over the cage if needed. 5. If all else fails and signal quality continues to remain poor, the sensor application site should be readjusted; press the “Start/Reset” button to restart the recording.

Anticipated Results The anticipated result for all three primary methods of obtaining and recording mouse ECG is the recording of the ECG tracings (Figs. 2 and 6) to be used for data analysis

by ECG analysis software. Whereas the anticipated result for echocardiography is the recording of echocardiographic images, often accompanied by ECG tracings (for examples please refer to our previous paper on echocardiography in Gao et al., 2011), and the anticipated result for methods involving invasive monitoring of arterial pressure is the recorded arterial pressure-wave form (Fig. 1), the anticipated result from the heart-rate monitoring systems is an analog or digital readout of the heart rate. The effects of physiologic and experimental conditions, sedation, volume status, time of day, age, gender, activity, and temperature on the measured heart rate has been well established. Therefore, there is often some disagreement regarding the measured heart rate obtained from different laboratories. However, as a general reference, some normal conscious heart rate values for a few of the more commonly used strains of mice can be found in Table 3. In an ideal experiment, arterial pressure should also be monitored. With abnormal blood pressure or unexpected deviation in either heart rate or respiratory rate, one should consider repeating the experiment. Considerations and experimental design The common aspect shared by the two noninvasive methods (pulse oximetry and noninvasive ECG monitoring) is that both of them share the advantage of being noninvasive and both are more suited for short-term monitoring. Although pulse oximetry provides additional information in terms of oxygen saturation, the proper interpretation of this information must take into account hemoglobin concentration and the possible presence of carboxyhemoglobin and methemoglobin. An important advantage of ECG systems over indirect measurements of heart rate is the ECG’s ability to discriminate heart beats originating from the SA node versus ectopic heart beats. This is of particular importance, since it allows for the exclusion of ectopic beats, thus allowing for the calculation and comparison of heart-rate variability. Noninvasive ECG monitoring provides information on rhythm in

Table 3 Conscious Heart Rate (HR) in Three Strains of Micea

Heart Rate and ECG Monitoring in Mice

Strain

C57/B6SJL

FVB

129Sv/J

HR (bpm)

588±14

617±26

522±11

MAP (mmHg) 103±1

99±2

102±8

Data resource Uechi et al. (1998a)

Lin et al. (2010)

Gross and Luft (2003)

a All values are mean±SE. MAP=mean arterial pressure; HR=heart rate.

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addition to rate. This allows for the measurements of QT, RR, and PR intervals. At a basic level, this allows for the detection of conduction abnormalities and the presence of arrhythmias. Furthermore, since RR interval represents the time between two consecutive beats, the variation of RR intervals represents heartrate variability; a decrease in this variability has been shown to be associated with increased mortality after myocardial infarction (Kleiger et al., 1987; Bigger et al., 1992) and congestive heart failure (Sandercock and Brodie, 2006). There are currently two major methods used for the comparison of heart-rate variability: a time-domain method, where the heartrate variability is compared over a given time period (usually 24 hr), versus a frequencydomain method where the frequencies of the heart rates are compared, most commonly via discrete Fourier transformation. Aside from heart-rate variability, the heart rate–corrected prolongation in QT has also been associated with an increase in cardiac arrhythmia and in cardiac mortality in the elderly (de Bruyne et al., 1999) as well as the general population (Montanez et al., 2004). Animal models that are capable of detecting drug-associated QT prolongation may prove to be useful for the safety screening of pharmacologic compounds. In mouse models, one accepted formula for the heart rate–associated QT correction is represented by Equation 1, above (Mitchell et al., 1998). In noninvasive ECG systems, the mouse must maintain continuous contact with the platform for recording; therefore, animal movement has the potential to disrupt this contact, and long-term continuous ECG monitoring is not possible. If longer-term continuous monitoring is desired, a tether allows for the monitoring of free-moving mice for an extended period of time under the direct supervision of the investigator. In addition, the monitoring of heartrate changes associated with exercise is possible with this system. The data are analyzed and graphically illustrated to reflect heart-rate changes and heart-rate variability over the entire study period. However, if long-term monitoring of circadian variability is needed, the implanted system should be used. This is considered the gold standard for the monitoring of heart rate in mice, as it allows for obtaining the most physiological values. Furthermore, telemetry systems capable of simultaneous recording of arterial pressure and ECG are now in development.

In brief, any particular method selected for monitoring heart rate in mice should depend on the experimental purpose, design, and duration. The methods and materials discussed in this article provide a practical blueprint for utilization of the chosen technique.

Time Considerations Noninvasive ECG system Expect acclimation time of at least 10 min and ∼10 to 15 min to complete the recording procedure. Tethered ECG system Expect ∼5 to 10 min to prepare and anesthetize the animal. While waiting for the anesthesia to take effect, the ECG wires can be prepared. The implantation of the wires can be completed in ∼10 to 20 min. Once the procedure is complete, allow the animal to recover overnight prior to recording. Actual time requirements depend upon the skill of the surgeon. In general, it takes ∼2 min to connect the animal to the recording system. If the harness and swivel are to be used for recording, an additional 5 min of time should be added for connecting the animal to the recording system. Implanted ECG telemetry system Expect ∼5 to 10 min to prepare and anesthetize the mouse. The implantation of the telemetry unit takes ∼5 to 20 min to complete. Actual time requirements depend upon the skill of the surgeon. Once the procedure is complete, allow the animal to recover for 3 to 5 days prior to recording. Pulse oximetry The mouse should be trained for at least 1 hr prior to recording (3 to 6 hr preferred). Signals are obtained instantaneously. Allow the animal to stabilize for at least 15 min prior to recording. Total preparation time prior to training/recording is 5 min.

Acknowledgements Supported in part by NIH grants HL033107, HL069020, AG027211, HL101420, HL093481, HL059139, HL095888, DK083826, and HL102472.

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Bigger, J.T. Jr., Fleiss, J.L., Steinman, R.C., Rolnitzky, L.M., Kleiger, R.E., and Rottman, J.N. 1992. Frequency domain measures of heart period variability and mortality after myocardial infarction. Circulation 85:164-171.

Lin, M., Harden, S.W., Li, L., Wurster, R.D., and Chen, Z. 2010. Impairment of baroreflex control of heart rate in conscious transgenic mice of type 1 diabetes (OVE26). Auton. Neurosci. 152:6774.

D’Angelo, D.D., Sakata, Y., Lorenz, J.N., Boivin, G.P., Walsh, R.A., Liggett, S.B., and Dorn, G.W. 2nd . 1997. Transgenic Galphaq overexpression induces cardiac contractile failure in mice. Proc. Natl. Acad. Sci. U.S.A. 94:8121-8126.

Lorenz, J.N. 2002. A practical guide to evaluating cardiovascular, renal, and pulmonary function in mice. Am. J. Physiol. Regul. Integr. Comp. Physiol. 282:R1565-R1582.

de Bruyne, M.C., Hoes, A.W., Kors, J.A., Hofman, A., van Bemmel, J.H., and Grobbee, D.E. 1999. Prolonged QT interval predicts cardiac and allcause mortality in the elderly. The Rotterdam Study. Eur. Heart J. 20:278-284. Erhardt, W., Hebestedt, A., Aschenbrenner, G., Pichotka, B., and Blumel, G. 1984. A comparative study with various anesthetics in mice (pentobarbitone, ketamine-xylazine, carfentanyletomidate). Res. Exp. Med. (Berl.) 184:159169. Gao, S., Ho, D., Vatner, D.E., and Vatner, S.F. 2011. Echocardiography in mice. Curr. Protoc. Mouse Biol. 1:71-83. Gross, V. and Luft, F.C. 2003. Exercising restraint in measuring blood pressure in conscious mice. Hypertension 41:879-881. Iyriboz, Y., Powers, S., Morrow, J., Ayers, D., and Landry, G. 1991. Accuracy of pulse oximeters in estimating heart rate at rest and during exercise. Br. J. Sports Med. 25:162-164. Kleiger, R.E., Miller, J.P., Bigger, J.T. Jr., and Moss, A.J. 1987. Decreased heart rate variability and its association with increased mortality after acute myocardial infarction. Am. J. Cardiol. 59:256-262. Kovoor, P., Wickman, K., Maguire, C.T., Pu, W., Gehrmann, J., Berul, C.I., and Clapham, D.E. 2001. Evaluation of the role of I(KACh) in atrial fibrillation using a mouse knockout model. J. Am. Coll. Cardiol. 37:2136-2143. Kramer, K., van Acker, S.A., Voss, H.P., Grimbergen, J.A., van der Vijgh, W.J., and Bast, A. 1993. Use of telemetry to record electrocardiogram and heart rate in freely moving mice. J. Pharmacol. Toxicol. Methods 30:209-215. Kurtz, T.W., Griffin, K.A., Bidani, A.K., Davisson, R.L., and Hall, J.E. 2005a. Recommendations for blood pressure measurement in animals: Summary of an AHA scientific statement from the Council on High Blood Pressure Research, Professional and Public Education Subcommittee. Arterioscler. Thromb. Vasc. Biol. 25:478479.

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Kurtz, T.W., Griffin, K.A., Bidani, A.K., Davisson, R.L., and Hall, J.E. 2005b. Recommendations for blood pressure measurement in humans and experimental animals. Part 2: Blood pressure measurement in experimental animals: A statement for professionals from the Subcommittee of Professional and Public Education of the American Heart Association Council on High Blood Pressure Research. Arterioscler. Thromb. Vasc. Biol. 25:e22-e33.

Mitchell, G.F., Jeron, A., and Koren, G. 1998. Measurement of heart rate and Q-T interval in the conscious mouse. Am. J. Physiol. 274:H747H751. Montanez, A., Ruskin, J.N., Hebert, P.R., Lamas, G.A., and Hennekens, C.H. 2004. Prolonged QTc interval and risks of total and cardiovascular mortality and sudden death in the general population: A review and qualitative overview of the prospective cohort studies. Arch. Intern. Med. 164:943-948. Odashima, M., Usui, S., Takagi, H., Hong, C., Liu, J., Yokota, M., and Sadoshima, J. 2007. Inhibition of endogenous Mst1 prevents apoptosis and cardiac dysfunction without affecting cardiac hypertrophy after myocardial infarction. Circ. Res. 100:1344-1352. Peter, P.S., Brady, J.E., Yan, L., Chen, W., Engelhardt, S., Wang, Y., Sadoshima, J., Vatner, S.F., and Vatner, D.E. 2007. Inhibition of p38 alpha MAPK rescues cardiomyopathy induced by overexpressed beta 2-adrenergic receptor, but not beta 1-adrenergic receptor. J. Clin. Invest. 117:1335-1343. Rivera-Ruiz, M., Cajavilca, C., and Varon, J. 2008. Einthoven’s string galvanometer: The first electrocardiograph. Tex. Heart Inst. J. 35:174-178. Sandercock, G.R. and Brodie, D.A. 2006. The role of heart rate variability in prognosis for different modes of death in chronic heart failure. Pacing Clin. Electrophysiol. 29:892-904. Stypmann, J. 2007. Doppler ultrasound in mice. Echocardiography 24:97-112. Uechi, M., Asai, K., Osaka, M., Smith, A., Sato, N., Wagner, T.E., Ishikawa, Y., Hayakawa, H., Vatner, D.E., Shannon, R.P., Homcy, C.J., and Vatner, S.F. 1998a. Depressed heart rate variability and arterial baroreflex in conscious transgenic mice with overexpression of cardiac Gsalpha. Circ. Res. 82:416-423. Uechi, M., Uechi, H., Nakayama, T., Wakao, Y., Ogasawara, T., Takase, K., and Takahashi, M. 1998b. The circadian variation of urinary N-acetyl-beta-d-glucosaminidase and gammaglutamyl transpeptidase in clinically healthy cats. J. Vet. Med. Sci. 60:1033-1034. Vatner, S.F. and Braunwald, E. 1975. Cardiovascular control mechanisms in the conscious state. N. Engl. J. Med. 293:970-976. Vatner, S.F., Takagi, G., Asai, K., and Shannon, R.P. 2002. Cardiovascular physiology in mice: Conscious measurements and effects of anesthesia. In Cardiovascular Physiology in the Genetically Engineered Mouse (B.D. Hoit, ed.) pp. 257-275. Kluwer Academic Publishers, New York.

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Xu, Q., Ming, Z., Dart, A.M., and Du, X.J. 2007. Optimizing dosage of ketamine and xylazine in murine echocardiography. Clin. Exp. Pharmacol. Physiol. 34:499-507. Yan, L., Vatner, D.E., O’Connor, J.P., Ivessa, A., Ge, H., Chen, W., Hirotani, S., Ishikawa, Y., Sadoshima, J., and Vatner, S.F. 2007. Type 5 adenylyl cyclase disruption increases longevity and protects against stress. Cell 130:247-258. Yang, X.P., Liu, Y.H., Rhaleb, N.E., Kurihara, N., Kim, H.E., and Carretero, O.A. 1999. Echocardiographic assessment of cardiac function in conscious and anesthetized mice. Am. J. Physiol. 277:H1967-H1974. Zhao, X., Ho, D., Gao, S., Hong, C., Vatner, D.E., and Vatner, S.F. 2011. Arterial pressure monitoring in mice. Curr. Protoc. Mouse Biol. 1:105-122.

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Exercise Performance Tests in Mice Stefan Marcaletti,1 Charles Thomas,2 and J´erˆome N. Feige1 1

MusculoSkeletal Diseases, Novartis Institute for Biomedical Research, Basel, Switzerland Center of Phenogenomics (CPG), Ecole Polytechnique F´ed´erale de Lausanne, Lausanne, Switzerland 2

ABSTRACT Maximal exercise performance is a multifactorial process in which the cardiovascular component, the innervation of the musculature, and the contractile and metabolic properties of skeletal muscle all play key roles. Here, protocols are provided for assessment of maximal running capacity of mice on a treadmill, with a combination of short high-intensity paradigms primarily intended to test for maximal power and cardiovascular function, and longer low-intensity paradigms to assess endurance and oxidative metabolism in skeletal muscle. The coupling of treadmill running to indirect calorimetry, to correlate performance measurements to maximal oxygen consumption, is also C 2011 by John Wiley & Sons, Inc. described. Curr. Protoc. Mouse Biol. 1:141-154  Keywords: exercise r running r treadmill r VO2 max r endurance power

INTRODUCTION There are numerous ways rodents can exercise including walking, jumping, swimming and running, for which various experimental approaches for measurement have been described (Kregel et al., 2006). Here, protocols are provided for measuring maximal locomotor performance of mice using forced treadmill running. Basic Protocol 1 presents two experimental designs that assess short-term high-intensity performance (referred to herein as power) and long-term low-intensity performance (referred to herein as endurance). Basic Protocol 2 then describes how to couple treadmill running to real-time gas exchange measurements to evaluate maximal oxygen consumption.

MEASUREMENT OF FORCED EXERCISE PERFORMANCE ON A TREADMILL

BASIC PROTOCOL 1

Treadmill running is an established technique to force mice to run at their maximal level of performance. Provided that the animals are properly familiarized with the equipment and that an aversive stimulation is delivered at the rear of the belt to stimulate running, the time and distance run by a given individual until it cannot cope with the speed of the belt are direct readouts of exercise performance. Performance tests typically involve protocols where running speed increases progressively in order to let animals warm up and gradually adapt their metabolic and cardiovascular parameters. Since the tolerable duration of running decreases as a function of increasing speed and depends on different types of metabolism (i.e., anaerobic glucose and phosphocreatine utilization for shortterm performance at high speed versus aerobic glucose and fatty acid oxidation for longer exercise at lower intensity; Savaglio and Carbone, 2000; Billat et al., 2005), several protocols have been established with various speed/duration profiles to model these different components. Here, two protocols are presented to probe for maximal power performance (short high-intensity run) and endurance (long low-intensity run).

Materials Appropriate mouse strain (e.g., C57BL/6J) and housing facility Spray bottle of distilled water Mild cleaning agent for cleaning the belt, e.g., 0.75% lysoform Current Protocols in Mouse Biology 1: 141-154, March 2011 Published online March 2011 in Wiley Online Library (wileyonlinelibrary.com). DOI: 10.1002/9780470942390.mo100160 C 2011 John Wiley & Sons, Inc. Copyright 

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Treadmill: see, e.g., Fig. 1; the ideal set-up should include: Revolving belt with adjustable speed (0 to 100 cm/sec) Adjustable slope for up- and downhill running (+20◦ /−20◦ ) Independent lanes with covered tops adapted to the size of mice (∼100 mm width and 450 mm length) A platform at the rear of the belt where the animal can escape when exhausted or in case of a major issue; this platform should, however, be equipped with a system to generate aversive stimulation that forces animals to run during the test (e.g., electrical stimulation grid with adjustable intensity from 0 to 2 mA and automatic detection of each stimulation received) Real-time control of belt speed, slope, time spent running, distance traveled, and number of aversive stimulations received A number of lanes adapted to the number of animals to assess: commercial systems have been developed, e.g., by Panlab (http://www.panlab.com/), Columbus Instruments (http://www.colinst.com/), and TSE (http://www. tse-systems.com/) NOTE: It is recommended but not essential to use a treadmill that communicates with a computer and allows the experimenter to control and record the treadmill parameters (velocity, distance traveled, aversive stimulations per min/cumulative number of aversive stimulations). NOTE: To ensure optimal test results, a few sessions of familiarization with the setup are required a few days preceding the actual treadmill test. For C57BL6/J mice, 1 to 2 sessions is typically sufficient (Fig. 2), but this number should be adapted to the particular strain of mice under consideration.

Familiarize animals with apparatus 1. Tag all mice to be used in the experiment with appropriate device/technique (e.g., ear clips, tattoo, probe) to follow performance longitudinally through time. 2. Set up treadmill to allow animal adaptation (5 min per day at 15 cm/sec, +5◦ slope and 0.3 mA electrical stimulation).

Exercise Performance Tests in Mice

Figure 1 Example of a commercial 5-lane treadmill setup for mice (Panlab). The treadmill is a dual system for mice and rats, but is shown in a mouse configuration including dedicated stimulation pads and lane adapters.

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3. Place each mouse in a dedicated lane and let them explore the new environment for a few minutes without any motion of the belt. 4. Start the treadmill under continuous observation. Although mice will spontaneously adapt to the system and start running, occasionally some assistance may be required to help an animal learn the desired behavior. In the extremely rare case where an animal gets injured, the experimenter should immediately stop aversive stimulation of the particular lane to allow for immediate rest at the rear of the belt and/or stop the belt and remove the animal before resuming the test.

5. After the treadmill test is finished, remove animals from the lanes and put them back into their housing cages. 6. Record the number of aversive stimulations received. 7. Clean all parts of the treadmill that were in contact with an animal with distilled water and remove feces and urine. 8. Disinfect the cleaned zones (e.g., with 0.75% lysoform). 9. Dry all parts prior to reuse of the treadmill. 10. Repeat daily until the vast majority of animals have reached a satisfactory level of familiarization to the apparatus (e.g., C57BL6/J mice should receive less than five electrical stimulations in the 5 min of the session).

Conduct performance tests 11. Set treadmill parameters for the one of the following tests: a. Power test: Belt speed: 15 cm/sec initially, increase 2 cm/sec every minute. Slope: +5◦ during the entire protocol.

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Duration: Until termination criteria are reached (prepare for 40 min when the treadmill is computer controlled). Aversive stimulation: mild electrical stimulation (e.g., 0.3 mA) or any other validated stimulation (see Critical Parameters). b. Endurance test: Belt speed: 15 cm/sec initially, increase 3 cm/sec every 12 min. Slope: +5◦ during the entire protocol. Duration: Until exhaustion criteria are reached (prepare for 4 hr when the treadmill is computer controlled). Aversive stimulation: mild electrical stimulation (e.g., 0.3 mA) or any other validated stimulation (see Critical Parameters). 12. Place each mouse in a dedicated lane. 13. Start the treadmill and run the test under continuous observation. 14. When the defined termination criteria are met (i.e., the mouse can no longer sustain exercise at the given speed), stop the belt and immediately allow the animal to stop running (i.e., by removing it from the treadmill or stopping the aversive stimulation and letting it rest at the rear of the belt). Examples of typical termination criteria are a stimulation rate higher than 20 electrical stimulations per minute or failure to re-engage on the treadmill despite aversive stimulation for more than 15 sec.

15. Record the running time, distance traveled, total number of aversive stimulations received, and maximum speed reached by each animal. 16. When all animals have reached the termination criteria, stop the treadmill and transfer animals back to their housing cages. 17. Clean all parts of the treadmill that were in contact with an animal with distilled water and remove feces and urine. 18. Disinfect the cleaned zones (e.g., with 0.75% lysoform). 19. Dry all parts prior to reuse of the treadmill.

Analyze data 20. Export the cumulative number of stimulations as a function of time and plot this for every mouse to validate that the termination criteria are correct. 21. Export the distance covered, the total running time and the maximal tolerated speed for each mouse, and plot the mean and error for each group. BASIC PROTOCOL 2

Exercise Performance Tests in Mice

VO2 max DETERMINATION WITH A TREADMILL COUPLED TO INDIRECT CALORIMETRY The coupling of treadmill running to an open circuit calorimeter (indirect calorimetry) allows assessment of maximal oxygen consumption (VO2 max) in mice. The concentration differences measured between the air of the chamber versus ambient air, along with the flow information, are used to compute oxygen consumption (VO2 ), carbon dioxide production (VCO2 ), and the respiratory exchange ratio (RER), defined as VCO2 /VO2 ratio. O2 and CO2 measurements are taken at regular intervals of time during the whole experiment. VO2 and VCO2 values are expressed as ml/kg/min after normalization for the body weight of the animal. However, the cost of such an approach in terms of time and equipment limits its extensive use.

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ambient air supply fans air sampler

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Figure 3

Example of a commercial metabolic treadmill setup for mice (Columbus Instruments).

Materials Appropriate mouse strain (e.g., C57BL/6J) and housing facility Spray bottle of distilled water Mild cleaning agent for cleaning the belt, e.g., 0.75% lysoform A metabolic treadmill (see, e.g., Fig. 3) consisting of a chamber tightly closed at both ends by removable walls (approximate volume, 2 liters); the chamber encompasses: A revolving belt with adjustable speed (0 to 120 cm/sec) adapted to the size of the mice (approximately 50 mm width and 26 cm length) A system at the rear of the belt to encourage animals to run during the test (e.g., electrical stimulation grid with adjustable intensity) An inlet port connected to a pump and an air-flow controller to ensure chamber ventilation with ambient air at a constant flow An outlet port connected to an air sampler delivering the air inside the chamber to the gas analyzer at regular intervals A fan at the front of the belt to ensure circulation of the air over the animal Ability to adjust the slope of the entire setup in 5◦ increments from −10◦ to +25◦ : commercial systems have been developed, e.g., by Columbus Instruments (http://www.colinst.com/) and TSE (http://www.tse-systems.com/) An open-circuit calorimeter (indirect calorimetry) (e.g., Oxymax, Columbus Instruments, http://www.colinst.com/) to monitor changes in gas concentration (O2 and CO2 ) in the air of the chamber; ambient air is used as a reference after calibration of the system with a calibration gas (20.5% O2 /0.5% CO2 with remainder N2 ) Computer and software communicating with the open-circuit calorimeter to define the settings of the experiment (air flow, sampling time, etc.) and to display measurements in real time (VO2 , VCO2 , and RER) throughout the experiment; ideally, the same software should allow to control and record the treadmill parameters Current Protocols in Mouse Biology

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Calibration gas (high-purity grade): exactly 20.5% O2 and 0.5% CO2 with remainder N2 (custom prepared by gas supplier) Scale to monitor body weight of the mouse Timer NOTE: To ensure optimal test results, a few sessions of familiarization to the setup are required every day preceding the actual treadmill test. For C57BL6/J mice 1 to 2 sessions is typically sufficient, but this number should be adapted to the particular strain of mice under consideration. 1. Turn on the system at least 2 hr before the experiment to allow warm-up and stabilization of the gas sensors. 2. Prior to the start of the experiment, calibrate the O2 and CO2 sensors according to the operating manual. Turn on the valve of the calibration gas tank connected to the gas analyzer. Follow the instructions provided by the graphical user interface of the software. Once calibration is completed, i.e., O2 and CO2 concentrations are set up in accordance with the concentrations of the calibration gas, turn off the valve. 3. Record the body weight of the mouse. 4. Configure the experiment. Experimental settings may vary according to the size of the metabolic treadmill and the indirect calorimetry system used. The settings given here refer to the requirements of Oxymax system from Columbus Instruments. a. Select the chamber (only one chamber for VO2 max protocols). b. Enter mouse identification information. c. Enter mouse body weight. d. Set the air-flow at 0.5 liters/min (for a chamber with a 2-liter volume). e. Configure the measurement settings as follows: Continuous measurements until manual stop of the gas measurements. One reference measurement with ambient air at the initiation of the protocol. The reference is measured for 30 sec. The purging time before and after the reference measurement is 60 sec. Since the reference is measured only once at the beginning of each session, it is important to work in a well ventilated environment (large room, mechanical ventilation) to avoid changes in O2 and CO2 concentration in ambient air due to the presence of the operator.

f. Measure gases from the chamber every 10 sec. g. Configure the ramping protocol of the belt speed as follows: Initial belt speed: 0 cm/sec for 5 min. Warm-up: 8 cm/sec for 5 min. After the warm-up phase, increase belt speed as follows: 15 cm/sec for 5 min 20 cm/sec for 5 min 25 cm/sec for 5 min Then increase belt speed by 3 cm/sec every 2 min. 5. Place the mouse in the chamber and close each end. 6. Start the timer and leave the mouse for 5 min without starting the protocol (gas measurement and belt off). Ensure that the air flow is on. Exercise Performance Tests in Mice

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This period allows the balancing of the gas concentration inside the chamber.

7. After this 5-min period, start the protocol (gas measurement on and ramping protocol on) with the graphical user interface of the software. Current Protocols in Mouse Biology

8. Turn on the stimulation device and mildly increase the intensity of electric stimulation until the mouse becomes sensitive (or use a prevalidated standard value). 9. Keep the mouse under continuous observation throughout the test. 10. Using the graphical user interface, display on the screen the measurement of VO2 and respiratory exchange ratio (RER) (see Fig. 4 for example).

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Figure 5 Individual oxygen consumption and respiratory exchange ratio (RER) during maximal oxygen consumption (VO2 max) protocol. 24-week-old male C57BL/6J mice were subjected to a VO2 max protocol with +10◦ treadmill angle. Left panel: Oxygen consumption (VO2 ) at each belt speed (n = 8). Right panel: Respiratory exchange ratio at each belt speed (n = 8). Exercise Performance Tests in Mice

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11. After 5 min at 0 cm/sec, increase the slope to +10◦ for the rest of the protocol. 12. Allow the animal to stop running when at least one of the defined termination criteria is met (i.e., it refuses to run or the RER rapidly increases; see Critical Parameters). 13. When the mouse has reached the termination criteria, stop the treadmill and transfer the animal back to its housing cage. 14. Record the running time and distance traveled. 15. Clean all parts of the treadmill that were in contact with an animal with distilled water and remove feces and urine. 16. Disinfect the cleaned zones (e.g., with 0.75% lysoform). 17. Dry all parts prior to reuse of the treadmill. 18. Export VO2 and RER measurement data per interval of time (1 interval = 10 sec). 19. For each mouse, calculate the average VO2 and RER for each belt speed. Plot a graphical representation of the data with VO2 or RER on the y axis and belt speed on the x axis (belt speed represents the intensity of the exercise). 20. Calculate the VO2 max for each mouse (Fig. 5). VO2 max is determined as the VO2 value when the mouse reaches the termination criteria defined in step 12.

COMMENTARY Background Information

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Despite an obvious connotation to a wide readership, the actual definition of exercise remains very broad, with many different interpretations of the term. Such differences arise from variations in duration, intensity, aim (acute performance versus training), and number of bouts, as well as metabolic and cardiovascular components of the exercise under consideration. Overall, exercise is defined as a broad spectrum of conditions through which an individual produces mechanical work over its basal level of activity. With this very broad definition, the authors fully agree with the recent viewpoint stating that the term exercise by itself is much too broad (Booth et al., 2010) and should always be used in conjunction with a definition of the type of exercise being discussed (e.g., endurance, resistance, strength, performance, training). Numerous procedures have been developed to assess various types of performance and to analyze the underlying key physiological parameters. For example, muscle contraction can be evaluated by measuring twitch and tetanic force in response to direct exogenous electrical stimulation of the afferent nerve, and cardiovascular phenotyping allows assessment of parameters such as heart rate, blood pressure, cardiac structure, and extraction volumes noninvasively. However, exercise performance involves the coordination of multiple physiolog-

ical parameters controlled by cardiovascular, respiratory, metabolic, and central cues, making global exercise modalities such as running or swimming better models of global performance. Despite the fact that swimming is not a natural mouse behavior, several protocols have been successfully developed to force mice to swim in water tanks both for training and performance purposes (Dawson and Horvath, 1970; Kregel et al., 2006). Maximal performance determination using swimming as an exercise modality is, however, hampered by the fact that it is difficult to define objective and humane exhaustion criteria. Treadmill running is therefore the widely accepted gold standard to evaluate maximal exercise performance in mice. During exercise, oxygen consumption (VO2 ) increases in order to adapt oxygen supplies to the needs of skeletal muscles. During the first phase of the exercise, VO2 increases proportionally to the intensity of the exercise. Beyond a certain degree of effort, named maximal aerobic power, VO2 cannot increase anymore despite the continuous increase of the workload. This stage corresponds to the maximal oxygen consumption or VO2 max. This parameter has been and is still extensively used in the clinic to investigate human cardiovascular and pulmonary disorders. VO2 max is tightly linked to cardiac, pulmonary, and muscle function. This parameter reflects the contribution Current Protocols in Mouse Biology

of the various steps from breath to oxidative phosphorylation inside muscle mitochondria, i.e., the diffusion of oxygen in pulmonary capillaries, its fixation in hemoglobin, its transport by the cardiovascular system, its diffusion towards myoglobin in skeletal muscles, and, finally, its use by the mitochondrial respiratory chain. When oxygen consumption is assessed as a function of exercise intensity in animal models, it is often difficult to reach an actual saturation of oxygen consumption, as the test is generally terminated by the animal refusing to run. VO2 max is therefore very often referred as VO2 peak, i.e., the peak of oxygen consumption for the animal’s maximal performance (see Fig. 5).

Critical Parameters Treadmill setup and experimental organization There are multiple commercial and userdesigned treadmill setups. The major technical parameters to be considered have been summarized in Basic Protocol 1. However, one critical parameter when testing large experimental groups is the number of lanes available. Typically, classical setups have five to six lanes, thereby making the procedure relatively low throughput, especially for endurance tests that can last up to a couple of hours. Whenever possible, it is therefore recommended to have several independent treadmills. An alternative solution is to run tests sequentially and/or to stagger experimental groups over multiple days. Under such circumstances, particular attention should be paid to potential confounding factors, such as the time of the day when the test is performed or the time elapsed since the last procedure, in order to minimize experimental variability. In addition, each session should include animals from the different experimental groups to avoid generating biased results. Technical considerations regarding gas analyzers for VO2 max protocols O2 and CO2 concentration are analyzed independently. Three technologies exist to measure oxygen: 1. Polarographic oxygen electrodes. They have a short half-life and can be unstable. They are inexpensive and have a 5% to 22% oxygen operating range. 2. Zirconium oxide–based sensors. These sensors are fast, accurate, and stable, and do not require intensive maintenance but are quite delicate. These sensors require a long warm-up phase before use and consume a lot of energy.

They have a full 5% to 100% oxygen operating range. 3. Paramagnetic analyzers. They are fast and stable and do not require a lot of maintenance. They have a full 5% to 100% oxygen operating range, but are only linear from 0% to 25% oxygen. They are expensive. Most of the systems available for indirect calorimetry in mice use zirconium oxide– based sensors (e.g., Oxymax, Columbus Instruments). Carbon dioxide sensors employ a single, nondispersed infrared beam. These sensors are extremely accurate, but are moisture sensitive. All analyzers are sensitive to the water vapor present in the expired gases. Therefore, a drying system is associated with the gas analyzer (Nafion tubes). Nafion tubes (e.g., Perma Pure, http://www.permapure.com) allow the diffusion of water vapor across their walls before the air from the chamber enters the analyzer. Nafion tubes need to be changed on a regular basis according to the extent of use of the system. Aversive stimulation for forced running Although mice are naturally prone to run, an aversive stimulation is required to force all animals to stay engaged in the test. This is typically achieved using mild electrical stimulation on a dedicated grid at the rear of the revolving belt. The intensity of the electrical stimulation should be optimized to the lowest possible level forcing the animal to re-engage on the belt without causing major distress (i.e., the mouse should not display signs of pain such as jumping or sudden high running speed). A constant current of 0.2 to 0.4 mA is generally sufficient to stimulate most strains of mice to run. Repeated treadmill testing may, however, cause de-sensitization of the animals and require slight increases in the intensity of the aversive currents. In all cases, electrical stimulation should always be balanced with the general welfare of the animal, and used with strict adherence to local veterinarian recommendations. Alternative aversive stimulation strategies have also been described. These mainly include pulses of air blown on the mouse when it crosses an infrared beam located at the rear of the belt, or manual prodding of the animal by the experimenter when it rests on the platform at the rear of the belt. All these techniques have advantages and drawbacks depending on the desired throughput of the test (i.e., number of animals analyzed in parallel), the reproducibility of the stimulation, the technical setup of the treadmill, and

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the level of stress imposed to the animal. The final choice of the aversive stimulus therefore remains to be defined by the experimenter according to their preferred options. Familiarization with the setup To minimize the number of aversive stimulations and to eliminate inherent bad runners (see below), animals should always be familiarized with the treadmill prior to the performance test. This can be achieved by a few short daily sessions at low speed. Typically, the number of aversive stimulations required to keep animals running during each familiarization session is used as a readout of adaptation to the instrument and should stabilize at low levels prior to initiation of the performance test. For C57BL6/J mice, one to two sessions of 5 min are sufficient to minimize the number of electrical aversive stimulations below five per session (Fig. 2), but the actual number of familiarization bouts needs to be validated for each mouse strain or transgenic model.

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Criteria to terminate the test Since the time spent running, the distance covered, and the maximal tolerated speed are the primary readouts used to assess running performance, it is critical to define objective criteria to terminate the test when the animal has reached its maximal performance. A number of parameters, such as the inability of the animal to right itself, low blood glucose, or glycogen depletion, have been proposed to characterize metabolic exhaustion when testing for endurance (Booth et al., 2010). These parameters are, however, not amenable to real-

time testing during the exercise test. The typical alternative is to use the number of aversive stimulations and/or the time spent unsuccessfully trying to re-engage on the treadmill as surrogate readouts of maximal performance. With appropriate habituation to the experimental setup and exclusion of inherent bad runners, the average electrical stimulation rate should typically remain very low during most of the test, until a clear terminal exponential increase of the stimulation rate indicates that the animal is reaching its maximal capacity (Fig. 6). During this phase, the experimenter will visually observe that the animal experiences difficulties when trying to re-engage on the belt and may rest for prolonged periods despite aversive stimulation. Although not available on all treadmills, it is highly recommended to have a real-time control of the aversive stimulation rate in order to have fully objective criteria to define maximal performance (e.g., by direct software readout, although this is not a classical parameter and may need to be programmed on request, or by controlling treadmill speed with 1-min intervals over which the number of stimulations can be integrated). When this is not possible, the experimenter can, however, resolve to use the failure to re-engage on the belt despite gentle manual prodding or continuous resting for more than 15 sec as interruption criteria. For VO2 max protocols, two termination criteria are usually defined (see below). The experiment is stopped when at least one of those is met.

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Criterion 1. Respiratory exchange ratio (RER) is ≥1, or dramatically increases within a short period of time (4 to 6 min) and then reaches a plateau. It is important to keep in mind that in the context of exercise protocols, RER is not equivalent to the respiratory quotient (RQ). RQ reflects substrate oxidation (RQ = 1 for carbohydrate oxidation; RQ = 0.82 for protein oxidation; RQ = 0.7 for lipid oxidation; and RQ≈0.85 with a balanced diet). Under stable conditions, RQ is equivalent to RER and cannot go above 1 (carbohydrates as the sole energy substrates). In the context of exercise, the RER increases progressively with the intensity of the exercise, and nonmetabolic CO2 is produced from intracellular buffering of protons by HCO3 – in the muscles (Peronnet and Aguilaniu, 2006). Criterion 2. The mouse is exhausted and refuses to run. This criterion is the most commonly reached. Exclusion criteria The inter-animal variability of exercise performance suffers from the fact that some animals are intrinsically bad runners, as they generally do not comply with the intended technological systems used to record performance. This is particularly problematic with forced treadmill performance tests where the aversive stimulation to force animals to run is inefficient in 5% to 10% of the mice, even after the initial adaptation to the apparatus as described in the protocol. To eliminate these true outliers resulting from the inability to adapt properly to the system and thereby minimize experimental variability, it is recommended to define objective exclusion criteria. Typically, these bad runners are easily identified during the initial sessions of familiarization to the setup and are excluded from the actual performance tests. Group size Exercise-based readouts have inherently relatively high levels of inter-individual variability, as performance is a multi-factorial process influenced by metabolic and cardiovascular as well as central effects. Consequently, group size should be sufficiently powered to allow meaningful statistical analyses and adapted to the variability of the exact experimental context (strain of mice, equipment, etc.). As a general recommendation, 10 to 12 mice per group yield robust and reliable results. Incline Running capacity is generally optimal at a slight uphill angle (e.g., 5◦ ). Experimenters wanting to make the tests more stringent can,

however, increase this angle up to 20◦ , which is well tolerated by wild-type mice despite an obvious decrease in running time and distance. In contrast, downhill running can be used to model exercise-induced muscle damage, as the alterations of muscle fibers will be exacerbated by gravity. For the VO2 max protocol, a slight uphill angle is also optimal to maximize performance (Kemi et al., 2002). A +5◦ to +10◦ angle is usually suitable for wild-type sedentary mice, but when studying trained mice, the angle should be increased to +15◦ or +20◦ to increase the intensity of the exercise and reach VO2 max faster. By contrast, with obese mice (genetically or diet-induced), it is recommended to maintain the incline of the belt at 0◦ . Time between tests and number of repeats Power performance tests consist of a short bout of high-intensity running from which mice can recover rapidly. Consequently, test/retest performance in the power paradigm are not affected when the two tests are repeated within a close interval (down to a few hours). It is therefore possible to repeat a performance test 3 to 4 hr, ideally 1 day, after a power test. The situation is, however, very different when performing endurance tests, as these long sessions induce profound metabolic challenges and physiological adaptations, and require much longer periods for the animal to recover. Therefore, it is typically recommended to let animals recover for at least 7 days after an endurance performance test. When designing longitudinal studies, one particularly critical parameter is the total number of performance tests and their frequency, as animals may desensitize to the aversive stimulations and learn to escape from the test prior to reaching maximal performance. The authors therefore recommend minimizing the total number of tests and validating that control animals from the strain of interest display constant performance in the longitudinal protocol under consideration. Impaired animals Some animals may be severely impaired in their running capacity by other experimental procedures (e.g., treatment or surgery) or by general alterations of morphology, behavior, coordination, or metabolism induced by genetic manipulations. Under such circumstances, the decision whether to conduct a performance test should be made based on the severity of the phenotype, the possibility of interpreting results with potential confounding factors, and ethical and general welfare

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considerations. In particular, special care should be taken when studying factors affecting the locomotor system and morphological alterations of feet and claws, as these may cause injuries during the tests. Circadian rhythm Mice are nocturnal animals that have circadian and metabolic rhythms synchronized for high spontaneous activity during the dark phase. Although this is typically inconvenient, some researchers recommend conducting exercise performance tests during this active nocturnal phase. The authors have, however, not observed any differences in power or endurance performance when comparing mice on regular day/night cycles to mice housed for over 2 weeks with day/night inversion and tested for running capacity in the dark.

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Genetic background Since exercise capacities are multifactorial and depend on various organs and physiological systems, sex as well as the genetic background will influence performance (Bernstein, 2003; Konhilas et al., 2004; Billat et al., 2005). For example, C57BL/6J mice have levels of performance amenable to reliably assessing over- and under-performance, but are outcompeted by other strains of mice such as DBA or BALB/c (Lightfoot et al., 2001; Lerman et al., 2002). In addition, we have observed that the response to aversive stimulation is very different in BALB/c than in C57BL/6J mice. It is therefore recommended to use mouse lines on a genetically pure background and to always match experimental groups from the same sex, strain, breeder, and housing environment. In

addition, different strains of mice respond differently to the treadmill profiles and aversive stimulations. Protocols described herein have been optimized for C57BL/6J mice but may require some adaptation for other stains of mice.

Anticipated Results Typical standard values for maximal treadmill performance of wild-type C57BL6/J mice are presented in Figure 7. Power tests consist of a short (20 min on average) run, which rapidly reaches high levels of intensity (55 cm/sec on average). As expected, the tolerable duration of running is much longer with the less intense endurant exercise in which running time reaches approximately 2 hr and 2 kilometers. Both exercise performance and VO2 max are significantly affected by the health status of the animal. For example, diet-induced obese mice display a significant reduction of both the distance traveled and VO2 max during maximal oxygen consumption protocols, while mice trained by spontaneous wheel running (see Thomas et al., 2011 for details) significantly improve their maximal running distance and their VO2 max (Fig. 8).

Time Considerations Familiarization to the exercise testing equipment is generally achieved in 1 to 2 days, but specific strains may require slightly longer adaptation periods. The average duration of power and endurance tests are shown in Figure 7. When planning the logistical aspects of the test, the

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Figure 8 VO2 max and running distance of sedentary and trained lean and diet-induced obese mice. 8 week old male C57BL/6J mice were fed with a standard chow diet (CD) or a high-fat diet (HFD) (60% kcal fat) for 16 weeks (n = 8 per group). These “sedentary” mice were subjected to a VO2 max measurement with a +10◦ angle for CD-fed mice and 0◦ angle for HFD-fed mice. Upon completion of this experiment, mice were maintained on these diets and allowed free access to a running wheel for 10 days. After this period, a second VO2 max protocol was performed with these “trained” animals under the same condition (+10◦ angle for CD-fed mice and 0◦ angle for HFD-fed mice). Data are represented as mean ± SEM and * represents a statistical significant difference (p<0.05) using a 1-way ANOVA followed by a Bonferroni test.

experimenter should not forget that these average values have coefficients of variation of ∼15%, and that the test cannot end before the best performer has reached maximal performance. In addition, there are numerous examples showing that training or pharmacological and genetic manipulations can enhance performance. Given that the number of animals to test generally exceeds the number of lanes available, the experimenter should design the experimental plan accordingly and stagger the testing over multiple days when required.

VO2 max protocols last for ∼35 to 45 min per mouse, but the overall duration of the experiment is generally long because classical experimental setups are limited to one or a low number of lanes due to cost constraints.

Literature Cited Bernstein, D. 2003. Exercise assessment of transgenic models of human cardiovascular disease. Physiol. Genomics 13:217-226. Billat, V.L., Mouisel, E., Roblot, N., and Melki, J. 2005. Inter- and intrastrain variation in mouse critical running speed. J. Appl. Physiol. 98:12581263.

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Booth, F.W., Laye, M.J., and Spangenburg, E.E. 2010. Gold standards for scientists who are conducting animal-based exercise studies. J. Appl. Physiol. 108:219-221. Dawson, C.A. and Horvath, S.M. 1970. Swimming in small laboratory animals. Med. Sci. Sports 2:51-78. Kemi, O.J., Loennechen, J.P., Wisloff, U., and Ellingsen, O. 2002. Intensity-controlled treadmill running in mice: Cardiac and skeletal muscle hypertrophy. J. Appl. Physiol. 93:13011309. Konhilas, J.P., Maass, A.H., Luckey, S.W., Stauffer, B.L., Olson, E.N., and Leinwand, L.A. 2004. Sex modifies exercise and cardiac adaptation in mice. Am. J. Physiol Heart Circ. Physiol. 287:H2768-H2776. Kregel, K.C., Allen, D.L., Booth, F.W., Fleshner, M., Henriksen, E.J., Musch, T.I., O’Leary, D.S., Parks, C.M., Poole, D.C., Ra’anan, A.W., Sheriff, D.D., Sturek, M.S., and Toth, L.A. 2006. Resource Book for the Design of Animal Exer-

cise Protocols. American Physiological Society, Bethesda, Md. Lerman, I., Harrison, B.C., Freeman, K., Hewett, T.E., Allen, D.L., Robbins, J., and Leinwand, L.A. (2002). Genetic variability in forced and voluntary endurance exercise performance in seven inbred mouse strains. J. Appl. Physiol. 92:2245-2255. Lightfoot, J.T., Turner, M.J., Debate, K.A., and Kleeberger, S.R. 2001. Interstrain variation in murine aerobic capacity. Med. Sci. Sports Exerc. 33:2053-2057. Peronnet, F. and Aguilaniu, B. 2006. Lactic acid buffering, nonmetabolic CO2 and exercise hyperventilation: A critical reappraisal. Respir. Physiol. Neurobiol. 150:4-18. Savaglio, S. and Carbone, V. 2000. Scaling in athletic world records. Nature 404:244. Thomas, C., Marcaletti, S., and Feige, J.N. 2011. Assessment of spontaneous locomotor and running activity in mice. Curr. Protoc. Mouse Biol. 1:185-198.

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Ethical Considerations in Mouse Experiments Bernard Baertschi1 and Marcel Gyger2 1 2

Institute for Biomedical Ethics, University of Geneva, Geneva, Switzerland EPFL—Center of Phenogenomics, Lausanne, Switzerland

ABSTRACT Mice count morally because they can be harmed. This raises a moral issue in animal experimentation. Three main ethical attitudes towards animals are reviewed here. The Kantian view denies moral value to animals because they lack reason. The second view, by Singer, considers animals as sentient creatures (i.e., able to suffer). Finally, Regan considers that animals are subjects of their own life; they are autonomous and therefore have moral rights. Singer is a reformist and allows animal experimentation under certain conditions. Regan is abolitionist, saying that animals have moral rights that cannot be negotiated. Current animal protection legislation strives to put in balance the human and animal interests to decide whether an animal experiment is morally justified or not. An ethical evaluation process is conducted based on the harm-benefit assessment of the experiment. The researcher has to implement the 3Rs (Replacement, Reduction, Refinement) to minimize the harms to the animals and make sure that the outcomes are scientifically significant and that the quality of the science is high, in order to maximize benefits to humans C 2011 by John Wiley & Sons, Inc. and animals. Curr. Protoc. Mouse Biol. 1:155-167  Keywords: ethics r mouse r animal experimentation r 3R

INTRODUCTION Why should we introduce ethical considerations into experiments with mice? The fundamental reason is that humans interact with animals differently than with objects, plants, or microbes. For example, when I walk along a path in the wild, I often kick a stone inadvertently. I do not bother about it (except if the stone could hurt somebody). I may pick up a bunch of weed to clean my shoes. I do not bother about that, either. But if I step on a tiny frog and hurt it, I grieve. Why? Because frogs matter to humans, contrary to stones and grass. Frogs matter because they are living beings, and because we tend to think that harming an animal is something we should avoid whenever we can. In short, animals do morally count for us, and it is not permissible to behave with respect to them just as we choose. Animals morally count because they can be harmed, but how much do they count? Human beings count, too, and sometimes, we meet with conflicts between animal and human interests. This is the case not only in animal experimentation: we eat farm animals; we destroy pests. Even with pets: can we say with confidence that we always act in their inter-

ests (think of castration)? Animals and human beings are related in many ways, and their interests interrelate in multifarious ways. In this overview, we will examine the case of mouse experiments. Laboratory mice are used all over the world and are the most frequently used laboratory animals in many countries. The principal reason is that, thanks to the particular reproductive biology of this species, mouse genetics is easy to manipulate. We will put our inquiry in the larger context of animal experimentation and the requirements that animal ethics exert upon on it. We will briefly discuss whether genetic manipulations are ethically defensible or not. Definitions of some of the philosophical terms used in this article are provided in a glossary in the appendix at the end of the text.

ANIMAL’S MORAL STATUS: SEVERAL THEORIES Moral status, intrinsic and extrinsic properties, and properties and values Stepping on a stone is not the same as stepping on a frog because we feel responsible for having “hurt” a living organism. In ethical terms, animals have a moral status. What is it,

Current Protocols in Mouse Biology 1: 155-167, March 2011 Published online March 2011 in Wiley Online Library (wileyonlinelibrary.com). DOI: 10.1002/9780470942390.mo100161 C 2011 John Wiley & Sons, Inc. Copyright 

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more precisely, to have a moral status? Mary Anne Warren states that: “To have moral status is to be morally considerable, or to have moral standing. It is to be an entity towards which moral agents have, or can have, moral obligations. If an entity has moral status, then we may not treat it in just any way we please” (Warren, 1997). Animals deserve moral consideration; therefore, we have moral duties toward them. But why? Giving a general answer to this question is easy: the moral status of an entity depends on certain of its intrinsic properties. They are not to be found “outside” of the object. My dog, born on June 9, 2000 in La Sarraz, with name Bobby, blue eyes, and black and white fur, has intrinsic properties such that we may relate to its feelings and capacity to suffer; however, my dog has also extrinsic properties or relational ones because this particular dog matters to me, as I have spent many years with it, shared many treks in the mountains, etc. Warren (1997) characterizes both properties as follows: “A thing’s intrinsic properties are those which it is logically possible for it to have had were it the only thing in existence. Its relational properties are those that it would be logically impossible for it to have had were it the only thing in existence.” And, as some intrinsic properties confer intrinsic value, some relational ones confer relational value. For example, ‘to be loved’ is a relational property that confers relational value upon the entity that is loved (the object of love); ‘to be hated’ brings relational disvalue. As this example shows, relational value can combine with intrinsic value to enhance the general value of the entity, or diminish it if it is a negative value (a disvalue). As we will see, the very different attitudes that we adopt toward mice, considered as laboratory animals, pets, or pests, do not depend on their intrinsic properties (all mice have the same relevant moral intrinsic properties) but on their relational ones. Intrinsic properties, as opposed to extrinsic properties, have to be found in the object itself. Going back to our first example about stones, weeds, and frogs, the latter is a living entity which is clearly different from a stone or other nonliving objects. A frog also has something more than weeds or other plants. What makes animals different from the rest of the world? Here philosophers and ethicists have different views of these properties; for Singer, the key property is sentience, for Regan, a kind of subjectivity (see below for more details).

Intrinsic properties like sentience or subjectivity are constitutive of moral status because they possess intrinsic value; beings who have moral status possess a value by and in themselves, and we ought to respect them because of this value. Regarding human beings, this intrinsic value is often named dignity. Aquinas said long ago: “Dignity means the goodness [or value] a thing possesses because of itself; utility, because of another thing” (Scriptum super Sententiis, liber 3, d. 35, q. 1, a. 4, q. 1, c. Available at: http://www. corpusthomisticum.org/iopera.html; accessed 7 May 2010). Moral status of an object takes roots in its intrinsic value that originates in its intrinsic properties. The moral status of mice and other animals calls for moral duties towards them. Are these duties identical to the ones we have towards humans or are they different? If they were identical, most studies on mice could not be possible; most animal experimentation should be banned. How different then are duties towards animals and humans? To answer this question, we have to go back to the relevant intrinsic properties of animals and humans. Those properties are identified on the basis of several predicates, such as those listed in Table 1 (Beauchamp and Walters, 1989). According to Pucetti, says LeRoy Walters, “the S*-predicates in the left-hand column can be applied to conscious nonpersons like dogs, whereas the R*-predicates in the right-hand column presuppose the possession of a conceptual scheme and the capacity to act as a moral agent. This latter capacity is, for Pucetti, the primary distinguishing feature of personhood, for persons are the only conscious entities who can adopt moral attitudes toward moral objects” (Beauchamp and Walters, 1989). In short, the moral status of these beings depends on what they intrinsically are, respectively, beings with properties sustaining S*—animals—and beings with properties sustaining R*—human beings. In comparing human beings and animals, we have highlighted two sets of properties. Does it mean that there exist at least two different moral statuses? Certain authors think like that, but others, strictly following the conception put forward by Warren, claim that there is only one moral status, the possession of which determines moral duties: “If an entity has moral status, then we may not treat it in just any way we please.” If we consider the objects that have moral status, their moral status is the same.

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Table 1 Morally Relevant Intrinsic Properties

S*-predicates/animals

R*-predicates/humans

To be in pain

To want to secure justice

To feel hungry

To summarize the point nicely

To be excited

To be an astute judge of character

To be afraid of you

To be a smug hypocrite

Traditionally only humans had been considered possessing moral status.

Indirect duties theories Human beings, and only them, have moral status, because they possess the only intrinsic property that has moral relevance: reason (autonomy, self-consciousness, self-mastery,. . . ). Kant is very clear on this point: “The fact that man is aware of an ego-concept raises him infinitely above all other creatures living on earth. Because of this, he is a person [. . . ]. He is a being who, by reason of his pre-eminence and dignity, is wholly different from things, such as the irrational animals, which he can master and rule at will.” (Kant, 1978; originally published 1798). Animals have no moral status because they are lacking in rationality; therefore they are things, that is, entities that can be possessed, sold or destroyed. We “can master and rule [animals] at will” says Kant. Does it mean that we can treat them with violence and cruelty? Kant answers clearly no. However the reason he gives has nothing to do with a wrong committed against them. Animals have no moral status; therefore they do not belong to the moral community, which comprises the set of beings who have moral duties toward each other. Animals can be harmed; they cannot be wronged. The reasoning against violence and cruelty thus consists in a duty toward human beings themselves: if I harm an animal without a good reason, I behave myself in a wrongful manner, because I manifest a vice of character—to be violent or cruel is a breach to a duty toward oneself, the duty to be gentle and benevolent— and the consequence thereof is that I will probably be violent and cruel toward my fellow human beings. For this reason, Kant condemns experiments with animals when they are unnecessary and done only for the sake of knowledge (Kant, 1996; originally published 1797). Such a conception was often embodied in the law before the 20th century. For example, in France, the Grammont Law (enacted in 1850) forbade bad treatment of animals, but

only in a public place. The reason is not that the animal would be wronged, but that passersby could be shocked by the cruel deeds. In brief, for this conception, we do not have direct moral duties toward animals, but only indirect ones: animals must not be harmed because it would wrong human beings (the doer and the observer).

Direct duties theories For indirect duties theories, the protection of animals is grounded on relational properties, i.e., the effects our deeds have on human beings. Nowadays, the great majority of our regulations are grounded on direct duty theory. It is forbidden to treat animals with violence and cruelty because such acts wrong their victims. Therefore, for those theories, animals possess a moral status; they have intrinsic properties asking for respect and protection. Utilitarian position (Singer) The first and most well-known author who has endorsed a direct duty theory is the philosopher Peter Singer. He followed Jeremy Bentham when he stated: “The question is not, can they reason? Nor can they talk? But, can they suffer?” (quoted in Singer, 1979). The property relevant for the possession of a moral status is not reason but sentience, i.e., the capacity for suffering and for happiness. Human beings can suffer or be happy, but they are not alone; animals also can. Therefore, animals belong to the moral community: they count morally because their happiness counts. As we can see, Singer, following Bentham (who was following Rousseau on this topic), enlarges the moral community. The relevant characteristic is no longer reason, possessed only by human beings, but sentience, belonging to human beings and animals (probably not all animals, but at least vertebrates). Reason is no longer the central intrinsic property, but sentience. Pathocentrism has replaced anthropocentrism, as we often call those

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conceptions. Singer even compares anthropocentrism to racism, and makes this comment: “Racists violate the principle of equality by giving greater weight to the interests of members of their own race when there is a clash between their interests and the interests of those of another race. [. . . ]. Similarly those, I would call, “speciesists” give greater weight to the interests of member of their own species where there is a clash between their interests and the interests of those of other species.” (Singer, 1979). Human beings tend to be speciesists, even when they put reason to the fore, because reason is a property that, on the earth and in their mind, only their own species possesses; in this sense anthropomorphism is speciesism. If sentience is the relevant moral property, then the interests of all sentient beings have the same importance. Therefore, all animals— human and non-human—are equal. We must give an equal consideration to their interests— the suffering of a mouse has the same value as the suffering of a human being. Does it mean that a mouse has the same moral importance as a human being? Not necessarily: human beings could be built so that their capacity for suffering would be much higher than that of mice. But the reverse could be true, also! It is an empirical question. For the moral point of view, as Singer is an utilitarian, he feels no necessity to enter into those questions: what counts once and for all is the quantity of happiness and of suffering in play. “How bad a pain is depends on how intense it is and how long it lasts, but pains of the same intensity and duration are equally bad, whether felt by humans or animals.” (Singer, 1979). In the case of an experiment with mice, what matters is the suffering of the mice and the benefits for the human beings (and maybe for the mice). Every benefit and every harm or suffering must be put into the balance. If the balance is in favor of human beings, the experiment is allowed; otherwise, it is not (and is immoral if performed in spite of the result of the balancing). To realize a thorough balance of interests is not an easy matter, and we obviously must have recourse to some approximations and evaluations (see “How to undertake the ethical review process,” below). However, ethology and animal psychology continue to progress. As a matter of principle, it must be noted that the Singerian position, which is at the root of the animal liberation movement, does not forbid animal experimentation. It is not an abolitionist position, but a reformist one; animal

experimentation is permissible when the sum of suffering is less than the sum of benefits. This seems reasonable and militates strongly in favor of measures like the three Rs (see “Points that the researcher should consider for a harm-benefit assessment in mouse experiments,” below). Singer’s perspective has its drawbacks too, as it would authorize an experiment performed on unwilling human beings in favor of a greater human happiness. Answers to this drawback of the utilitarian theory exist, but this does not concern us, because our topic of interest is animal, not human experimentation. Rights theory (Tom Regan) Peter Singer’s position is reformist; Tom Regan’s one is abolitionist. He objects to the use of animals for the sake of human interests—animals ought not to be used as mere means for the benefit of human beings; they must not be instrumentalized. What are his arguments? Regan is not a utilitarian; therefore, for him, it is not the capacity to suffer, i.e., sentience, that counts. For him it is Autonomy. “Animals, as individuals who retain their psychological identity over time, have a welfare that is not unrelated to their ability to act autonomously (i.e., as they prefer)” (Regan, 1984). Autonomy is a rational property, and traditionally it has been the basis for the ascription of moral rights. Do animals have moral rights? Regan claims that they have. But how can he justify this claim, since animals are deprived of rationality? To understand Regan’s position, some distinctions are in order. If traditionally, and especially in Kant’s thought, autonomy is a rational property (the property of being able to lead one’s life and to make choices and decisions after having examined the various possibilities) at the root of the various liberties and rights, this kind of autonomy is not the only one. To be autonomous still means to have the capacity to realize one’s preferences: “Individuals are autonomous if they have preferences and have the ability to initiate action with the view to satisfying them” (Regan, 1984). Animals (at least some of them, and in particular mice) do have this property; they are, in this sense, the subject of their lives (Regan, 1984). Animals and human beings are subjects of their own lives; consequently, they possess moral rights. It is easy to understand the reason for that. If a being has preferences he wants to satisfy, if he has projects he wants to realize, if he has a welfare that he cares for, then all those endeavors must be protected, in the

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sense that they must not be interfered with without good reason. Such a being has a moral right to pursue his endeavors, and human utility does not justify a violation of such rights. Of course, animals are not human beings; notably, they are not moral agents (they do not act from a moral point of view), but moral patients (they possess interests with a moral weight). However, since moral patients as well as moral agents are subjects of their lives, they possess rights. Therefore, “it is not an act of kindness to treat animal respectfully, it is an act of justice” (Regan, 1984), i.e., an act of respect for their rights. As animals and human beings are on a par concerning their rights, we understand why Regan is an abolitionist. Since it is immoral to experiment on human beings for the sake of the other human beings without their consent, it is immoral to experiment on animals without their consent. Since they are unable to give their consent, animal experimentation is immoral (like experimentation on children or on mentally handicapped human beings). More precisely, Regan spells three objections against animal experimentation. 1. The right to medical treatment is an acquired right that we have with regard to society or the medical profession. It does not authorize governments to subsidize research that violates the basic rights of animals, or entitle scientists to conduct such experiments (Regan, 1984). 2. “Risks are not morally transferable to those who do not voluntary choose to take them” (Regan, 1984). Life is risky, and it is the function of medical and scientific research work to minimize, even to cancel, those risks. To do so, they must perform experiments, a risky procedure for the subjects involved in them. If those subjects consent to run those risks, they can be morally accepted; if they do not consent (because they don’t want to, or because they are unable to), then this consists in an unacceptable instrumentalization. 3. It is morally impermissible to utilize beings that have an intrinsic value (Regan says “inherent value”) like mere resources for other beings (Regan, 1984). Animals, like human beings, have a moral status; therefore they have an intrinsic value that should be respected. Hierarchical theory There is one feature common to direct duties theories we have examined so far, and to indirect duties theories: their monism or “centrism.” A conception of moral status is monistic when it considers that there exists only one

moral status—either you have a moral status or you have no moral status. Those theories differ with respect to the property that is relevant for the possession of such a status— reason (according to Kant and many authors in our western tradition), sentience (according to Singer and the utilitarians), or subject-of-alife (according to Regan). This property sets a threshold: if you are above, you possess a moral status; if you are beneath, you don’t possess any. Monism breeds ‘centrism’— anthropocentrism (only beings endowed with reason, i.e., human beings, possess a moral status) or pathocentrism (only beings endowed with sentience, i.e., which can feel pleasure and suffer, possess a moral status). There is, however, another way to consider moral status: pluralism. A conception of moral status is pluralistic when it considers that there exist several kinds of moral status, depending on different properties, e.g., reason for persons, sentience for animals, or life for plants. Pluralism does not breed “centrism” but “hierarchism.” It places the different moral statuses on a scale, usually with human beings at the top, animals a little beneath, then plants, and, perhaps, microorganisms. This conception is widespread in our societies. It is often refined with subhierarchies, especially in the realm of animal experimentation. It is better to use mice than chimpanzees, rats than primates, and so on, because animals that are placed higher up on the scale have more intrinsic value. But it is not only folk ethics. The Nuffield Council on Bioethics (1996), for example, claims that it is better to use a swine than a chimpanzee for xenotransplantations. Some authors object that such a view is the remains of a pre-Darwinian conception (the great chain of beings, the scala naturae). But this is not necessarily the case: all depends on the manner in which you conceive the hierarchy. You do not have to have recourse to biological criteria; complexity or other capacities can be appealed to, especially mental ones that are gradual or scalar, e.g., (self-)consciousness.

What makes the difference between a pet mouse, a lab mouse, and a pest mouse? A hierarchical conception explains many judgments that we pass on the use of animals in experiments, but not all. What makes the difference in consideration between a pet mouse, a lab mouse, and a pest mouse? Mice may be considered as almost a family member, as a subject in a medical study, or simply as a

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pest to be destroyed. Although these mice are almost identical in cognitive and emotional performance, and although their capabilities to feel pain and to suffer are certainly not very different—i.e., their intrinsic relevant properties are the same—our relationships to them are however quite diverse, even contradictory. As we have said, our attitudes depend here not on their intrinsic properties, but on their relational ones. Differences in relationships bring with them different ethical considerations. But is it morally justified to assign different moral status to different mice based on their relationships with humans? Following the different direct duty theories we have examined, we ought to say that it is not justified to assign different moral status to different mice (Mason and Littin, 2003; Meerburg et al., 2008). As they have the same morally relevant properties, they have the same moral status. Therefore, their interests must be considered accordingly. The fact that a mouse has a different relationship to us does not change anything with respect to its intrinsic value. Consequently, if we adopt severe norms for animal experimentation, we should accept severe ones for pest destruction. Of course, the consideration of all interests involved may shift the balance (if you are not an abolitionist). Pest mice threaten our interests; lab mice do not. This fact can easily justify a difference in treatment, but not a negligence of the interests of the mice.

An Ethical Step Further: Animal Dignity The special case of the Swiss Constitution and the law for protection of animals Like all countries, Switzerland has a law for the protection of animals (LPA). But in contrast to others, this law is in part based on an appeal to the animal’s dignity—and even to the dignity of creation. In the Swiss Constitution, we read: “The Confederation shall legislate on the use of the reproductive and genetic material of animals, plants, and other organisms. In doing so, it shall take into account the dignity of creation and the security of man, animal and environment, and shall protect the genetic multiplicity of animal and vegetal species.” (Swiss Constitution, art. 120). Ethical Considerations in Mouse Experiments

Animal and human dignity Traditionally, dignity has been a concept restricted to human beings. In the 20th century, it has been more and more used, in the aftermath

of WWII and of the Universal Declaration of Human Rights. We read in its first article: “All human beings are born free and equal in dignity and rights. They are endowed with reason and conscience.” But what does it mean exactly to respect human dignity? Mainly two things, usually expressed in two bans: 1. a ban on instrumentalization, i.e., on utilizing human beings as if they were mere objects; 2. a ban on degrading treatment or humiliation. These two bans are on the forefront in internationally important texts (e.g., Convention for the protection of human rights and dignity of the human being with regard to the application of biology and medicine (Oviedo Convention), and Universal Declaration on the Human Genome and Human Rights proclaimed by UNESCO). These texts are meant to protect human beings against abuses by medicine and biology that reduce them to the status of objects for experiment, treating them as nonpersons (Beyleveld and Brownsword, 2002). The ban on instrumentalization is the most widely voiced of the two bans, but it is in a sense less fundamental than the ban on degrading treatments, because instrumentalization is only one kind of degrading treatment—it degrades a human being to the status of a thing. Nevertheless, if the two bans are kept separate, it is because each points to a different paradigm: the paradigm of instrumentalization is slavery, whereas the paradigm of degrading treatment is torture. Historically, slavery and torture are perhaps the two main domains where human beings have been (and sometimes still are) treated totally disrespectfully. Animals are not human beings; in particular, they are not “endowed with reason and conscience,” two properties at the root of human dignity. Moreover, although intentional torture is condemned when carried out against animals, methods for killing mice as pests are not always soft, nor is instrumentalization. In a sense, animals are our slaves. Pet and lab mice are used uniquely for human goals: a paradigm of instrumentalization. What does it mean then to respect the dignity of animals? For abolitionists, the answer is straightforward—to respect animal dignity is to give up animal experiments and to refrain from interfering with their lives. For those who adhere to other conceptions, the answer will depend on the intrinsic morally relevant property of animals. To possess interest (that can be satisfied or frustrated) is characteristic of sentient animals. It is

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consequently not surprising that this property has been chosen as a ground for moral consideration, i.e., dignity. Therefore, it seems natural to claim that to respect animal dignity amounts to satisfying their interests. More precisely, as the question of respect of dignity arises when there exists a conflict of interests between human interests and animal interests, the Swiss Federal Ethics Committee on Non-Human Biotechnology (ECNH, 2001) has stated: “We disregard an animal’s dignity if we fail to make the possibility of violation the subject of an evaluation of interests, i.e., if we give it no consideration and take it for granted that human interests take precedence” (ECNH, 2001). Human and animal interests must be put in a balance. If our decision follows the tilt of the balance, then we respect animals‘ dignity; if we go on with the experiment contrary to the tilt of the balance, we violate animals’ dignity. The recent Swiss LPA has confirmed this view. It defines dignity as the proper value of an animal and states that there is an infringement of the dignity of the animal when the burden on it cannot be justified by human dominant interests. Under “burden,” we must understand pains or harms, anxiety or debasement, profound modifications of its phenotype or capacities, and excessive instrumentalization (LPA, art. 3). Note that in some other readings of the Swiss law and especially in the application decree, we find also another view. Animal dignity would be per se violated by certain painful or anxiogenic interventions, without any reference to a weighing of interests. Under this second interpretation, it would be lawful and permissible to violate the dignity of animals when important human interests are at stake; under the first (classical) interpretation, the importance of these human interests would prevent the act from constituting such a violation, if the interests of the animal have been taken into account (see Krepper, 2010; Swiss Academies of Arts and Sciences, 2010). To follow the second interpretation will of course favor the belief that animal experimentation is morally—if not legally—dubious.

Does genetic manipulation of the species raise special issues? The answer to this question depends on which stance we adopt. For some authors, genetic manipulation can easily be considered as a violation of dignity. If changes in phenotypic traits count as harms, and if such harms are on the same footing as pain, anxiety, or debase-

ment, then phenotypic changes due to a genetic manipulation will constitute infringements of dignity. On the contrary, if we adopt the Swiss conception of animal dignity, genetic manipulations will count only if they are against the satisfaction of an animal’s interest, and they will constitute a violation of dignity only if the human interests at stake are not greater than the animal’s interest. Under this interpretation, creating “monsters” by genetic engineering—be it true or false, depending on what conception you entertain of a monster—is not a problem if the balance is in favor of human interests.

Ethical Foundation of National Legislation on Animal Experimentation Among the different ethical views expressed above, legislators have generally incorporated into law a pathocentric view, with its emphasis on reformism. Pathocentrism is realized by the law pointing heavily towards the fact that animals should not be in pain, or experience suffering, stress, or in anxiety when in experiments. Legislation has also incorporated hierarchism in animal experimentation regulations, as “lower” species should replace “higher” ones when the result is comparable.

The Use of Ethical Tools to Assess Mouse Experiments What kind of tool is used? How should it be decided whether a mouse experiment is allowed or not, i.e., how to take seriously mice’s interests? The ethical tool applied almost universally in animal experimentation is the harm-benefit assessment. The usual image to describe such a tool is the balance, which will compare the torque on an arm weighing the harm to the animal produced by the experiment to the torque on the other arm weighing the benefit generated by the experiment to humans (and/or animals) in terms of health, environment, and knowledge (three overriding human interests). Instead of the balance, Bateson (1986) proposes a cube with its three dimensions representing quality of research, probability of benefit, and animal suffering, to decide whether a specific animal study should be carried out or not. Quantification tools have been developed as score sheets (Porter, 1992; Boisvert and Porter, 1995; Stafleu et al., 1999); there is also a Web-based self-assessment score sheet, e.g., http://tki.samw.ch/, developed by the Swiss Academy of Sciences.

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The process of weighing harms and benefits is often named ethical review process or ERP.

How to undertake the ethical review process? Although it is easy to understand that the experiment might be acceptable or unacceptable depending onto which side the arms of the balance rotate, or in which part of the cube the project situates, the balance and cube metaphors are misleading. They suppose that the units for measuring harm to the animals are identical to the units measuring benefit for humans and/or animals and quality of research. They suppose that the harm-currency exchange rate is one-to-one with that of the benefit- or quality-of-research currency. This is of course not the case. Moreover, it is difficult to figure out a currency measuring objectively the amount of benefit, the quality of science, or the harm by adding units of benefit, of quality, or of harm. These parameters are partly incommensurable. It is therefore clear that the weighing that one has to do is not a quantitative procedure with mathematical precision. It is rather a question of moral judgment, which will depend on the people doing it. We may compare the process to that of a judge weighing the plea or sentence in the “scales of justice” (Smith and Boyd, 1991).

Who performs the ethical review process?

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Countries have established different ways to do an ERP. The process can take place at the level of the institution like the Institutional Animal Care and Use Committee (IACUC) in the United States of America, or at the level of the state, the region, or the country (for Europe, see review by Smith et al., 2007). It is recommended to do an ERP with a panel of people representing different points of view on animal experimentation. The panel should have a large array of expertise. It should engage in open discussions, and members of the panel should be ready to compromise. Some national regulations call for an ERP done by a single person. This process should always be supervised by a third party. It is also important that the ERP be done locally because it is important to know how animals are used and cared for in the local institutions, as well as the level of training of their researchers and technicians. Researchers submitting a mouse experimentation project should, in some countries are required to, present their own harm-benefit assessment. Every applicant should go through

this assessment to be aware of the ethical issues raised by his/her own work and improve his/her research at the highest ethical standards. The ultimate question that each researcher should ask to him/herself is: “Is it right or wrong to use mice in my project?”

Points that the researcher should consider for a harm-benefit assessment in mouse experiments Points to review for minimizing harm to the animals: Ideally, the best way to minimize harm to animals is to eliminate it by replacing all in vivo work with nonsentient or nonliving alternatives. As we know, there is a long road to travel before this can be achieved. In the meantime, we should systematically apply the 3R concept: Replace, Reduce, and Refine, that Russell and Birch (1959) (see abridged version in Balls, 2009) have developed and published more than 50 years ago. As we allot efforts to find alternatives to animal experimentation (Replacement), we should at the same time minimize the number of animals we use (Reduction), and improve housing conditions, techniques and procedures that we apply to the animals in order to minimize invasiveness and to improve welfare (Refinement). Much national and international legislation have integrated the 3R concept in order to apply a fair ethical harm—benefit assessment to in vivo studies. Replacement of animals: First of all, the researcher has to prove that an animal model is an absolute necessity to achieve the aims. A thorough database search has to be presented to convince both the investigator and the ERP panel that this is the best approach to the problem. Whenever possible, replacementalternatives data should be provided to demonstrate that the animal experiment is the inevitable next step to achieve the intended aims. However, it seems that the research community is not well aware of the available tools to find and review replacement alternatives (Leenaars et al., 2009). An exhaustive literature search in multiple electronic databases is not always simple; advice from librarians may be recommended. Hooijmans et al. (2010a) and Chilov et al. (2007) propose a search filter to collect, respectively, all animal experimentation and alternatives in specific databases such as, for example, the most well-known free database, PubMed (http://www.ncbi.nlm.nih.gov/pubmed). Other fee-for-service databases are also available, like Scopus (http://www.scopus.com) or Web

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of Science (http://apps.isiknowledge.com). Through these database searches, researchers may find a better animal model than the one originally in mind, or alternatives avoiding the use of an animal model. Reduction of number of animals: Russell and Birch (1959) (see abridged version in Balls, 2009) have also insisted on the optimal design of the experiment in order to use the minimum number of animals to achieve statistical significance. An excellent review on design and sample size determination especially devoted to animal experimentation has been written by Festing et al. (2002). However, reduction of number of animals is not a simple matter of statistics. Unnecessary duplication of experiments resulting from poor design, poor training of experimenters, and poor database searches for similar experiments must also be avoided. Therefore, it is important to search genetically modified mice depositories. The KOMP database (http://www.komp.org/), the International Knockout Mouse Consortium (http://www.knockoutmouse.org/), The Jackson Laboratory (http://www.jax.org/ and http://www.informatics.jax.org/), the EMMA consortium (http://www.emmanet.org), and the Japan Mouse/Rat Strain resources database (http://www.shigen.nig.ac.jp/mouse/jmsr/top. jsp), among many others, may be consulted (also see http://www.mmrrc.org/about/ resources.html). Refinement of housing, procedures, and techniques: Refinement can be defined as “. . . methods in animal research which alleviate or minimize the pain, distress, or other adverse effects suffered by the animals involved and/or enhance animal well-being.” (Smaje et al., 1998). Refinement has to be considered throughout the lifespan, including the death, of the laboratory animals. This is probably the one component of the 3Rs that depends heavily on the training of people for the best practices for housing animals, for procedures, and for application of the least invasive techniques. Highly trained people to run the animal facilities and funding for comfortable housing for the animals and for buying and using bioimaging machinery for longitudinal studies and diagnostic equipment that requires minimal biological sample volume are major contributors to Refinement. The listing of all possible means to refine animal studies is long. Here we review briefly the major issues in Refinement of mouse experiments:

1. Husbandry and environment of the animals: • Well equipped animal house, with optimal heating, ventilation and air conditioning (HVAC) equipments. • Unobstructed flows of clean and dirty material, animal, personal and waste. • Optimal enriched housing for laboratory animals that spend their entire lives in cages. • High hygienic standards in animal facilities housing many genetically modified mice coming from laboratories all over the world. • Least invasive animal identification system. 2. Procedures for the care of animals before, during and after procedures: • Transport and adaptation to the experimental environment. • Handling and restraint techniques. • Animal biopsies. • Injection and sampling of biological material. • Anesthesia, analgesia, and surgery. • Post-operative care. • Pain identification, evaluation, and alleviation. • Score sheets to assess humane endpoints and to take actions. Consultation of genetically modified mouse databases which incorporate issues on deleterious phenotypes can greatly help in creating such score sheets (see also http://www.eumodic.org/). • State-of-the-art euthanasia.

The specific costs of creating and using genetically modified mice Worldwide statistics on the number of animals used in experimentation show the preeminence of the mouse model; this species is very well adapted to genetic manipulation, reproduces rapidly, and is easy to house. The genetic model is often far more predictive when testing hypotheses regarding fundamental biological processes or when mimicking human diseases than the classical models used in the past. This signifies that a refinement has been achieved. However, we have to remember that to get this higher-quality (genetically engineered) animal model, many animals have been generated and euthanized during the process. Therefore it is crucial to review databases on genetically modified mice to search for the model of interest. This saves time, money, and animals. However, the GM model approach may in some circumstances create a conflict between the Refinement and Reduction

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component of the 3Rs that the ERP has to solve. It is therefore very important to review the specific costs linked to genetic engineering of mice and to take appropriate actions. Costs are of three kinds: 1. the procedures used to create the GM line; 2. the husbandry to establish and to maintain the GM lines which generate often a large number of animals that have to be discarded; 3. the possible negative impact of the genetic manipulation on the welfare of the individual animals. The review on “refinement and reduction applied to the production of GM mice” published by Robinson et al. (2003) gives excellent advice, from creation to archiving of GM mice, and for implementation of best practices in the lab and the animal facilities (see also Australian Animal Welfare Committee, 2006). Before starting any creation of a GM mouse line, researchers have to make sure that the GM mouse of interest is not already available (see under section “Reduction of number of animals,” listing several mice databases). If not, the genetic engineering process may start with an appropriate design of a transgene. At this stage, procedures to create the GM line have to be reviewed and the following main issues have to be considered: • Selection of mouse strains appropriate as embryo source; high plug rates; implantation; and survival of embryos and pups. • Protocols for superovulation. • Surgery, anesthesia, analgesia, and postoperative followup for best practices of vasectomy and transfer of embryos to foster mothers. • Size/weight/age optimization of female embryo donors and selection of the least aggressive but still sexually aroused stud males. When founders are available, expansion and maintenance of the GM lines raise husbandry issues. The following have to be addressed: • Best practices in colony management. • Implementation of high hygiene standards to prevent health problems. • Pair housing of pregnant mice receiving eggs of the same microinjection experiment instead of single housing. • Enrichment of cages housing single vasectomized and stud mice. • Choice of the least invasive methods to take biopsies for genotyping and identification of animals. When both genotyping and identification of the same individual are required,

consider the dual-purpose approach (ear punch or phalange amputation). • Selection of training of personnel for the most appropriate euthanasia procedures for culling GM mice colonies. • Sending cryopreserved embryos or sperm instead of live animals for inter-laboratory exchange, for purposes of welfare and hygiene. • Archiving GM lines by cryopreservation of embryos, sperm or ovarian tissue. Finally, to minimize welfare problems linked to the genetic modifications, the following have to be considered: • Use of inducible promoters and conditional transgenes to avoid life-long genetically linked welfare problems. • Breeding of homozygotes if no welfare problem is detected. • Phenotyping of new lines from birth onward for early detection of welfare problems. • If welfare problems arise, set clear humane endpoints to avoid excessive harm to the animals.

Maximizing the benefits to humans or animals To justify the harm inflicted upon the laboratory animals, researchers should answer two questions: (1) What is the contribution of my research to the improvement of human (and/or animal) health; and (2) in which timeframe will I achieve the short-, medium-, and long-term aims of the project? Additionally, they should adhere to the duty of following the highest scientific quality standards. Below, “quality of research” is defined. Relevance of research to society and opportunities for reaching specific aims depend heavily on directions taken by society, politics of science, granting agencies, and objectives of large institutions and laboratories. This debate involves every researcher. Quality of science Peer review is the quality-control measure that scientists use to assess quality of their research. It correlates funding to publication of scientific papers in high-impact journals. A published paper of in vivo studies is just the tip of an iceberg of information including the housing and care of animals in facilities, experimental procedures on living beings, study design, allocation of animals to different experimental groups, blinded analysis of biological samples and experimental outcomes, etc. A survey made by Kilkenny et al. (2009) on research using animals shows that reporting

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is often incomplete and not transparent and that design and statistical methods are poor. Lack of accuracy and transparency, and incomplete reporting in animal experimentation, make review and meta-analysis of research fields almost impossible (Hooijmans et al., 2010b; Kilkenny et al., 2010). It means that, often, clinical-trial design cannot be backed up by animal-model results. Such situations further muddle the already controversial field of animal experimentation; moreover, it hampers the benefit side of the harm-benefit assessment of animal experiments. Kilkenny et al. (2009) and Hooijmans et al. (2010b) show also that researchers have to work directly with animal facility teams to grasp the entire complexity of an animal experiment. Therefore, there is an ethical need to perform high-quality work in the lab as well as in the animal house. Progress to achieving high quality science in the field of in vivo studies can be made on the lab side by applying the ARRIVE initiative (for Animal Research: Reporting In Vivo Experiments; Kilkenny et al., 2010) and/or use the checklist for animal experimentation reporting issued by Hooijmans et al. (2010b) from the 3R Center of Radboud University, Nijmegen, The Netherlands. For animal housing, implementation of a quality control procedure is helpful for achieving high-quality housing and care of laboratory animals (Dirnagl, 2010).

CONCLUSIONS Animal experimentation is an important aspect of life science and medical research. It is even mandatory for drug development. Animals, however, are not things we can use as we please; they have a moral standing because they possess interests that can be thwarted. In short, human beings can harm them. From an ethical point of view, harming animals is not impermissible, but ought to be justified. Current legislation of animal protection is pathocentric in essence: human and animal interests have to be weighed to decide whether an animal experiment is morally justified. Experiments with mice must therefore go through a harm-benefit assessment, with the 3Rs being a prerequisite to such assessment. But all animals are not on the same footing; our societies tend to adopt a hierarchical concept of animals, where certain animals count for more than others. For instance, mice have the capacity to suffer and to flourish, but they do not have a mental life as rich as apes or other nonhuman primates. Therefore, their moral standing

is lower, and experiments with this species are easier to justify morally than the same studies on nonhuman primates such as apes. In a reformist perspective of animal experimentation, the 3Rs approach develops its full strength by obligating scientists to think through their experiments thoroughly. In conclusion, ethical considerations with respect to mouse experiments do not only require consideration and improvement of animal welfare, but also improvement of the quality of science.

LITERATURE CITED Australian National Health and Medical Research Council’s Animal Welfare Committee. 2006. Guidelines for the generation, breeding, care and use of genetically modified and cloned animals for scientific purposes. See http://www. nhmrc.gov.au/publications/synopses/ea17syn. htm (August 2, 2010). Balls, M. 2009. The Three Rs and the Humanity Criterion: An abridged version of The Principles of Humane Experimental Technique by WM.S. Russell and R.L. Burch. FRAME, Nottingham, U.K. Bateson, P. 1986. When to experiment on animals. New Scientist 109:30-32. Beauchamp, T. and Walters, L. 1989. Contemporary Issues in Bioethics. Wadsworth Publishing Group, Belmont, Calif. Beyleveld, D. and Brownsword, R. 2002. Human Dignity in Bioethics and Biolaw. Oxford UP, Oxford. Boisvert, D.P and Porter, D.G. 1995. Ethical scoring systems. In Report of the 1995 World Congress on Alternatives, Alternative Methods in Toxicology and Life Sciences Series 11 (A.M. Goldberg and L.F.M. van Zutphen, eds.) pp.637-641. Mary Ann Liebert, Inc. Publishers, New York. Chilov, M., Matsoukas, K., Ispahany, N., Allen, T.Y., and Lustbader, J.W. 2007. Using MeSH to search for alternatives to the use of animals in research. Med. Ref. Serv. Q. 26:55-74. Dirnagl, U. 2010. Quality control and standard operating procedures. Neuromethods 47:239248. Ethics Committee on Non-Human Biotechnology (ECNH) 2001. The Dignity of Animals. Bern, Switzerland. Festing, M.F.W., Overend, Ph., Das, R.G., Borja, M.C., and Berdoy, M. 2002. The Design of Animal Experiments. Reducing the use of animals in research through better experimental design. Laboratory Animal Handbooks N. 14. Laboratory Animals Ltd. The Royal Statistical Society Medicine Press Ltd., London. Hooijmans, C.R., Tillema, A., Leenaars, M. and Ritskes-Hoitinga, M. 2010a. Enhancing search efficiency by means of a search filter for finding all studies on animal experimentation in PubMed. Lab. Anim. 44:170-175.

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Hooijmans, C.R., Leenaars, M., and RitskesHoitinga, M. 2010b. A gold standard publication checklist to improve the quality of animal studies, to fully integrate the three Rs, and to make systematic reviews more feasible. Altern. Lab. Anim. 38:167-182. Kant, I. 1978. Anthropology from a Pragmatic Point of View (1798): Translated by Victor Lyle Dowdell, revised and edited by Rudnick. Southern Illinois University, Carbondale and Edwardsville, Ill. Kant, I., 1996. Metaphysics of Morals (1797). Cambridge University Press, Cambridge, U.K. Kilkenny, C., Parsons, N., Kadyszewski, E., Festing, M.F., Cuthill, I.C., Fry, D., Hutton, J., and Altman, D.G. 2009. Survey of the quality of experimental design, statistical analysis and reporting of research using animals. PloS ONE 4:e7824. Kilkenny, C., Browne, W.J., Cuthill, I.C., Emerson, M., and Altman, D.G. 2010. Improving bioscience research reporting: The ARRIVE guidelines for reporting animal research. PLoS Biol. 8:e1000412. Krepper, P. 2010. Tierw¨urde im Recht: am Beispiel von Tierversuchen. Aktuelle Juristische Praxis 4:303-313. Leenaars, M., Savenije, B., Nagtegaal, A., Van Der Vaart, L., and Ritskes-Hoitinga, M. 2009. Assessing the search for and implementation of the three Rs: A survey among scientists. Altern. Lab. Anim. 37:297-303. Mason, G. and Littin, K.E. 2003. The humaneness of rodent pest control. Anim. Welfare 12:1-37. Meerburg, B.G., Brom, F.W.A., and Kijlstra, A. 2008. The ethics of rodent control. Pest Manag. Sci. 64:1205-1211. Nuffield Council on Bioethics. 1996. Animalto-Human Transplants, the Ethics or Xenotransplantation. http://www.nuffieldbioethics. org/xenotransplantation. Porter, D.G 1992. Ethical scores for animal experiments. Nature 356:101-102. Regan, T. 1984. The Case for Animal Rights. Routledge, London. Robinson, V., Morton, D.B., Anderson, O., Carver, F.A., Francis, R.J., Hubrecht, R., Jenkins, E., Mathers, K.E., Raymond, R., Rosewell, I., Wallace, J., and Wells, D.J. 2003. Refinement and reduction in production of genetically modified mice. Sixth report of the BVAAWF/ FRAME/RSPCA/UFAW Joint Working Group. Lab. Anim. 37:1-51. Russell, W.M.S. and Birch, R.L. 1959. The principles of humane experimental techniques. http://altweb.jhsph.edu/pubs/books/humane exp/het-toc; abridged version reprinted in Balls (2009). Ethical Considerations in Mouse Experiments

Singer, P. 1979. Practical Ethics. Oxford University Press, Oxford. Smaje, L., Smith, J.A., Combes, R.D., Ewbank, R., Gregory, J.A., Jennings, M., Moore, G.J.,

and Morton, D.B. 1998. Advancing refinement of laboratory animal use. Lab. Anim. 32:137142. Smith, J.A. and Boyd, K.M. (eds.) 1991. Lives in Balance: The Ethics of Using Animals in Biomedical Research. Oxford University Press, Oxford. Smith, J.A., van den Broek, F.A.R., Canto Martorell, J., Hackbarth, H., Ruksenas, O., and Zeller, W. 2007. Principles and practice in ethical review of animal experiments across Europe: Summary of the report of the FELASA Working Group on Ethical Evaluation of Animal Experiments. Lab. Anim. 41:143-160. Stafleu, F.R., Tramper, R., Vorstenbosch, J., and Joles, J.A. 1999. The ethical acceptability of animal experiments: A proposal for a system to support decision-making. Lab. Anim. 33:95303. Swiss Academies of Arts and Sciences. 2010. The dignity of animals and the evaluation of interests in the Swiss Animal Protection Act. http://www.swiss-academies.ch Warren, M.A. 1997. Moral Status. Oxford University Press, Oxford, United Kingdom.

APPENDIX: GLOSSARY Anthropocentrism: A moral theory is anthropocentrist if it puts human being at the top of the moral realm. Usually it gives moral status only to human beings (but see hierarchism). Autonomy: A being is autonomous if he can choose between various possibilities and act accordingly, i.e., freely. Traditionally, autonomy requires the capacity to ponder about one’s own desires (see Reason), but certain authors attribute autonomy to animals, as they can act accordingly to their desires or preferences. Biocentrism: A moral theory is biocentrist if it believes that the morally relevant intrinsic property is life. For biocentrism, all living beings possess moral status. Dignity: Dignity is synonymous with intrinsic value, and denotes a high intrinsic value. It has therefore traditionally been kept for human beings. Harm-benefit assessment: This is a weighing between the likely adverse effects on the animals and the benefits to human beings or other species likely to accrue as the result of the research. Hierarchism: A moral theory is hierarchist if it believes that there exist several moral statuses ranked on a scale of value. Usually it puts human beings at the top of the scale, then animals followed by plants, etc.

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Indirect duty: A person has an indirect duty towards a being (e.g., an animal) if this being is only an opportunity for the person to fulfill a direct duty towards another being (e.g., a human person). Intrinsic property: A being possesses two kinds of properties, intrinsic and extrinsic. A property is intrinsic if it belongs to the being independently of his environment; in any other case, the property is extrinsic. “To have four legs” is an intrinsic property for a rat, whereas “To be bigger than a mouse” is an extrinsic one. Intrinsic value: A being can have two kinds of value, intrinsic and extrinsic. His value is intrinsic if it depends on his morally relevant intrinsic properties; in any other case, his value is extrinsic. “To be sentient” gives an intrinsic value to a rat, whereas “to be loved” gives an extrinsic one. Moral agent: A being is a moral agent if he has moral duties towards other beings; he is a moral patient if he has only moral rights and no moral duties. Adult human beings are moral agents, children and, for certain authors, animals, are moral patients. Moral status: This denotes the place that a being occupies in the moral realm. The place of a human being is different from the place of a rat. Their moral status is therefore not

the same. To determine a being’s moral status, we must take his morally relevant intrinsic properties and his intrinsic value into account. Pathocentrism: A moral theory is pathocentrist if it believes that the morally relevant intrinsic property is sentience, i.e., the capacity for suffering and for happiness. For pathocentrism, only sentient beings, i.e., human beings and animals, possess moral status. 3Rs: Described first by Russell and Birch (1959), three principles that guide the use of animals in experimentation: (1) replacement (use of alternatives to animals whenever it is possible to reach the same scientific goals); reduction (use of minimal number of animals to obtain scientifically significant results); and refinement (use of any method and procedure to decrease or eliminate pain, suffering, distress, fear, or anxiety, and to increase the welfare of animals that cannot be replaced by other methods). Reason: This is a multifarious capacity that comprises in particular autonomy, selfconsciousness, conscience, and self-mastery. Traditionally, this capacity is kept for human beings. Sentience: This is the capacity for suffering and for happiness. For pathocentrism, this capacity determines moral status.

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Lentiviral Vector Mediated Transgenesis Isabelle Barde,1 Sonia Verp,1 Sandra Offner,1 and Didier Trono1 1

School of Life Sciences and “Frontiers in Genetics” National Program, Ecole Polytechnique F´ed´erale de Lausanne (EPFL), Lausanne, Switzerland

ABSTRACT The genetic manipulation of rodents through the generation of fully transgenic animals or via the modification of selective cells or organs is a procedure of paramount importance for biomedical research, either to address fundamental questions or to develop preclinical models of human diseases. Lentiviral vectors occupy the front stage in this scene, as they can mediate the integration and stable expression of transgenes both in vitro and in vivo. Widely used to modify a variety of cells, including re-implantable somatic and embryonic stem cells, lentiviral vectors can also be directly administered in vivo, for instance in the brain. However, perhaps their most spectacular research application is in the generation of transgenic animals. Compared with the three-decade-old DNA pronuclear injection technique, lentivector-mediated transgenesis is simple, cheap, and highly efficient. Furthermore, it can take full advantage of the great diversity of lentiviral vectors developed for other applications, and thus allows for ubiquitous or tissue-specific or constitutive or externally controllable transgene expression, as well as RNAi-mediated gene C 2011 by John Wiley & Sons, Inc. knockdown. Curr. Protoc. Mouse Biol. 1:169-184  Keywords: lentiviral vector r transgenesis r transgenic animals It was with retroviruses that the generation of transgenic mice was first attempted some thirty years ago. However, while infection of 2- to 4-cell-stage embryos succeeded in yielding genotypically positive animals capable of transmitting the transgene to their offspring, the foreign genetic material was systematically silenced due to proviral DNA methylation (Jahner and Jaenisch, 1985). Shortly thereafter, it was discovered that the direct injection of naked DNA into the pronucleus of a fertilized oocyte resulted in its integration into the host genome, and that most transgenes introduced by this technique were subsequently expressed and transmitted to progeny (Gordon et al., 1980). It thus became the standard method to create transgenic animals. Nevertheless, this approach suffers from obvious shortcomings: it is technically demanding and moderately efficient, with only a fraction of transgenic animals appropriately expressing the foreign sequence, and it is largely limited to mice because only mice produce oocytes whose transparency is suitable for injection of a clearly visible pronucleus.

BASIC PROTOCOL

In 2002, the old idea of retrovirally generated transgenic animal resurfaced, owing to the demonstration that lentiviral vectors could mediate efficient in vivo gene delivery of transgenes that largely escaped epigenetic silencing (Lois et al., 2002). Lentivector-mediated transgenesis is relatively easy to perform and leads to high percentages of provirus-positive animals. Moreover, a wide variety of lentiviral vectors have been developed that can all be used in transgenic animals, thus allowing for a broad range of genetic manipulations including externally controllable expression and knockdown, the latter offering an economically advantageous alternative to stable knockout (Tiscornia et al., 2003; Szulc et al., 2006). In a typical procedure, the lentiviral vector is injected beneath the zona pellucida of a fertilized oocyte, and it then penetrates by fusion between the viral and plasma membranes.

Current Protocols in Mouse Biology 1: 169-184, March 2011 Published online March 2011 in Wiley Online Library (wileyonlinelibrary.com). DOI: 10.1002/9780470942390.mo100169 C 2011 John Wiley & Sons, Inc. Copyright 

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Its RNA genome then undergoes reverse transcription, yielding a double-stranded DNA copy that integrates into the host cell chromosomes. The kinetics of lentiviral vector integration into the target genome defines the degree of genotypic mosaicism found in the resulting animal. The genotypic characterization of the transgenic mice obtained from oocytes injected with lentiviral vectors demonstrates that integration occurs rapidly following this procedure, at the 2- to 4-cell stage, so that the degree of mosaicism is low in the resulting animals (Sauvain et al., 2008). Moreover, the phenotypic consequences of this mosaicism can usually be minimized through the use of vector doses high enough to induce multiple proviral copies per embryo, ensuring that all cells harbor at least one integrant. As already noted in human cells, lentiviral vector integration in transgenic mice favors certain genes, particularly the ones active at the time of infection, occurs indifferently with respect to transcriptional orientation, without bias for introns or exons, and tends to concentrate in the middle of the transcribed region (Sauvain et al., 2008). The advantage of this protocol is that it is simple and highly efficient, resulting in an average (calculation based on approximately 10,000 injections performed) of 67% of the resulting animals carrying the provirus.

Materials Mouse hybrid strain B6D2F1 derived from C57BL/6JxDBA2J (females and males purchased at 5 and 8 weeks of age, respectively from Charles River Laboratories) Pregnant mare serum (PMS; see recipe) Human chorionic gonadotropin (HCG; see recipe) Egg medium (see recipe) Embryo-tested mineral oil, sterile filtered, (Sigma, cat. no. M5310) 70% ethanol Hyaluronidase (H-3506 Sigma) NMRI female 7 weeks old minimum (Charles River Laboratories) Vasectomized males NMRI, 2 to 10 months old (Charles River Laboratories) Lentiviral vector (see Barde et al., 2010) Ketarum (see recipe) Artificial tears: Viscotears (Novartis Pharma Schweiz AG, Bern, Switzerland) Phosphate-buffered saline (PBS) Heparin-Na 25000 IE/5 ml (Braun, 3511014; http://www.bbraunusa.com/) Histopaque-1083 (Sigma, cat. no. H8889) PBS containing 1% (v/v) fetal bovine serum (FBS) DNAeasy Genomic DNA Extraction Kit (Qiagen, cat. no. 69509) pTitin (available from Addgene, http://www.addgene.org), a pRRL vector in which the target sequence of the Titin primers used for normalization has been cloned (this plasmid allows one to perform a standard curve) Kit for preparing qPCR master mix (TaqMan universal PCR master mix, no AmpErase UNG; Applied Biosystems, cat. no. 4324020, including 2× reaction buffer) Primers and probe for Gag, WPRE, and Titin detection (see Table 1)

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U-100 insulin syringe, 29-G, 0.33 mm × 12.7 mm, Microfine (Becton Dickinson) Petri dishes (3 cm, sterile) Surgical draping Surgical instruments Understage illumination stereomicroscope (Leica, MZ7.5) Mouth pipet aspirator tube assemblies for calibrated microcapillary pipets (Sigma, cat. no. A5177-5EA) Capillaries for mouth pipet (Drummond microdispenser; 2-000-050; Milian)

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Table 1 Primers and Probes for the Genotyping of Transgenic Animals Obtained by Lentiviral Vector–Mediated Transgenesisa

Sequence detected Primer/probe name Primer/probe sequence Gag

WPRE

Titin

Gag Forward

GGAGCTAGAACGATTCGCAGTTA

Gag Reverse

GGTGTAGCTGTCCCAGTATTTGTC

Gag probe

ACAGCCTTCTGATGTTTCTAACAGGCCAGG

WPRE forward

GGCACTGACAATTCCGTGGT

WPRE reverse

AGGGACGTAGCAGAAGGACG

WPRE probe

ACGTCCTTTCCATGGCTGCTCGC

Titin forward

AAAACGAGCAGTGACGTGAGC

Titin reverse

TTCAGTCATGCTGCTAGCGC

Titin probe

TGCACGGAAGCGTCTCGTCTCAGTC

Probe fluorophores

FAM-BHQ

FAM-BHQ

FAM-BHQ

a Gag oligos are used for amplification of HIV-1 derived vector sequences and are specific for the 5 end of the gag gene. This sequence is present

in all HIV-1-derived vectors, as it is part of the extended packaging signal. WPRE oligos amplify the WPRE sequence present in almost all later-generation LV vectors (see Commentary). Titin oligos are used to normalize for the amount of genomic DNA and are specific for the mouse titin gene. Stocks of probes and primers usually come lyophilized and are diluted to 10 μM in water.

Capillary for injection micropipet: borosilicate standard wall with filament, 1.2 mm O.D., 0.69 mm I.D., 100 mm length, (Harvard Apparatus, cat. no. 300044) Puller for preparing injection micropipet (INJECT+MATIC, http://www.injectmatic.com/) Sterile holding pipet vacutips (Vaudaux-Eppendorf, 5175 108.000) Sequencing tip microloader (Vaudaux-Eppendorf, 5242 956) Inverted microscope (Leica AS TP) with TransferMan NK2 micromanipulators (Eppendorf) and microinjector (INJECT+MATIC, http://www.injectmatic.com/) Hypodermic needles, microlance, 30-G, 12 -in. Heating pad 5 × 12.5 cm (40-90-2-07 FHC) Suture clips MicroAmp 96-well optical reaction plate (Applied Biosystems) Optical adhesive Film (Applied Biosystems) Centrifuge with microtiter plate carrier Real-time PCR machine (e.g., 7900HT Sequence Detector, Applied Biosystems) Computer running SDS7900HT software (Applied Biosystems) and Microsoft Excel Additional reagents and equipment for sacrifice of mice (Donovan and Brown, 2006), flow cytometry (Robinson et al., 2010), and quantitative real-time PCR (qPCR; Fraga et al., 2008) Induce superovulation 1. At 5 pm on day 1, inject five B6D2F1 females (6 to10 weeks old) intraperitoneally (i.p.) with pregnant mare serum (PMS) at 10 IU/mouse, using an insulin syringe. Females are superovulated 3 days before collection of the embryos.

2. After 46 to 48 hr, inject the females i.p. with human chorionic gonadotropin (HCG), 10 IU/mouse, using an insulin syringe . PMS is used to mimic the oocyte maturation effect of the endogenous follicle-stimulating hormone (FSH). HCG is administered to mimic the ovulation induction effect of luteinizing hormone (LH).

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3. Immediately following these injections (at approximately 4 to 5 pm on day 3), individually mate each female with a male counterpart, setting up five mating pairs. B6D2F1 stud males are singly housed and are used as breeders in the prime of their reproductive life, that is, between 2 and 8 months with 1 week of rest after each mating. In addition, a record of their performance should be maintained, as males that have been mated successfully in the past (as indicated by a vaginal plug) are likely to do the same in the future. Other strains can be used for transgenesis; however, the age, time, and hormone doses need to be set up to obtain enough embryos. The conditions described here can be used for the C57BL/c6 strain.

Harvest embryos 4. First thing in the morning (day 4), prepare all of the Petri dishes described below by equilibrating them in a CO2 incubator at 37◦ C for a minimum of 30 min (Fig. 1): Two 3-cm dishes each containing 1 ml of egg medium (Fig. 1A). Two 3-cm dishes each containing six droplets of 25 μl egg medium covered by embryo-tested mineral oil (Fig. 1B). One lid of a 3-cm dish, to be used as injection chamber, containing two droplets of 20 μl egg medium covered by embryo-tested mineral oil (Fig. 1C). 5. Inspect the females visually in the morning—0.5 days post coitum (dpc)—for a copulation plug. To do so, gently hold each mouse by the tail. Visually inspect the female genitals for the presence of a crusty whitish plug called the vaginal plug that is the proof of a successful mating. This plug is the semen of the male that coagulates inside the vagina. The check should be made in the morning as the plug dissolves with time, and mating usually occurs in the middle of the night.

6. Sacrifice the B6D2F1 females showing a copulation plug, by cervical dislocation (Donovan and Brown, 2006) at 0.5 dpc.

Lentiviral Vector Mediated Transgenesis

Figure 1 Specific materials required for the manipulation of embryos. A 3-cm Petri dish with 1 ml egg medium (A); a 3-cm dish with six droplets of 25 μl of egg medium covered by embryo-tested mineral oil (B); injection chamber: one lid of a 3 cm dish with two droplets of 20 μl of egg medium covered by embryo-tested mineral oil (C); and a mouth pipet (D) containing a fine capillary at its tip (E).

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Figure 2 Harvest of embryos. The dissected part of the ovary and oviduct are placed in the 3-cm dish with 1 ml prewarmed egg medium (A). The ampulla (Am) is the bulged part of the oviduct and is normally easily visible if the superovulation was successful. The ampulla is opened by carefully tearing the wall using fine forceps (B), and the embryos (Em) then pop out (C). The embryos are still surrounded by numerous small cells (D) that the hyaluronidase disaggregates (E).

7. Place the mice on a clean draping with their ventral sides up. Wipe the abdominal fur with 70% ethanol. Using a clean pair of dissecting scissors, make an incision along the ventral midline from the pubis to the rib cage. Resect the skin and mesentery and move the viscera aside to expose the ovaries, oviducts, and uterine horns. The ovaries are easily recognized at the tip of the uterus as round whitish structures covered by the peri-ovarian fat pad. On average, 30 to 40 embryos can be collected from each mouse.

8. With the help of forceps, gently hold the uterine horn just above the oviduct and make an incision with fine scissors below the forceps. Make a second cut in the middle of the ovary, making sure not to touch the ampulla. Immediately put both the ovary and the oviduct in the 3-cm dish containing 1 ml prewarmed egg medium (Fig. 2). 9. Under an under-stage illumination stereomicroscope, open the ampullas by carefully tearing their wall using fine forceps; the embryos will then pop out (Fig. 2). The ampulla is the bulged part of the oviduct (Fig. 2A) and is normally easily visible if the superovulation was successful. At this point, the embryos are still surrounded by numerous small cells forming the cumulus oophorus, which the hyaluronidase will disaggregate.

10. Add a few milligrams of hyaluronidase (using the tip of the forceps) to the medium and incubate for 3 min at 37◦ C in order to remove the cumulus oophorus. 11. Rapidly transfer the embryos (using the mouth pipet in Fig. 1D) to the second 3-cm dish containing 1 ml egg medium. Try to take as little medium as possible while pipetting the embryos. To handle the embryos, we use a mouth pipet containing a fine capillary at its tip (Fig. 1D). This fine capillary is home-made by stretching it with a flame.

12. Wash the embryos in the first three droplets in the Petri dish by pipetting them up and down four to five times, and then transfer them to the last three droplets by taking the least amount of medium possible (Fig. 1B), with no more than 50 embryos per droplets. At this stage, the embryos are counted and kept in the incubator until injection with the lentiviral vector.

Inject lentiviral vector All the injection steps should be performed in a state-of-the art P2 laboratory (WHO, 2004).

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13. Prepare the injection micropipet by pulling borosilicate glass capillaries with the puller apparatus. Also prepare the holding pipet to be used in step 18. Refer to http://www.injectmatic.com/produits.php?lang=fr&idProduit=2 for more information. 14. Thaw a 15-μl aliquot of high-titer lentiviral vector on ice (Barde et al., 2010), briefly vortex, and microcentrifuge 5 min at maximum speed, room temperature, to remove potential aggregates. All lentiviral vectors must have been correctly produced and titered (see Barde et al., 2010). A cut-off limit of 5 × 108 transducing units (TU)/ml is the rule, and the quality of the preparation must be checked with a p24 ELISA: the ratio of infectious particles to physical particles must be 1/100 to 1/1000 at most. The vectors used in this protocol are HIV-1 derived lentiviral vector and were produced with a second-generation packaging system (for vector details, see Fig. 6).

15. Load the injection micropipet with 3 μl of the lentiviral vector, using a sequencing tip that is thin enough to be inserted into the capillary. Take care not to introduce air bubbles, as the injection will then be more difficult. The presence of air bubbles can be checked under a low-magnification loupe.

16. Position the holding pipet and the injection micropipet on their respective micromanipulators. 17. Add 50 embryos per droplet into the injection chamber and place it on the plate of the inverted microscope for injection. Using the low-magnification lens, focus on the embryos at the center of the field to visualize them during the correct positioning of the pipets. Embryos have been fertilized and thus present two polar bodies, two pronuclei, and a nice round shape.

18. Place the injection micropipet on pressure mode. While keeping the pressure on, break the tip of the injection micropipet by flicking it on the holding pipet. 19. Using gentle suction through the holding pipet, hold and maneuver an embryo. Then introduce the injection micropipet through the zona pellucida into the perivitelline space, and deliver 10 to 100 pl of the vector stock (Fig. 3). Be careful not to breach the plasma membrane with the injection micropipet, as this might kill the embryo. On average 100 embryos are injected per lentiviral vector tested.

Lentiviral Vector Mediated Transgenesis

Figure 3 Perivitelline injection of the lentiviral vector stock. An embryo is held on the left with the holding pipet and on the right the injection micropipet containing the lentiviral vector stock is introduced through the zona pellucida (A). The injection micropipet is gently pushed into the perivitelline space (B) and delivers 10 to 100 pl of the vector stock leading to visible swelling of the area (C).

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20. At the end of the procedure, wash the injected eggs successively in three of the egg medium droplets placed in the other 3-cm Petri dish, and distribute them into the other three droplets, with no more than 50 embryos per droplet. Leave overnight at 37◦ C in a CO2 incubator. As the use of mouth pipetting to transfer the transduced embryos can be hazardous, it is recommended to interpose a 0.22-μm filter in the middle of the plastic tube of the mouth pipet (Fig. 1D).

21. Mate 30 foster NMRI females with vasectomized males (two females per male). Usually, 30 NMRI females are necessary to obtain between three and five successful matings. To increase this efficiency, the mating can be started earlier; for example, 2 days before the injection, mate ten females in order to have some plugs ready on the day of injection. This makes it possible to transfer some embryos directly and save them in case there is no plug the day after the vector injection. In this case only 20 females are mated for injection the day after.

Transfer embryos 22. After the overnight incubation (step 20), check under a stereomicroscope to confirm that most of the embryos (usually around 80%) have progressed to the 2-cell stage. Only these are deemed to be viable and will be transferred into the NMRI mated females.

23. In the morning, anesthetize a female with an observed vaginal plug by i.p. injection of 100 μl Ketarum per 10 g of body weight. Protect the eyes of the mouse with a drop of artificial tears in each eye. 24. Disinfect the two flanks of the mouse with 70% ethanol. 25. Place the animal on one side, perform a 1-cm incision in the skin over the lateral lumbar region with sharp scissors to visualize, through the peritoneum, the underlying ovarian fat pad and attached ovary. 26. Make a 0.5-cm incision in the body wall overlying the ovarian fat pad, and, using blunt forceps, gently remove the fat pad along with the attached ovary and uterine horn from the peritoneal cavity. 27. Under a stereomicroscope, visualize the ampulla: it must appear as a dilatation of the oviduct. Collect 8 to 10 transduced embryos inside the capillary of a mouth pipet. IMPORTANT NOTE: Load the embryos into the capillary with a minimal amount of medium to increase transfer efficiency.

28. Determine the orientation of the ampulla, grasp gently, with fine forceps, the nondilated proximal part, and make a small hole with a 30-G, 12 -in. hypodermic needle. Insert the embryo-containing capillary into this hole and inject the embryos gently towards the ampulla. Ideally, a small air bubble should be visible (Fig. 4).

29. Put the uterine horn, ovary, and fat pad back into the body cavity. Seal the skin incision with suture clips. Repeat the procedure on the other uterine horn. 30. Following the surgery, place the recipient females on a heating pad (37◦ C). Observe the mice for several minutes to ensure that their breathing appears normal for their anesthetized condition (they will usually stay down for 15 to 20 min following the surgical procedure). After recovery from the uterine transfer surgery, house the recipient females two per cage.

Genotype the transgenic animals 31. At a point 4 weeks after the birth of the transgenic offspring, collect 100 μl of blood from the tail vein of each of the transgenic animals. For this, prewarm the animal

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A

M F

Figure 4 Transfer of embryos into the ampulla. Under a stereomicroscope, the ampulla (A) is held with fine forceps (F), and the embryo-containing capillary of the mouth pipet (M) is introduced into the hole previously made with a hypodermic needle. The embryos are injected gently towards the ampulla.

under a heating lamp for 5 min until the animal is warm; use a scalpel to do a small incision on a tail vein and collect the blood in a 5-ml tube containing 1 ml of PBS with one drop of heparin (5000 IE/ml). 32. Isolate leukocytes on a Histopaque gradient according to the manufacturer’s instructions. Basically, vortex the tube and add 750 μl of Histopaque solution slowly at the bottom. Centrifuge 8 min at 400 × g, room temperature, with the brake off. Harvest the ring in the middle of the tube and wash it with 1 ml PBS containing 1% FBS, by centrifuging 5 min at 300 × g, 4◦ C, and removing the supernatant.

Determine injection efficiency 33. Analyze the percentage of lentivirus-expressing cells directly by flow cytometry (Robinson et al., 2010) for those lentiviral vectors that allow expression in leukocytes of a fluorescent protein or any product that can be detected with an antibody. 34. Determine the number of proviruses integrated by analyzing the cells using quantitative real-time PCR (qPCR; Fraga et al., 2008). For this, extract cell DNA from each individual sample using a DNAeasy kit according to the manufacturer’s recommendations. For the DNA elution step, use 100 μl of AE buffer from the kit instead of 200 μl. Store DNA at –20◦ C until use. 35. Prepare a plasmid standard curve for the quantitative real time PCR: a. First, adjust accurately the DNA concentration of the pTitin plasmid to 1 mg/ml (the Titin concentration is 1.2 × 1011 molecules/μl). Calculation of the molecule number is as follows: Lentiviral Vector Mediated Transgenesis

7584 (no. of bp of pTitin) × 660 (average molecular weight of a base pair) = 5 × 106 g = ∼1 mol; if plasmid concentration is 1 mg/ml then 1 × 10−6 g = ∼2 × 10−13 mol. 2 × 10−13 mol × 6.02 × 1023 (Avogadro number) = 1.2 × 1011 molecules/μl.

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b. The first point of the standard curve is 1 × 107 molecules in 8 μl = 1.25 × 106 molecules/μl. c. Generate the standard curve by making serial 10-fold dilutions until there are 10 molecules in 8 μl. d. Dilute the plasmid as shown in Figure 5A. 36. Perform qPCR reaction (also see Fraga et al., 2008): a. Prepare a mix (containing everything but the sample DNA) for the number of wells needed for the qPCR analysis, including all samples and standards in duplicates according to the following recipe (9 μl per well): 8.5 μl 2× reaction buffer from Applied Biosystem qPCR MasterMix 0.17 μl forward primer (10 μM) 0.17 μl reverse primer (10 μM) 0.17 μl probe (10 μM). The Taq polymerase is part of the 2× reaction buffer. The primers for Gag, WPRE, and Titin are amplified in separate reactions.

b. Place 9 μl of this mix into each well of a 96-well optical reaction plate. c. Add 8 μl of sample DNA (from steps 34 to 35) to each of the appropriate wells (see Fig. 5B). DNA concentration of sample must be between 50 to 100 ng in 8 μl. d. Close plate with optical film. e. Centrifuge the plate 1 min at 200 × g, room temperature, in a centrifuge with a microtiter plate carrier, to bring all liquid to the bottom of the wells. f. Place the 96-well plate in the real-time PCR machine and run the appropriate program depending on the fluorochromes (FAM) and quenchers (BHQ = non fluorescent) used in the TaqMan probes. Use the following temperature cycling protocol: 1 cycle: 50 cycles:

10 min 95◦ C 15 sec 95◦ C 1 min 60◦ C

(initial denaturation) (denaturation) (annealing).

Be aware that the precise settings of a qPCR protocol depend on the real-time PCR machine and mix used. This aspect is beyond the scope of this protocol (see also Fraga et al., 2008). If not familiar with qPCR techniques, one should seek advice from a local qPCR expert or from the technical support department of the real-time PCR machine supplier.

Analyze the results 37. Analyze the amplification reactions using the SDS7900HT software. a. In the setup of the SDS document, assign, for each point of the standard curve, the corresponding value and the standard status. b. The SDS program will automatically calculate the standard curve and the quantity of each unknown sample for each gene of interest. An example of amplification profiles of HIV sequences in human DNA is given in Figure 5C (as displayed by SDS).

38. Set the threshold values (Ct) where the amplification curve is the steepest, both for the gene of interest (Gag or WPRE) and for the internal control (Titin). These Ct values are the number of cycles required for the amplification curve to cross the absorbance threshold values. Also see Fraga et al. (2008).

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Figure 5

(legend appears on following next page)

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39. Export the results as a Microsoft Excel spreadsheet and calculate the HIV sequences copy number per cell for each sample using the following formula: Vector copy number = (quantity mean of Gag or WPRE sequence)/(quantity mean of Titin/2). The 2-fold factor reflects the fact that the Titin sequence is present in two copies per genome (two alleles). Using standard DNA-extraction procedures in a laboratory context where HIV sequences are often handled, one can expect a level of background contamination with HIV sequences corresponding to cells containing 1 copy per 1000 genomes. In this case, consider higher copy numbers for calibration. Using careful DNA extraction procedures and standardization as described above, one can expect reproducibility within a 2-fold range. Investigators should consult a local qPCR expert if a more stringent quantitative PCR procedure is needed.

REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps.

Egg medium 500 ml 1× Minimum Essential Medium (Invitrogen, cat. no. 41090028) 5 ml L-glutamine/penicillin/streptomycin (Brunschwig, P11-013; http://www. brunschwig-ch.com/) 1.3 g sodium lactate (Fluka, cat. no. 71718) 17.7 mg sodium pyruvate (Sigma, cat. no. S8636) 2 g BSA (Sigma, cat. no. A3311) All reagents are mixed under sterile conditions Divide into 25-ml aliquots Store up to 1 year at −20◦ C Filter (0.22 μm) before use Human chorionic gonadotropin (HCG) Purchase 1500 UI Chorulon (Veterinaria AG, http://www.veterinaria.ch/). Dilute the powder in 15 ml of 1× PBS to obtain a final concentration of 10 UI/100 μl. Divide into 0.6-ml aliquots in microcentrifuge tubes and store up to 1 year at –80◦ C.

Ketarum Mix 1.2 ml ketamine (Ketasol, Dr. Graeub AG, http://www.graeub.com/) with 0.8 ml xylazine (Xylasol, Dr. Graeub AG, http://www.graeub.com/) and make up to 10 ml with 1× PBS. The solution can be stored up to 1 month at 4◦ C.

Pregnant mare serum (PMS) 1000 UI Folligon (Veterinaria AG, http://www.veterinaria.ch/). Dilute this powder in 10 ml of 1× PBS to obtain a final concentration of 10 UI/100 μl. Divide into 0.6-ml aliquots in microcentrifuge tubes and store up to 1 year at –80◦ C. Figure 5 (figure appears on previous page) Setup and analysis for the qPCR genotyping of the transgenic animals. (A) Dilution of the pTitin plasmid standard for the qPCR titration. (B) Example of template of the 96-well plate for the qPCR assay. (C) Representative qPCR analysis. Genomic DNA from transgenic mouse blood was subjected to qPCR amplification and monitoring using a Perkin-Elmer 7900HT (Applied Biosystems) with sets of primers and probes specific for HIV Gag or Titin sequences. Amplification plots were displayed and cycle threshold values (Ct) were set as described in the text. SDS software allows calculation of the standard curve (square point) and determination of Gag and Titin quantity for each animal (cross point). Values of Gag and Titin quantities exported into an Excel worksheet. Current Protocols in Mouse Biology

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COMMENTARY Background Information

Lentiviral Vector Mediated Transgenesis

The pronuclear injection of naked DNA remains the method of choice to establish stable lineages of transgenic mice. However, it is technically demanding, restricted to the mouse, and relatively inefficient, as many lines need to be screened in order to identify one that stably transmits and correctly expresses the transgene. These limitations have prompted the search for new techniques simpler to perform, more efficient, and applicable to multiple species. The development of lentiviral vectors and the demonstration of their ability to mediate the integration and stable expression of transgenes in vivo (Naldini et al., 1996) suggested that they might represent a useful tool for generating transgenic animals. The proofof-principle of this concept was provided in 2002, with the lentivector-mediated generation of transgenic mice and rats (Lois et al., 2002). Remarkably, lentiviral vectors turned out to be far less susceptible to epigenetic silencing than their murine leukemia virus (MLV) counterparts, which had been dismissed three decades earlier for their inability to sustain transgene expression due to proviral DNA methylation (Jahner et al., 1982). It remains to be formally determined whether newer-generation MLV vectors, devoid of silencing cis-acting sequences (Wolf and Goff, 2007), might suffer from the same drawback. In the original lentivector-mediated transgenesis experiment, a vector carrying a GFP reporter gene driven by a ubiquitous promoter was injected between the plasma membrane of fertilized oocytes and the glycoproteinic outer membrane, called the zona pellucida. The space between these two membranes is called the perivitelline space, and can be clearly visualized next to the polar body where the zona pellucida loops out. Once in this perivitelline space, the vector anchors to the plasma membrane, penetrates into the cytoplasm by receptor-mediated endocytosis, and finally integrates into the chromosomal DNA. The results obtained were encouraging, as more than 30% of injected embryos produced transgenic animals, compared to 3% using pronuclear injection. Moreover, these transgenic animals expressed the transgene in more than 80% of the cases. Interestingly, the approach was successfully applied to the rat, a species known to be difficult to manipulate genetically. This experiment produced transgenic rats at a frequency of 59% using the same lentiviral construct, with a rate of transgene expression as high as 40%. These

figures should be compared with the 17.4% efficiency of rat transgenesis using pronuclear injection. Finally, expression of GFP could be restricted to a given tissue using tissue-specific promoters such as those of myogenin, active in skeletal muscles, or lck, active in T cells only (Lois et al., 2002). Lentiviral vector– mediated transgenesis was not only superior to pronuclear injection for generating transgenic animals from different species than laboratory mice, but drastically reduced the technical skills required for such operation. In an attempt to further simplify the procedure, lentiviral vectors were coincubated for a few hours with embryos, the zona pellucida of which had been previously removed by acidic treatment. Embryos were then extensively washed and transferred to surrogate mothers. The efficiency dropped significantly with this procedure but the percentage of transgenic animals was still comparable to that obtained by pronuclear injection (Okada et al., 2007). In our experience, even though the transduction of “depellucidated” embryos is an alternative to perivitelline injection, this method, while theoretically advantageous because it does not require any heavy equipment, carries two major drawbacks. First, the embryos devoid of zona pellucida cannot be transferred before the morula stage, when their cells no longer run the risk of dispersing, and therefore need to be kept in vitro for a rather long time. Second, following the removal of the zona pellucida, the embryos become very sticky and difficult to manipulate, decreasing the overall efficiency of the technique. One of the main advantages of lentivectormediated transgenesis is the opportunity to exploit the large sum of work successfully invested to develop lentiviral vectors capable of a wide range of biological activities, from constitutive transgene expression to externally controllable gene knockdown (Fig. 6). However, the genotypic mosaicism of the FO animals (issued from the injected oocytes) leads to the differential segregation of individual integrants in their F1 progeny. While possible, the establishment of stable transgenic lines requires proper back-crossing and phenotypic/genotypic screening rounds. In contrast, the ease of generation of large F0 cohorts makes it such that in most situations the phenotypic/genotypic screening rounds are not necessary, since animals can be generated “at will” and tested “in bulk.”

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Figure 6 Examples of some lentiviral vector designs, for gene overexpression (A), gene knock-down with a shRNA (B), and gene regulation (C). The vectors are self-inactivating: the 5 U3 region is from the Rous sarcoma virus (RSV); they all bear the encapsidation signal (including a short gag-derived sequence), the Rev-responsive element (RRE), the central polypurine tract with its termination sequence (cPPT/cTS), and the woodchuck hepatitis virus responsive element (WPRE). “Promoter” designates the transcriptional unit responsible for expression of the transgene, and can be either ubiquitous (human phosphoglycerate kinase, hPGK; ubiquitin, UbC) or tissue specific (e.g., calmodulin-dependent protein kinase II, CamKII; liver-specific transthyretin, mTTR). IRES refers to the internal ribosome entry site of the encephalomyocarditis virus, which directly recruits ribosomes, thus allowing bicistronic expression. pA is a monodirectional poly(A) site derived from the bovine growth hormone gene. The reporter can be, for example, the enhanced green fluorescent protein (eGFP), or one of its variants such as CFP, YFP, dsRed, or Tomato. For gene knockdown, the shRNA is expressed under the control of the H1 or U6 polymerase III promoters. For gene regulation, one of the most powerful designs is based on the Tet-on system allowing gene expression in the presence of doxycycline, an analog of tetracycline. The gene of interest, which can be either a cDNA or an miRNA-embedded shRNA, is under the control of the tetracycline-regulated minimal CMV promoter (TRE), and the tetracycline-dependent transactivator (rtTA, or improved version such as rtTA2S-M2) is expressed under the control of a constitutive promoter, which can also be a ubiquitous or tissue-specific one.

Critical Parameters and Troubleshooting The most important parameters for efficient lentiviral-mediated transgenesis are the vector titer and the quality of its preparation. For lentiviral vector design and preparation, please refer to Barde et al. (2010). In 293T cells, a lentiviral vector stock with a titer below 1 × 108 transducing units (TU)/ml will rarely yield transgenic animals, as will an infec-

tious particle/physical particle ratio much below 1/100. It could be that the early embryo expresses restriction factors that neutralize the incoming vector, and that this can only be overcome by high doses of vector, or that the zona pellucida itself acts as a physical anti-viral barrier that “soaks up” viral particles, which are then unable to reach the plasma membrane. However, it is more likely that the injected dose, even if difficult to assess precisely, is

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Table 2 Troubleshooting Guide for Lentiviral Vector Mediated Transgenesis

Problem

Possible cause

Solution

No plug

Poor breeder male: first time or too old Change the male

Only few embryos recovered after superovulation

Female or male are too old Problem with the hormones

Change female or male Check the correct timing of hormone injections

Vector stock plugged the microcapillary

Poor quality of vector stock

Microcentrifuge the vector stock 5 min at maximum speed and use the supernatant

Low rate of two-cell-stage embryos after the injection

Embryos were not fertilized Poor quality of vector stock

Change male if poor breeder Retry with another vector production

Low yield of animal birth

Poor quality of vector stock

Retry with another vector batch Try to do an inducible expression of the cassette of interest

Embryonic toxicity of the expression cassette

Lentiviral Vector Mediated Transgenesis

Low yield of transgenic Low volume injected due to low animals vector quality or low vector titer

Retry once; if failure, then retry with a new vector production

Transgenic animals The cassette expression is low or is determined by qPCR do not expressed: weak promoter or not express the methylation during embryogenesis transgene of interest

Improve cassette expression by using another promoter; if possible include a gene reporter

simply low. Between 10 and 100 pl of viral stock can be injected; thus, with a titer of 1 × 109 TU/ml, the number of infectious particles placed in the perivitelline zone is around 10 to 100. The quality of the preparation is also a key parameter, since an overly impure solution will not only plug the microcapillary but also increase the quantity of potentially toxic material delivered around the oocyte, leading to its death within minutes of the injection. Lentiviral vectors tend to integrate within the transcribed region of expressed genes. This results in promoting expression of the transgene, in most cases. However, care should be invested in choosing the right transcriptional unit. Some promoters are indeed irreversibly silenced during embryogenesis (e.g., EF1α, CMV) and thus must be avoided. For robust, ubiquitous expression the hPGK or Ubc promoters will be favored. Other designs will allow for specific applications such as tissue-specific or drug-controllable expression or knockdown (Fig. 6). The cloning capacity of lentiviral vectors is limited to 10 to 12 kb. Longer inserts will interfere with infectivity, potentially decreasing viral titer below a critical threshold. In these cases, pronuclear injection might be prefer-

able. Care should also be taken in the design of the vector, to avoid inactivating its gene-transfer potential through the addition of detrimental poly(A) signals, splice sites, or recombination-prone sequences. See also Table 2 for a list of common problems and solutions. Lentiviral vector–mediated transgenesis generally results in genotypic mosaicism, because integration is not immediate. The more delayed this event, the more mosaic the resulting animal. Nevertheless, genotypic characterization of the transgenic mice demonstrates that the integration is on average completed by the 2- to 4-cell stage, so that the degree of mosaicism is low in the resulting animals (Sauvain et al., 2008). Moreover, from a practical standpoint, a genetic mosaicism within transgenic animals is not of great importance as long as they contain several proviruses, since the likelihood is great that each cell will harbor at least one integrant. Indeed, whereas pronuclear injection of naked DNA results in the entire foreign DNA integrating into one single locus, each lentiviral provirus will target a different site. Due to the low rates of lentiviral inactivation during embryogenesis, most proviruses will

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subsequently express the transgene. Thus, despite genotypic heterogeneity, the transgenic animal will present a homogenous phenotype. Nevertheless, such mosaicism can be a limitation for the establishment of pure inbred lines, as each provirus will segregate independently. To circumvent this problem, integration site analysis with a method such as LAM PCR can be used to track integrants along the breeding to generate single-integrant homozygous animals. It is noteworthy that the rates of recovery of homozygous animals after back-crosses appear to be lower than predicted from pure Mendelian transmission, probably because of the functional inactivation of genes targeted by the proviruses (Sauvain et al., 2008).

from preparation of embryos to injection and transfer are completed in 1 week. Pups are born 3 weeks after the transfer, and the genotyping is performed on 4-week-old animals. When embryos are the scope of the analysis, the procedure is correspondingly faster.

Anticipated Results

Fraga D., Meulia T, and Fenster S. 2008 Realtime PCR. Curr. Protoc. Essential Lab. Tech. 00:10.3.1-10.3.34

When applied optimally, the procedure described here yields between 10 and 20 transgenic animals per injected lentiviral vector. The average vector copy per animal can vary from 1 to 20 with a mean around 2 to 5. Optimally, 100 fertilized oocytes should be injected to obtain approximately 80 live 2-cellstage embryos the day after lentiviral vector injection, allowing the transplantation of at least three foster mothers with 16 to 20 embryos each. Under these conditions, eight animals on average will be born from each foster (range of five to twelve). In our laboratory, the average yield of genotypically positive pups is 65%, but ranges from 32% to 100% as a function of the vector and mouse strain used. Our best results are obtained with B6D2F1 mice; the efficiency decreases to 45% with the C57Bl/6 strain.

Time Considerations The whole procedure takes 8 weeks if live animals are sought (see Fig. 7). All the steps

Acknowledgements Isabelle Barde and Sonia Verp (coauthors) contributed equally to this manuscript.

Literature Cited Barde, I., Salmon, P., and Trono, D. 2010. Production and titration of lentiviral vectors. Curr. Protoc. Neurosci. 53:4.21.1-4.21.23. Donovan, J. and Brown, P. 2006. Euthanasia. Curr. Protoc. Immunol. 73:1.8.1-1.8.4.

Gordon, J.W., Scangos, G.A., Plotkin, D.J., Barbosa, J.A., and Ruddle, F.H. 1980. Genetic transformation of mouse embryos by microinjection of purified DNA. Proc. Natl. Acad. Sci. U.S.A. 77:7380-7384. Jahner, D. and Jaenisch, R. 1985. Retrovirusinduced de novo methylation of flanking host sequences correlates with gene inactivity. Nature 315:594-597. Jahner, D., Stuhlmann, H., Stewart, C.L., Harbers, K., L¨ohler, J., Simon, I., and Jaenisch, R. 1982. De novo methylation and expression of retroviral genomes during mouse embryogenesis. Nature 298:623-628. Lois, C., Hong, E.J., Pease, S., Brown, E.J., and Baltimore, D. 2002. Germline transmission and tissue-specific expression of transgenes delivered by lentiviral vectors. Science 295:868-872. Naldini, L., Bl¨omer, U., Gallay, P., Ory, D., Mulligan, R., Gage, F.H., Verma, I.M., and Trono, D. 1996. In vivo gene delivery and stable transduction of nondividing cells by a lentiviral vector. Science 272:263-267.

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Okada, Y., Ueshin, Y., Isotani, A., Saito-Fujita, T., Nakashima, H., Kimura, K., Mizoguchi, A., Oh-Hora, M., Mori, Y., Ogata, M., Oshima, R.G., Okabe, M., and Ikawa, M. 2007. Complementation of placental defects and embryonic lethality by trophoblast-specific lentiviral gene transfer. Nat. Biotechnol. 25:233-237. Robinson, J.P., Darzynkiewicz, Z., Hoffman, R., Nolan, J.P., Orfao, A., Rabinovitch, P.S., and Watkins, S. (eds.) 2010. Current Protocols in Cytometry. John Wiley & Sons, Hoboken, N.J. Sauvain, M.O., Dorr, A.P., Stevenson, B., Quazzola, A., Naef, F., Wiznerowicz, M., Sch¨utz, F., Jongeneel, V., Duboule, D., Spitz, F., and Trono, D. 2008. Genotypic features of lentivirus transgenic mice. J. Virol. 82:7111-7119.

Szulc, J., Wiznerowicz, M., Sauvain, M.O., Trono, D., and Aebischer, P. 2006. A versatile tool for conditional gene expression and knockdown. Nat. Methods 3:109-116. Tiscornia, G., Singer, O., Ikawa, M., and Verma, I.M. 2003. A general method for gene knockdown in mice by using lentiviral vectors expressing small interfering RNA. Proc. Natl. Acad. Sci. U.S.A. 100:1844-1848. WHO. 2004. Laboratory Biosafety Manual, 3rd ed. World Health Organization, Geneva. Wolf, D. and Goff, S.P. 2007. TRIM28 mediates primer binding site-targeted silencing of murine leukemia virus in embryonic cells. Cell 131:4657.

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Assessment of Spontaneous Locomotor and Running Activity in Mice Charles Thomas,1 Stefan Marcaletti,2 and J´erˆome N. Feige2 ´ Center of Phenogenomics (CPG), Ecole Polytechnique F´ed´erale de Lausanne, Lausanne, Switzerland 2 MusculoSkeletal Diseases, Novartis Institute for Biomedical Research, Basel, Switzerland 1

ABSTRACT The locomotor activity of laboratory mice is a global behavioral trait which can be valuable for the primary phenotyping of genetically engineered mouse models as well as mouse models of pathologies affecting the central and peripheral nervous systems, the musculoskeletal system, and the control of energy homeostasis. Basal levels of mouse locomotion can be recorded using infrared monitoring of movements, and further information can be gathered by giving the animal access to a running wheel, which will greatly enhance its spontaneous physical activity. Described here are two detailed protocols to evaluate basal locomotor activity and spontaneous wheel running. Curr. C 2011 by John Wiley & Sons, Inc. Protoc. Mouse Biol. 1:185-198  Keywords: activity r locomotion r wheel running r training

INTRODUCTION The spontaneous locomotor activity of a mouse is a multifactorial parameter primarily controlled by the brain, but also influenced by stress and disease, as well as cardiovascular and metabolic cues. As such, it is an easily accessible parameter providing general information on the physiological and neurological state of the animal for the primary phenotyping of mouse models. In addition, giving free access to a running wheel with which mice will significantly enhance their level of spontaneous physical activity can provide complementary information with stronger focus on peripheral parameters such as cardiovascular function, skeletal muscle contraction, or metabolic homeostasis. Here, two protocols are provided for assessment of spontaneous levels of locomotor activity when housed in experimental cages where locomotion is restricted (see Basic Protocol 1), and of running activity when the housing environment is enriched with running wheels (see Basic Protocol 2). Additional traits and properties are discussed in the Commentary.

ASSESSMENT OF SPONTANEOUS LOCOMOTOR ACTIVITY USING INFRARED DETECTION

BASIC PROTOCOL 1

Infrared detection is one of the best methods to measure mouse locomotor activity, and can be used either in regular housing cages (referred to as home cages) or in combination with metabolic cages where gas exchanges are measured and urine and feces collected to extract metabolic parameters.

Materials Mice (e.g., C57BL/6J) Housing room dedicated to this experiment with restricted access of lab personnel during recording Locomotor activity monitoring system (see Fig. 1): e.g., TSE (http://www.tse-systems.com/), Columbus Instruments (http://www.colinst.com/),

Current Protocols in Mouse Biology 1: 185-198, March 2011 Published online March 2011 in Wiley Online Library (wileyonlinelibrary.com). DOI: 10.1002/9780470942390.mo100170 C 2011 John Wiley & Sons, Inc. Copyright 

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Schematic representation of an infrared-based activity monitoring cage.

or PanLab (http://www.panlab.com/), typically composed of several of the following units: Cage for single mouse housing (with a design and feeding/drinking systems as close as possible to the home cage) Infrared transmitters and receivers for horizontal movement in x direction Infrared transmitters and receivers for horizontal movement in y direction (optional) Infrared transmitters and receivers for vertical movement in z direction (rearings) Food and water consumption monitoring devices (optional, allows correlation of activity with feeding/drinking patterns) NOTE: The position of all infrared transmitters and receivers should be adjustable, and the inter-beam spacing is typically on the order of 1 to 3 cm. Figure 2 provides an example of a home-cage monitoring system (Fig. 2A) and of a metabolic cage including actimetry infrared beams (Fig. 2B). Ideally, home-cage monitoring is the preferred option, as the presence of bedding and a familiar environment minimizes the stress induced by a change of housing conditions. However, metabolic cages can offer advantages such as the ability to measure energy expenditure and to collect urine and feces. NOTE: A computer and software supplied by the manufacturer of the system are required to quantify beam breaks and process the data. Each system has specificities in terms of data processing, but general considerations are addressed in steps 11 and 12, below.

Spontaneous Locomotor and Running Activity in Mice

Prepare equipment and animals 1. Adapt the height of all horizontal infrared transmitters and receivers to the height of a mouse standing on its four legs (Fig. 1, side view). When using a system with bedding, it is important to anticipate that the animal will move the bedding when nesting and to set a sufficient height to avoid detecting these events.

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Figure 2 Example of systems for locomotor activity measurements in home cages (A) and in metabolic cages (B). The home-cage monitoring system (supplied by TSE) monitors horizontal activity in both directions (x and y) as well as vertical activity (z), whereas the metabolic cages (CLAMS, Columbus Instruments) measure vertical activity and horizontal activity in only one direction (x). Both systems integrate different technologies to measure feeding and drinking behavior in real time. Note that the overall design of the home cage system (bedding, feeding/drinking systems) is closer to usual mouse housing cages, but metabolic cages also allow measuring indirect calorimetry and collecting urine and feces.

2. Adapt the height of all vertical transmitters and receivers to the height of a mouse standing on its hind legs. In order to detect only the rearings of the animal, the height should be sufficient to avoid detecting movements in horizontal planes when the mouse is ambulating on its four legs.

3. Remove any object from the cage that might interfere with the detection of the infrared beams (e.g., animal environment enrichment). 4. Start the software and test that all infrared beams are active by manually breaking each individual beam. If some beams do not respond, clean them according to the manufacturer’s instructions and repeat this step until all are functioning correctly. 5. Reset all parameters.

Measure spontaneous locomotor activity 6. Place one animal per cage in the locomotor activity monitoring system. 7. When the mice are in cages different from the home cages (e.g., metabolic cages or cages without bedding or with different feeding/drinking devices), or when animals were previously group housed, let the animals familiarize themselves with the new environment for 24 to 48 hr. 8. Start recording the data for all cages at the same time and run for at least one full circadian period (24 hr). In cases where the throughput is limited because of low numbers of cages, at least one full dark period (during which mice are active) should be covered.

9. Stop recording and return animals to their home cages.

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10. Clean the cages and change the bedding. See Critical Parameters for discussion of measures of mouse anatomy and physiology that can compromise the study, e.g., dysfunctions that lead to poor mobility, or affect circadian rhythm, motivation, and endurance.

Analyze data 11. For each cage, determine the number of beam breaks in every direction as a function of time. When both x and y horizontal movements are detected, the software generally allows calculation of the horizontal distance traveled. Alternatively, the sum of x and y beam breaks can be used as a measurement of total horizontal activity. Some software packages also quantify a parameter called Xamb, which only quantifies horizontal movement across multiple beams in order to distinguish real locomotion from small body movements (e.g., tail flicking). See Critical Parameters for more discussion.

12. Sum the number of beam breaks or the distance traveled in 30- to 60-min time intervals. The duration of binning intervals can be adjusted to modulate temporal resolution. However, the authors do not recommend using bins smaller than 30 min, as inter-animal variability will generally become significant. BASIC PROTOCOL 2

ASSESSMENT OF SPONTANEOUS RUNNING ACTIVITY IN WHEELS The spontaneous locomotor activity of laboratory mice is generally limited by the size of housing cages, as cage size influences locomotor activity (Poon et al., 1997), and wild-derived mice have higher levels of activity than laboratory mice (Nishi et al., 2010). However, physical activity of laboratory mice can be greatly enhanced by giving access to a running wheel that mice are free to use at their convenience.

Materials Mice (e.g., C57BL/6J) Housing room dedicated to this experiment with restricted access of lab personnel during recording Cages with free running wheels and food/water supply: e.g., Lafayette (http://www.lafayetteinstrument.com/), PanLab (http://www.panlab.com/), TSE (http://www.tse-systems.com/), Tecniplast (http://www.tecniplast.it/); Figure 3 shows two examples of typical setups—the wheel diameter is variable depending on the manufacturer but should be ∼12 to 30 cm, and wheels should be equipped with a system to quantify the number of revolutions Computer and software supplied by the manufacturer of the system to quantify the number of revolutions as a function of time Prepare equipment 1. Start the software and verify that all wheel detectors are active by manually rotating each wheel. If some wheels do not respond, clean the revolution detector according to the manufacturer’s instructions and repeat this step until all systems are functioning correctly. 2. Reset all parameters.

Perform experiment 3. Place one animal per cage. Spontaneous Locomotor and Running Activity in Mice

4. Start recording the data for all cages at the same time. Typically, the number of revolutions is converted by the software into a distance, and the total distance per time interval (see step 8 below for details on temporal resolution), as

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Figure 3 Example of cages used to record spontaneous wheel running. (A) A system from Lafayette Instruments, and (B), a system from Tecniplast/Bioseb. Both systems integrate computercontrolled recording of the number of wheel revolutions.

well as the time spent running and the average speed of revolution during running events, are recorded.

5. Run the experiment for at least one full circadian period (24 hr) for up to several weeks when wheel running is used as an exercise training modality (see Anticipated Results). When long-term experiments are performed over several days, it is recommended to regularly verify that data acquisition is occurring correctly by making sure that data are being generated for all cages (e.g., revolution sensors may sometimes malfunction because of dust). Also, minimize changing of bedding, food, and water, and record when these events occur, as they may interfere with measurements by waking the animals during the light phase.

6. Stop recording and return animals to their home cages. 7. Clean the cages and change the bedding. CAUTION: When cages have to be cleaned in washing machines and/or autoclaved, it is very important to remove all of the hardware used to detect wheel revolutions.

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Analyze data 8. For each cage, determine the running distance as a function of time. 9. Sum the total distance run over 30- to 60-min time intervals when studying circadian running patterns. The size of the binning intervals can be adjusted to modulate temporal resolution. However, the authors do not recommend using bins smaller than 30 min, as inter-animal variability will generally become significant. When longer experiments are analyzed, the total distance run daily can be used as a global measure of activity. In addition, some software programs allow determination of the average running speed during running events and the total time spent running.

COMMENTARY Background Information Spontaneous locomotor activity can be analyzed using three main technologies: infrared movement detection, video tracking, and telemetry. While the last method can be useful in situations where activity needs to be monitored in animals housed in groups, or when other telemetric measurements have to be recorded (e.g., temperature, heart rate, blood pressure), telemetric chips require surgery for implantation and suffer from higher variability, most likely because the detection of movement relies on positional tracking of the radiofrequency emitter from the chip. The noninvasive alternatives are generally preferred. Video tracking offers the advantage of allowing elaborate quantification of activity patterns and permitting the same equipment to be used for behavioral tests. However, the systems are generally not compatible with classical housing cages, and their price precludes duplicating the setups in order to analyze several experimental groups in parallel—two factors that generally restrict the duration of videotracking experiments to a few hours. However, because locomotor activity is strongly influenced by circadian rhythms, it is important to measure activity patterns over at least an entire circadian period (and ideally several). Therefore, infrared-based measurement of activity is currently the most widely used approach.

Critical Parameters

Spontaneous Locomotor and Running Activity in Mice

Experimental setup and organization. Multiple commercially available setups are available to monitor spontaneous locomotor activity and free wheel running. Home-cage monitoring is the preferred option, as this minimizes stress and variability of the measurements. When this is not possible, adequate familiarization to the new cage is key to preventing biases linked to the stress of the new environment. For most strains of mice, 24 to 48 hr has been found to give satisfactory famil-

iarization, but familiarization of new strains of mice should be tested and considered satisfactory when activity patterns are similar over two circadian periods. For locomotor activity monitoring, the height of the horizontal beams is important to avoid unwanted detection of bedding movements, while the height of the vertical beams is crucial to detect rearings but not horizontal locomotion. Also, any environment enrichment (e.g., houses or paper for nesting) should be avoided, as this will bias movement detection by creating zones where infrared beam breaks cannot be detected. In addition, it is important to make sure that the cages remain transparent to infrared beams, as repeated washing and autoclaving of plastic cages can alter the infrared transmission. This is typically done using the “beam testing” function of the software by manually breaking every beam within the cage and checking that each beam break is recorded. When designing an activity experiment, it is important to consider the housing conditions before and after the test, as it is absolutely essential to use single-cage housing during activity monitoring to assign activity and running patterns to each individual mouse. When animals are group housed prior to the test, a familiarization period of at least 24 hr (or until activity patterns are stable) is recommended to avoid introducing behavioral biases linked to the new housing environment. Housing density prior to the test can affect behavior (Davidson et al., 2007), so it is recommended to match all animals for housing density during the entire experiment. Conversely, when animals have to be returned to grouped cages after the test, it is recommended to minimize the total duration of monitoring in a singlehousing environment to prevent fighting when the animals are returned to the grouped cages (i.e., 48 to 72 hr of single housing is generally tolerable but this can be adapted according to sex, age, and genetic background).

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In terms of logistics, a critical parameter when testing large experimental groups is the number of cages available. Ideally, one would like to be able to test two experimental groups in parallel to exclude all potential confounding factors from the environment (see below). Unfortunately, this is not always possible due to the cost of the systems. When such limitations occur, experimental groups should be staggered over several days, but animals from all groups should be included in every session to prevent biases linked to inter-session environmental factors. Data acquisition and analysis. Systems from different suppliers differ in the way locomotor activity is measured, as the number of dimensions in which movement is analyzed, as well as data processing, may vary according to the setup and software package used. To assess the correlation between x and y horizontal activity, we analyzed two different strains of mice fed either regular chow or high-fat diet to cover a wide range of activity levels. Figure 4A illustrates the excellent correlation between x and y horizontal locomotor activity under these experimental conditions. These results suggest that the measurement of activity in the second horizontal dimension may be of moderate relevance and can be dispensable, at least for the strains and conditions tested here. However, the main advantage of having measurements of the second horizontal dimension is to extract the total distance traveled, which also strongly correlates with the total horizontal activity (Fig. 4B). In addition, the patterns of activity are very similar whether or not all beam breaks are quantified or whether or not true locomotion is discriminated from small body movements by assessing multiple beam breaks (Xamb) only (Fig. 6A,C). Finally, the correlation of horizontal and vertical activity is also relatively good (Fig. 4C), as both walking and rearing are linked behaviors controlled by central mechanisms (Archer, 1973). Nevertheless, the fact that this correlation is not as good as in the two horizontal directions demonstrates that some behavioral differences exist between walking and rearing. These differences are mainly linked to the fact that rearing of the animal on its hind legs represents either an explorative behavior to analyze its environment or a feeding/drinking behavior to access the food and water supplies provided at the top of the cage (Archer, 1973). To that end, a recent methodology which discriminates the movements of the animal for drinking and feeding from ambulatory activity

is particularly interesting for further deconvoluting the causes of variations in locomotor activity (Goulding et al., 2008). Temporal resolution is important to evaluate the patterns of activity across the circadian period. Appropriate binning to minimize the influence of biological variability while maximizing temporal resolution is key (Dowse and Ringo, 1994). As a general recommendation for the protocols described herein, 30- to 60min bins represent a good compromise. When the analysis of temporal patterns is of primary interest and the ultradian periodicities are to be analyzed with high resolution, deconvolution of temporal data can be achieved by sophisticated analytical procedures, as described in Dowse et al. (2010). Environmental factors and stress. Spontaneous locomotor and running activities are behavioral traits linked to sleep-wake periods and strongly influenced by stress (de Visser et al., 2007; Marston et al., 2008). As a result, environmental parameters, such as temperature, humidity, light patterns, and noise, must be tightly controlled. In particular, it is crucial that no human interference occur during the period of recording (e.g., to change cages or manipulate animals used for other experiments). To achieve this, it is highly recommend that a dedicated room be reserved for these tests. Intra-group variability and group size. Circadian activity patterns have moderate levels of inter-individual variability, but are governed by complex central, as well as metabolic and cardiovascular, effects (see “Physiological parameters affecting activity,” below, for further discussion). Consequently, group size should be sufficiently powered to allow meaningful statistical analyses and adapted to the variability of the exact experimental context (e.g., strain of mice, equipment). When global activity rather than circadian profiles is of interest, variability can also be constrained by integrating data over the entire dark and light phases, but such integration may mask differences in the temporal distribution of activity patterns. Given that the coefficients of variation of activity readouts for strains such as C57BL/6 and DBA/2 mice are in the range of 10% to 20%, studies with 8 to 10 mice per group are sufficient to detect changes in activity of approximately 20% with a level of significance of 5% and a power of 90%. In addition, day-to-day variations in activity patterns within individual animals are lower than inter-animal variability, provided that experimental and environmental conditions are

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Figure 4 The levels of locomotor activity in the three dimensions are correlated. The locomotor activity of wildtype mice on different genetic backgrounds (C57BL/6J and DBA2/J ) and under different dietary challenges (chow diet and high-fat diet for 12 weeks) was measured over 24 hr using home-cage monitoring on the TSE setup described in Fig. 2A, with a 12 hr:12 hr dark:light period. The total beam breaks and distance covered during the light and dark phases were determined, and horizontal activity in the two dimensions was compared (A), and correlated to the distance traveled (B), and to vertical locomotor activity (C).

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kept constant. When studying the effect of a treatment or of an inducible transgenic modification, baseline activity measurements prior to treatment/induction can therefore enhance statistical power by allowing transversal evaluation of the same animal. Sex, age, and genetic background. Sex, age, and genetic background will influence both spontaneous locomotor and wheel-running activities (Koide et al., 2000; Lerman et al., 2002; Konhilas et al., 2004; Lightfoot et al.,

2004; Turner et al., 2005; Ghosh et al., 2010), with C57BL/6 mice being among the most active strains of laboratory mice (Lightfoot et al., 2004). A particularly useful resource illustrating the influence of genetic background and experimental conditions is the Mouse Phenome Database, available at http://phenome.jax.org. It is therefore strongly recommended to use mouse lines on a genetically pure background and to always match experimental groups from the same age and

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Figure 6 Horizontal and vertical locomotor activity of unchallenged or obese wild-type mice in metabolic cages. The locomotor activity of wild-type male C57BL/6J mice described in Figure 5 was measured over 24 hr using metabolic monitoring on the Columbus Instruments CLAMS setup described in Fig. 2B, after a 24-hr familiarization to the new cage environment. Activity counts were integrated over 1-hr intervals (A-C), while feeding and drinking behavior was integrated over 2-hr periods (D). Data are represented as mean ± SEM and * represents a statistical significant difference (p < 0.05) using a 1-way ANOVA followed by a Bonferroni test.

sex. Housing conditions (Poon et al., 1997) and diet (Figs. 5 and 6) will also affect activity and should be controlled to minimize variability.

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Physiological parameters affecting activity. Spontaneous physical activity can be influenced by morphological or genetic defects of various physiological systems or in acute or chronic diseases. The brain is the central regulator of spontaneous activity, as it controls motor function and motivation, and coordinates activity patterns according to circadian, sleep/wake, and feeding periods (Grillner, 1981). Many genetic, pharmacological, or surgical interventions can induce either hypo- or hyper-activity by affecting several of the neurotransmitter systems and different regions of the brain (Viggiano, 2008). Interference with

genes of the core circadian clock and with genes regulating narcolepsy can also result in major dysregulation of both the levels of activity and of their distribution across the circadian period (Vanitallie, 2006; Dibner et al., 2010). Other possible causes of changes in physical activity are structural abnormalities of bones or skeletal muscle (Hara et al., 2002; Connolly et al., 2003). Respiratory and cardiovascular parameters are also major determinants of activity, as defective supply of oxygen and nutrients to skeletal muscle can impair contraction. Consistently, mouse models with chronic obstructive pulmonary diseases and cardiomyopathies may show altered spontaneous locomotor activity (Freeman et al., 2001; Luthje et al., 2009). Finally, metabolic disorders also impact locomotion, as glucose and lipid

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homeostasis in skeletal muscle and in other organs that interact with muscle to provide the energy required for contraction are important for locomotor activity (Goulding et al., 2008; Hennige et al., 2010). Given the multi-factorial influences that converge on the control of spontaneous locomotor and running activities, the authors recommend these tests in combination with other phenotyping and post-mortem analyses to elucidate the potential causes of the phenotypes observed.

Anticipated Results The typical spontaneous locomotor activity of male C57BL/6J mice in a home-cage environment is shown on Figure 5. Because mice are nocturnal, the highest activity is observed during the dark phase when animals are awake. Typically, a first peak of activity is observed in the first 4 to 6 hr of the night, followed by a period of lower activity during the middle of the dark phase. A second hyperactive phase occurs during the last 2 hr of the night and continues for 1 to 2 hr after the light switches on. Despite the fact that horizontal activity is greater than vertical activity (compare axes of Fig. 5A-B versus C-D), circadian activity during the dark phase is strongly correlated in the horizontal and vertical directions, although relative horizontal activity is higher during the light phase. Similar circadian patterns are observed when locomotor activity is measured in metabolic cages after correct familiarization to the new environment (Fig. 6). In addition, when the same mice are analyzed both in the home-cage and metabolic-cage sys-

tems, there is a good correlation between the activity measured with both setups (Fig. 7), demonstrating that experiments performed on different setups can be compared to a certain extent. As discussed above, activity patterns can be altered in response to genetic defects of the nervous, cardiovascular, or musculoskeletal systems. An example of such alterations involves the effects of a high-fat diet in male C57BL/6 mice (Figs. 5 and 6), which most likely results both from locomotor defects linked to the overweight condition and to deleterious consequences of the excess fat in the musculature and in the brain (Kohsaka et al., 2007). Interestingly, a recent study demonstrated that feeding C57BL/6J mice a high-fat diet induces a reduction of locomotor activity even at early time points (Bjursell et al., 2008), and suggests through correlative simulations that this drop in activity may actually be a determinant for the development of obesity. Conversely, increased physical activity is beneficial to metabolic diseases (Ghosh et al., 2010). Nevertheless, it is important to keep in mind that the complex relationship between metabolic homeostasis and physical activity is strongly influenced by the sex and genetic background of the animal, as well as experimental conditions such as the type of diet and the duration of a dietary intervention (Funkat et al., 2004; Feige et al., 2008; Ghosh et al., 2010). When C57BL/6J mice are given free access to a running wheel, they will spontaneously start running following the circadian profile

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Spontaneous Locomotor and Running Activity in Mice

depicted in Figure 8A. The running phase is restricted to the dark period, where the distance covered every hour is on the order of 800 to 1000 m at the beginning of the night and then drops to lower values towards the end of the dark period. The total distance of spontaneous wheel running ranges from 6 to 8 km per day (Fig. 8B), and it is very important to note that the access to wheels enhances locomotor activity by approximately 10-fold, as the distance traveled daily in home cages without wheels is on the order of 600

to 900 meters. Because this dramatic increase in physical activity is sustained through time (Fig. 8B), it is possible to use spontaneous wheel running as an exercise training modality to enhance performance. For example, 2 weeks of spontaneous wheel running enhanced the performance of C57BL/6J mice in treadmill performance tests probing either for endurance or power (Fig. 9; see Marcaletti et al., 2011, for details on performance tests). Interestingly, wheel running had a more pronounced training effect on endurance, with

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the maximal distance increasing by 75% in the endurance test, most likely because wheel running induces mainly prolonged oxidative training with moderate-intensity rather than high-intensity resistance training (Ghosh et al., 2010).

Davidson, L.P., Chedester, A.L., and Cole, M.N. 2007. Effects of cage density on behavior in young adult mice. Comp. Med. 57:355-359.

Time Considerations

Dibner, C., Schibler, U., and Albrecht, U. 2010. The mammalian circadian timing system: Organization and coordination of central and peripheral clocks. Annu. Rev. Physiol. 72:517-549.

The setup of the two protocols described herein is relatively rapid, and the experiments then typically run without any intervention by the experimenter. Locomotor activity should cover a full circadian period, but durations longer than 24 hr covering multiple circadian periods are beneficial to the statistical power and the reliability of the results. In cases where the cages for activity monitoring are different from the regular housing conditions, a minimum of 24 hr of familiarization to the new environment is required, and this familiarization period may need to be extended until activity patterns are stable for particular strains or experimental conditions. This can be done using an independent set of cages dedicated to familiarization, where the costly detectors have not been built in. In the case where such a familiarization setup is not available, the data recorded over the first 24 hr of the experiment should be discarded. Familiarization with the wheel is extremely rapid, as the circadian profile over the first 24 hr is very similar to the subsequent periods (data not shown), and the daily distance covered is stable during the first few days and then slightly increases, most likely due to a training effect (Fig. 8B). When the circadian wheel running pattern is of interest, it is therefore sufficient to assess running activity over a 24-hr period, although integration over several circadian periods is also beneficial. Training experiments to enhance performance using spontaneous wheel running typically last 2 to 4 weeks.

Literature Cited Archer, J. 1973. Tests for emotionality in rats and mice: A review. Anim. Behav. 21:205-235. Bjursell, M., Gerdin, A.K., Lelliott, C.J., Egecioglu, E., Elmgren, A., Tornell, J., Oscarsson, J., and Bohlooly, Y. 2008. Acutely reduced locomotor activity is a major contributor to Western diet–induced obesity in mice. Am. J. Physiol Endocrinol. Metab. 294:E251-E260. Connolly, C.K., Li, G., Bunn, J.R., Mushipe, M., Dickson, G.R., and Marsh, D.R. 2003. A reliable externally fixated murine femoral fracture model that accounts for variation in movement between animals. J. Orthop. Res. 21:843849.

de Visser, L., van den Bos, R., Stoker, A.K., Kas, M.J., and Spruijt, B.M. 2007. Effects of genetic background and environmental novelty on wheel running as a rewarding behaviour in mice. Behav. Brain Res. 177:290-297.

Dowse, H.B. and Ringo, J.M. 1994. Summing locomotor activity data into “bins”: How to avoid artifact in spectral analysis. Biol. Rhythm Res. 25:2-14. Dowse, H., Umemori, J., and Koide, T. 2010. Ultradian components in the locomotor activity rhythms of the genetically normal mouse, Mus musculus. J. Exp. Biol. 213:1788-1795. Feige, J.N., Lagouge, M., and Auwerx, J. 2008. Dietary manipulation of mouse metabolism. Curr. Protoc. Mol. Biol. 84:29B.5.1-29B.5.12. Freeman, K., Lerman, I., Kranias, E.G., Bohlmeyer, T., Bristow, M.R., Lefkowitz, R.J., Iaccarino, G., Koch, W.J., and Leinwand, L.A. 2001. Alterations in cardiac adrenergic signaling and calcium cycling differentially affect the progression of cardiomyopathy. J. Clin. Invest. 107:967974. Funkat, A., Massa, C.M., Jovanovska, V., Proietto, J., and Andrikopoulos, S. 2004. Metabolic adaptations of three inbred strains of mice (C57BL/6, DBA/2, and 129T2) in response to a high-fat diet. J. Nutr. 134:3264-3269. Ghosh, S., Golbidi, S., Werner, I., Verchere, B.C., and Laher, I. 2010. Selecting exercise regimens and strains to modify obesity and diabetes in rodents: an overview. Clin. Sci. (Lond.) 119:5774. Goulding, E.H., Schenk, A.K., Juneja, P., MacKay, A.W., Wade, J.M., and Tecott, L.H. 2008. A robust automated system elucidates mouse home cage behavioral structure. Proc. Natl. Acad. Sci. U.S.A 105:20575-20582. Grillner, S. 1981. Control of locomotion in bipeds, tetrapods and fish. In Handbook of Physiology, Motor Control (V. Brooks, ed.) pp. 1179-1236. Waverly Press, New York. Hara, H., Nolan, P.M., Scott, M.O., Bucan, M., Wakayama, Y., and Fischbeck, K.H. 2002. Running endurance abnormality in mdx mice. Muscle Nerve 25:207-211. Hennige, A.M., Heni, M., Machann, J., Staiger, H., Sartorius, T., Hoene, M., Lehmann, R., Weigert, C., Peter, A., Bornemann, A., Kroeber, S., Pujol, A., Franckhauser, S., Bosch, F., Schick, F., Lammers, R., and Haring, H.U. 2010. Enforced expression of protein kinase C in skeletal muscle causes physical inactivity, fatty liver and insulin resistance in the brain. J. Cell Mol. Med. 14:903913. Kohsaka, A., Laposky, A.D., Ramsey, K.M., Estrada, C., Joshu, C., Kobayashi, Y., Turek,

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F.W., and Bass, J. 2007. High-fat diet disrupts behavioral and molecular circadian rhythms in mice. Cell Metab. 6:414-421.

Marcaletti, S., Thomas, C., and Feige, J.N. 2011. Exercise performance tests in mice. Curr. Protoc. Mouse Biol. 1:141-154.

Koide, T., Moriwaki, K., Ikeda, K., Niki, H., and Shiroishi, T. 2000. Multi-phenotype behavioral characterization of inbred strains derived from wild stocks of Mus musculus. Mamm. Genome 11:664-670.

Marston, O.J., Williams, R.H., Canal, M.M., Samuels, R.E., Upton, N., and Piggins, H.D. 2008. Circadian and dark-pulse activation of orexin/hypocretin neurons. Mol. Brain 1:19.

Konhilas, J.P., Maass, A.H., Luckey, S.W., Stauffer, B.L., Olson, E.N., and Leinwand, L.A. 2004. Sex modifies exercise and cardiac adaptation in mice. Am. J. Physiol. Heart Circ. Physiol. 287:H2768-H2776. Lerman, I., Harrison, B.C., Freeman, K., Hewett, T.E., Allen, D.L., Robbins, J., and Leinwand, L.A. 2002. Genetic variability in forced and voluntary endurance exercise performance in seven inbred mouse strains. J. Appl. Physiol. 92:22452255. Lightfoot, J.T., Turner, M.J., Daves, M., Vordermark, A., and Kleeberger, S.R. 2004. Genetic influence on daily wheel running activity level. Physiol. Genomics 19:270-276. Luthje, L., Raupach, T., Michels, H., Unsold, B., Hasenfuss, G., Kogler, H., and Andreas, S. 2009. Exercise intolerance and systemic manifestations of pulmonary emphysema in a mouse model. Respir. Res. 10:7.

Nishi, A., Ishii, A., Takahashi, A., Shiroishi, T., and Koide, T. 2010. QTL analysis of measures of mouse home-cage activity using B6/MSM consomic strains. Mamm. Genome 21:477485. Poon, A.M., Wu, B.M., Poon, P.W., Cheung, E.P., Chan, F.H., and Lam, F.K. 1997. Effect of cage size on ultradian locomotor rhythms of laboratory mice. Physiol. Behav. 62:1253-1258. Turner, M.J., Kleeberger, S.R., and Lightfoot, J.T. 2005. Influence of genetic background on daily running-wheel activity differs with aging. Physiol. Genomics 22:76-85. Vanitallie, T.B. 2006. Sleep and energy balance: Interactive homeostatic systems. Metabolism 55:S30-S35. Viggiano, D. 2008. The hyperactive syndrome: Metanalysis of genetic alterations, pharmacological treatments and brain lesions which increase locomotor activity. Behav. Brain Res. 194:1-14.

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Design and Generation of Gene-Targeting Vectors ¨ 1,2 Benedikt Wefers,1 Wolfgang Wurst,1,2,3,4 and Ralf Kuhn 1

German Research Center for Environmental Health, Munich, Germany Technical University Munich, Munich, Germany 3 Max-Planck-Institute of Psychiatry, Munich, Germany 4 Deutsches Zentrum f¨ur Neurodegenerative Erkrankungen e.V. (DZNE), Munich, Germany 2

ABSTRACT This unit provides an overview of the major types of mutant alleles that can be generated by gene targeting in ES cells. It presents the growing public resources of premade gene targeting vectors, modified ES cells, and mutant mice. General guidelines for the design of targeting vectors are followed by protocols for the construction of vectors to generate knockout (KO), conditional KO, and subtle mutant alleles. Curr. Protoc. Mouse Biol. C 2011 by John Wiley & Sons, Inc. 1:199-211  Keywords: gene targeting r conditional KO mice r targeting vector design r knockout mice r EUCOMM/KOMP

INTRODUCTION The first section of this unit provides an overview of various types of mutant alleles that can be produced by gene targeting and gene trapping in ES cells. The second section describes the growing public resources of premade gene targeting vectors, modified ES cells, and mutant mice. General guidelines for the design of targeting vectors are followed by protocols for the construction of vectors to generate knockout (KO), conditional KO, and subtle mutant alleles that can be implemented in any molecular biology laboratory.

Mutant Alleles Generated by Gene Targeting In the last two decades, gene targeting in ES cells has been extensively used as a powerful tool to generate predesigned mouse mutants and to study gene function in vivo (Capecchi, 1989, 2005). To generate the reported ∼4000 KO mouse strains, various vector designs were developed to achieve specific gene manipulations. The vast majority of the approaches to design mutant alleles can be classified into five categories (described below and shown in Fig. 1). Germline (classical) KO alleles The classical KO approach disrupts a target gene in the germline by the insertion of a neomycin selection marker gene (neo) into an exon (Fig. 1A). As the first available vector design, the vast majority of KO mice have been created following this approach. Since these “conventional” or “classical” KO mice are homozygous for a null allele in the germline, they provide appropriate models of inherited diseases, leading to embryonic or early postnatal lethality in ∼30% of cases. Besides this application, germline KO mice are often useful to study gene function in adults; however, conditional KO alleles are preferred by many researchers. The published resource of KO mice can be explored using the gene-specific query tool at the mouse genome informatics (MGI) Web page of The Jackson Laboratory (http://www.informatics.jax.org). See Targeting vectors for germline knockout alleles below for the construction of KO vectors and Hasty et al. (2000) for further reading. Current Protocols in Mouse Biology 1: 199-211, March 2011 Published online March 2011 in Wiley Online Library (wileyonlinelibrary.com). DOI: 10.1002/9780470942390.mo100179 C 2011 John Wiley & Sons, Inc. Copyright 

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Figure 1 Mutant alleles generated by gene targeting and gene trapping. To generate a germline KO allele a neomycin-cassette is inserted into the critical exon 2 of the target gene. This leads to the disruption of the gene’s translational reading frame and the premature termination occurs at the neo-polyA site (A). In a gene trap allele, a βgeo-cassette, harboring a splice acceptor (SA), the β-galactosidase/neo coding region and a polyA sequence, is inserted, resulting in the truncation of the wild-type transcript (B). Using gene targeting, a subtle mutation can be inserted into an exon of the target gene (C). In a conditional knockout allele (D), the critical exon 2 is flanked by two Cre recombinase recognition sites (loxP sites). Upon Cre expression, the critical exon is excised, leading to a reading frame shift upon splicing to the downstream exon 3. The “knockout first, conditional-ready” allele (E), which is used by the EUCOMM/KOMP consortia, allows classical and conditional gene targeting (see Fig. 2). Exons of the target gene are shown as numbered rectangles.

Design and Generation of Gene-Targeting Vectors

Gene-trap KO alleles Gene-trap mutagenesis is based on the random integration of a generic gene disruption vector across the genome of embryonic stem cells. Such vectors simultaneously mutate a gene at the site of insertion and report its expression by insertion of a β-galactosidase reporter (lacZ) gene (Fig. 1B). Since a single vector can be used to target a large number of genes, gene trapping is a high-throughput insertional mutagenesis approach used to establish libraries of mutant ES cell clones. Gene-trap vectors are promoterless and contain a splice acceptor element and an ATG-less hybrid-coding region for the lacZ reporter and the neo selection marker (βgeo). Therefore, gene trapping allows only the targeting of genes that are sufficiently expressed in ES cells. The inclusion of pairs of wild-type and mutant loxP and FRT sites in conditional gene-trap vectors further allows the inversion of the gene-trap cassette by Cre or FLP recombinase for conditional gene inactivation (Schnutgen et al., 2003, 2005). The >400,000 gene-trap ES cell clones that are publicly available can be searched at the International Gene Trap Consortium (IGTC) Web page (http://www.genetrap.org) and the genomic vector insertion sites are mapped

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on the Ensembl mouse genome server (http://www.ensembl.org). For further reading, see Evans et al. (1997), Hansen et al. (2003), and K¨uhn and Wurst (2005).

Subtle mutation alleles In many instances, human diseases have been associated with disease alleles harboring missense mutations that cause a partial loss- or gain-of-function of the affected protein. The effect of such mutations can be studied in mouse models by codon exchange in the related mouse gene (Fig. 1C). See Targeting vectors for insertion of subtle mutations below for the construction of vectors that introduce subtle (e.g., missense) mutations. Conditional KO alleles Conditional mutagenesis enables inactivation of a target gene via Cre/loxP recombination in somatic cells. In the conditional allele, a functionally essential (critical) exon is flanked by two loxP sites that do not interfere with gene function (Fig. 1D). Conditional mutant mice are obtained by crossing the conditional strain with a transgenic line that expresses Cre recombinase or an externally inducible Cre fusion protein under control of a cell type-specific promoter. Thereby, the inactivation of the target gene is either restricted to a selected cell type without temporal control or can be induced at a chosen time. Conditional mutagenesis enables the study of gene function in a very precise manner in a cell type of choice. The published resources for conditional KO mice and relevant citations can be identified using the gene-specific query tool at the mouse genome informatics Web page of the Jackson laboratory (http://www.informatics.jax.org). See Targeting vectors for conditional knockout alleles below for the construction of conditional KO vectors and Branda and Dymecki (2004), Kwan (2002), Rajewsky et al. (1996), and Torres and K¨uhn (1997) for further reading. KO first, conditional-ready alleles The large-scale EUCOMM/KOMP mutagenesis programs, which are coordinated by the International Knockout Mouse Consortium (IKMC), generate a genome-wide resource of gene specific targeting vectors, targeted ES cell clones, and mutant mouse strains. The EUCOMM/KOMP alleles are mostly of the “KO first, conditional-ready” type (Fig. 1E). This design initially disrupts the target gene by the intronic insertion of a gene disruption cassette (KO first, including a splice acceptor, lacZ reporter, and neo-resistance gene) that is flanked by FRT sites. In addition, two loxP sites flank a “critical” exon of the target gene. Upon FLP-mediated deletion of the gene disruption cassette, the EUCOMM/KOMP KO allele is converted into a standard conditional allele. These resources are accessible via the EUCOMM, KOMP, and IKMC Web pages (http://www.eucomm.org, http://www.komp.org, and http://www.knockoutmouse.org). Since this resource will ultimately provide a genome-wide collection of KO and conditional KO reagents, it is described in more detail in Ready-made EUCOMM/KOMP targeting vectors, ES cells and mouse lines below. Project Planning Due to the substantial existing and growing sources of information on targeting vectors, ES cell clones, and mutant mouse strains, a mutagenesis project should be started by searching the relevant databases for preexisting materials to avoid potential duplication. This can be conveniently achieved by searching for the name of the gene of interest at the MGI (http://www.informatics.jax.org), IGTC (http://www.genetrap.org), and IKMC (http://www.knockoutmouse.org) Web pages. If these databases do not yield materials relevant to the project requirements, it may be necessary to consider generating a novel gene-targeting vector, as described in the protocols below.

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Ready-Made EUCOMM/KOMP Targeting Vectors, ES Cells, and Mouse Lines The EUCOMM/KOMP alleles are mostly of the “KO first, conditional-ready” type (Fig. 2) (Friedel et al., 2007). This design initially disrupts the target gene by the intronic insertion of a gene disruption cassette that includes a splice acceptor element, a βgalactosidase reporter gene, and a neo-resistance gene. In these “knock-out first” alleles, gene function is inactivated by splicing of upstream exons to the splice acceptor site of the targeting cassette (targeted mutation 1a, Tm1a, Fig. 2A). Most EUCOMM/KOMP targeting vectors contain a targeting cassette with a neo resistance that is driven by its own β-actin promoter to allow the targeting of all genes, irrespective of their expression status in mouse ES cells. Promoterless vectors, which utilize the promoter of the targeted gene to drive the expression of the neo resistance, are also used for the targeting of genes that are expressed in ES cells. In addition to the gene disruption cassette, the targeted EUCOMM/KOMP alleles contain two loxP sites that flank a critical exon of the target gene (Fig. 2A). The loxP-flanked critical exon, as the target for Cre-mediated excision, is determined by the following criteria: (1) its deletion causes a translational reading frame shift in the remaining mRNA that leads to the production of a shortened, mutant protein, (2) the exon is present in all transcript splice variants of the targeted gene, (3) the size of the exon is <1 kb, to ensure that the distance of the loxP sites is minimal for efficient Cre recombination, and (4) the size of the flanking introns is at least 0.5 kb, so that loxP sites can be placed in nonconserved regions that are not required for endogenous splicing. In many cases, the second exon of a target gene fulfills these critical criteria, but in some instances, several small exons are combined into a group of critical exons. Design and Generation of Gene-Targeting Vectors

The “KO first,” Tm1a allele can be converted into a classical KO allele by breeding the Tm1a mice with a Cre transgenic germline deleter strain. The neomycin selection cassette and the critical exon are deleted in the germline, resulting in a β-galactosidase-tagged, classical KO allele (Tm1b, “beta-Gal reporter,” Fig. 2B). Furthermore, the Tm1a-type

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allele can be converted into a Tm1c allele for conditional mutagenesis by the use of the two FLP recombinase recognition (FRT) sites that flank the gene disruption cassette. The FLP-mediated deletion is typically performed by breeding the Tm1a mice with an FLP transgenic germline deleter strain (for a line in the C57BL/6N background see Kranz et al., 2010). Upon the removal of the gene disruption cassette, the remaining loxP sites that are flanking the critical exon delineate the configuration of a conditional allele (Tm1c, “conditional,” Fig. 2C). This functional conditional allele can subsequently be inactivated in somatic cells by breeding Tm1c mice to a mouse line that expresses Cre recombinase in a time- and tissue-specific manner, resulting in the nonfunctional Tm1d allele (“deletion,” Fig. 2D). At present, the EUCOMM/KOMP resource offers >10,000 targeting vectors, >8,000 genes targeted in the C57BL/6N-derived ES cell line JM8 (Pettitt et al., 2009), and >700 established mouse strains. These resources are accessible through the programs EUCOMM and KOMP (http://www.eucomm.org; http://www.komp.org) and the IKMC (http://www.knockoutmouse.org) Web pages. The vectors that are generated by EUCOMM/KOMP can also be used to assemble vectors with other targeting cassettes. For this purpose, intermediate versions of the targeting vectors can be ordered, and a targeting cassette of choice can be inserted by Gateway (Invitrogen) cloning.

Design Rules for Gene Targeting Vectors Upon the introduction of genomic DNA fragments into ES cells, the DNA becomes degraded in the vast majority of cells. However, some cells either integrate the DNA into a random position of the genome or align and recombine the fragment with an endogenous homologous gene copy by homologous recombination (HR). The term “gene targeting” refers to the utilization of the HR mechanism to introduce a preplanned mutation into a selected target gene. For this purpose, specific gene targeting vectors can be constructed that combine several kilobases (kb) of gene-derived genomic sequence with a mutation of interest. Since HR of a gene-targeting vector with an endogenous locus occurs at a low rate of ∼10−5 –10−6 of electroporated ES cells, a drug selection marker gene must be included into these vectors, in order to expand the small number of cells containing stable integrations. The efficiency at which gene targeting is achieved is usually expressed as the ratio of drug-resistant ES cell clones harboring homologous recombined alleles versus clones containing random vector integrations. This ratio varies greatly between ∼0.1% and 30%, depending on the targeted gene and the specific design of gene targeting vectors. Some genomic regions, in particular the Y-chromosome, have so far been resistant to gene targeting attempts, for unclear reasons. Gene targeting in ES cells was first described in 1987 for the Hprt locus in the classical publication by Thomas and Capecchi, which defined the length of the vector homology region as a critical factor for the HR frequency (Thomas and Capecchi, 1987). In addition, several other parameters and vector design rules have been found useful to maximize the recovery of targeted ES cell clones and to ensure that the genetic modification results into the intended mutant allele. Length of the genomic homology regions The combined length of the vector genomic homology regions should ideally be in the range of 6 to 12 kb. If the genomic sequence is unevenly split among the vector homology arms, the shortest arm should include 1 to 2 kb. Source of genomic DNA for vector construction It has been found that even small sequence differences of just 0.1% between the homology sequences of a targeting vector and the genomic target locus leads to a strong decline of the HR frequency (te Riele et al., 1992). Therefore, targeting vectors should be built from genomic DNA that is derived from the same mouse strain that was used for the derivation

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of the ES cell line employed for gene targeting. Since the mouse genome sequence is derived from the C57BL/6J strain, any region of this genome can be easily obtained as sequence-confirmed BAC clones. Clone identifiers can be retrieved from the Ensembl genome browser (http://www.ensembl.org) and ordered from various distributors (e.g., ImaGenes, http://www.imagenes-bio.de). To ensure efficient recombination, targeting constructs based on the C57BL/6 genomic sequence should be transfected into C57BL/6derived ES cell lines, such as the JM8 line (Pettitt et al., 2009) or (C57BL/6x129)-F1 hybrid lines, e.g., V6.5 or IDG3.2 cells (Eggan et al., 2001; Hitz et al., 2009).

Positive and negative selection markers Targeting vectors must include a drug-selectable resistance marker gene in order to positively select ES cell clones harboring stable vector integrations. The most commonly used selection marker is the neomycin phosphotransferase gene (neo) that confers resistance to the neomycin analog G418. The use of a neo expression cassette driven by the Pgk1 gene promoter that is available from Addgene (http://www.addgene.org; plasmid 13442) is most common. See Targeting vectors for conditional knockout alleles below for a neo-cassette flanked by FRT sites that can be deleted by FLP recombinase. In addition, the selection of feeder-dependent ES cells requires the use of embryonic fibroblasts established from a neo-transgenic mouse strain, e.g., C57BL/6J-Tg(pPGKneobpA)3Ems/J mice (http://www.jax.org; stock 002356). Negative selection marker genes are used to reduce the number of G418-resistant ES cell colonies harboring random vector insertions and thereby to enrich for homologous recombined clones. The commonly used negative selection markers are the herpes simplex virus (HSV)-derived thymidine kinase (TK) gene (Mansour et al., 1990) and the diphteria toxin A-chain (DTA) gene (Yagi et al., 1990). Negative selection using HSVTK requires the addition of Gancyclovir or FIAU into the culture medium. HSV-TK or DTA expression cassettes are placed at the end of one of the targeting vectors homology arms and will exert their cytotoxic function upon the random genomic integration of the complete vector. In case of HR, the heterologous HSV-TK or DTA sequences are not incorporated into the genome. Random vector integration often results in the partial deletion of vector sequences. If such an event inactivates the negative selection marker, random integrant ES cell clones survive the selection procedure. Therefore, negative selection usually results in a 3- to 10-fold enrichment of homologous recombined ES cell clones. The HSV-TK cassette can be retrieved from, e.g., the plasmid pEasyFlox (Addgene, plasmid 11725) and the DTA cassette from the Addgene plasmid 13440.

Structure of the target locus The design of a gene-targeting vector requires careful analysis of the genomic structure of the target locus. For this purpose, the genomic sequence should first be downloaded from a genome database (e.g., http://www.ensembl.org), imported into sequence analysis software (e.g., VectorNTI, Invitrogen; DNA workbench, CLC bio), and annotated for the position of the exons, alternatively spliced exons, the exon reading frames, functional protein domains, and repetitive elements. This information enables one to build a reliable targeting vector that mutagenizes the target gene in the desired manner. Further details for the construction of KO, conditional KO, and subtle mutation alleles are given in the protocols below.

Design and Generation of Gene-Targeting Vectors

Screening for Recombinant ES Cell Clones The development of a reliable screening strategy for the identification of recombinant ES cell clones is as important as the design of the targeting vector. PCR-based screening strategies can help to identify recombinant clones (Gomez-Rodriguez et al., 2008), but the final proof of the integrity of a targeted locus can be achieved only by Southern

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blot analysis. Therefore, we recommend performing the screening of transfected ES cell colonies directly by Southern blotting (Southern, 2006). This can be conveniently achieved by the culture of ES cells, DNA preparation, and DNA restriction in 96-well plates (Nagy et al., 2003). For the design of a Southern-based screening strategy, a sequence model of the recombined locus should be constructed and compared to the wild-type gene using software for DNA sequence analysis. Using these data, genomic segments for use as labeled hybridization probes can be identified. For a targeted locus, two sequence areas of 0.7 to 1 kb, which are located upstream and downstream of the targeting vector homology regions and that are free of repetitive elements, should be identified. These segments can be amplified by PCR, cloned, and used as 5 - and 3 genomic probes for Southern blot hybridization. The Southern blot screening of several hundred or thousands of DNA samples requires considerable amounts of a restriction enzyme able to detect a clear size difference between the wild-type locus and the targeted allele. To identify restriction enzymes suitable for Southern screening the wild-type and mutant alleles should be analyzed and compared for the distribution of recognition sites of inexpensive restriction enzymes (e.g., EcoRI, BamHI, HindIII, EcoRV, PstI, and XbaI). In most cases, this analysis results in the identification of at least one enzyme suitable for Southern screening. Alternatively, to facilitate the identification of mutant ES cell clones, the design of the targeting vector can be adapted to include suitable restriction sites between the homology regions and the neo selection marker. PROTOCOLS FOR THE CONSTRUCTION OF GENE TARGETING VECTORS Since gene-targeting vectors are assembled from genomic and heterologous sequences, any method for the handling and joining of recombinant DNA fragments can be used for vector construction. However, the most popular and convenient protocols are based on HR in E. coli (recombineering, ET cloning; Testa et al., 2003, 2004; Valenzuela et al., 2003; Chan et al., 2007; Wu et al., 2008; Lee et al., 2009; Fu et al. 2010) and the directed cloning of PCR-amplified genomic fragments into plasmid backbones that provide the heterologous vector elements. As recombineering and ET cloning protocols require a set of specialized reagents and expertise, we therefore provide here protocols that are based on simple PCR and cloning methods and that can be implemented in any molecular biology laboratory.

Materials pGKneobpA (Addgene, http://www.addgene.org, plasmid 13442) pEasyfloxII-DTA (submitted to Addgene by R. K¨uhn) Genomic BAC clones (e.g., from ImaGenes, http://www.imagenes-bio.de or Source BioScience, http://www.lifesciences.sourcebioscience.com) Proofreading DNA polymerase, e.g., Phusion (New England Biolabs) or Herculase-II (Stratagene) DNA oligonucleotides as PCR primers Bioinformatic software for sequence analysis and vector design (e.g., VectorNTI, CLC Sequence Viewer, or Geneious) Software for PCR primer design (e.g., VectorNTI or Primer3, http://frodo.wi.mit.edu/primer3/) Generation of Targeting Vectors: General Procedure The steps below apply to the construction of any type of gene targeting vector, requiring the analysis of the target gene’s structure, the selection of PCR primers, and the development of a Southern blotting strategy. Specific rules for the construction of knockout, conditional, or subtle mutation targeting vectors are given in the following three subsections. 1. Download the genomic sequence of the target gene from a genome database (e.g., Ensembl) Current Protocols in Mouse Biology

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2. Analyze and annotate the target sequence for exon positions, alternatively spliced exons, reading frames, and functional protein domains to identify a critical exon using sequence analysis software (follow guidelines described above in Design rules for gene targeting vectors). 3. Develop a Southern blot (Southern, 2006) screening strategy using 5 - and 3 -probes and identify suitable restriction enzymes following the guidelines described in Design rules for gene targeting vectors. 4. Order an BAC clone containing the target gene (see Design rules for gene targeting vectors, above) from a genomic library (e.g., RPCI-23 for C57BL/6-derived ES cells). 5. Design suitable primer pairs for the PCR amplification of the homology arms (see detailed information below in Targeting vectors for germline knockout alleles, Targeting vectors for conditional knockout alleles, and Targeting vectors for insertion of subtle mutations). 6. Amplify homology arms by high-fidelity PCR using a proofreading polymerase. 7. Clone the homology arms into an appropriate vector backbone to obtain the targeting vector (see detailed information in Targeting vectors for germline knockout alleles, Targeting vectors for conditional knockout alleles, and Targeting vectors for insertion of subtle mutations). 8. Confirm the integrity of targeting vector homology regions by sequence analysis to avoid the transfer of undesired mutations into the target gene. 9. Upon transfection of ES cells, select for recombinant clones using the positive selection marker provided by the vector backbone (most commonly neomycin). 10. Screen for correctly targeted clones by Southern blotting (Southern, 2006). Generation of Specific Targeting Vectors Since homologous recombination can be used to generate different types of targeted alleles, the following sections present information for the generation of targeting vectors for germline KO and conditional KO alleles, and for the insertion of subtle mutations.

Targeting vectors for germline knockout alleles For the construction of a targeting vector to generate a classical germline knockout allele, use the pGKneobpA plasmid backbone that includes KpnI, ApaI, SalI, ClaI, HindII, EcoRV, and EcoRI sites for the integration of the 5 homology arm (segment A) and NotI and SacII sites for the integration of the 3 homology arm (segment B) (Fig. 3B). Furthermore, for positive selection, the vector contains a neo expression cassette. The combined size of the two homology arms should be 6 to 12 kb, with each arm comprising 2 to 5 kb. The 5 end of the 5 homology arm should include a unique restriction site for the linearization of the targeting vector prior to electroporation into ES cells. The junction of the two homology arms must be located in a critical exon of the target gene to disrupt the target protein by insertion of the neo expression cassette (Fig. 3A). Moreover, the primers for PCR amplification must include appropriate restriction sites for cloning of the fragment into the pGKneobpA plasmid.

Design and Generation of Gene-Targeting Vectors

Upon cloning of the homology arms into the two restriction sites of the vector, the targeting vector contains the homology arms and the critical exon, which is disrupted by the neo expression cassette (Fig. 3C). The presence of the neo cassette in the targeted knockout allele makes it easy to identify recombined clones: e.g., compared to the wild-type

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Figure 3 Targeting vectors for germline knockout alleles. Genomic DNA or a BAC clone (A) is used for the PCR amplification of homology arms, which are then cloned into the plasmid backbone (B). The resulting targeting vector (C) is integrated into the target gene locus by homologous recombination, leading to a neomycin-positive knockout allele (D). Exons of the target gene are shown as numbered rectangles.

allele, a Southern blot using the 5 probe and restriction enzyme 1 (“Re1”) identifies an increase in band size of ∼2 kb, while using the 3 probe with restriction enzyme 2 (“Re2”) leads to a smaller fragment as compared to the wild-type allele (Figure 3D).

Targeting vectors for conditional knockout alleles For the construction of a conditional targeting vector, we recommend the generic plasmid backbone pEasyfloxII-DTA, which contains two loxP sites, a FRT-flanked neo expression cassette for positive selection, and a DTA gene for negative selection (Fig. 4B). This vector harbors SfiI, NotI, AscI, AsiSI, and PacI sites for the insertion of the 5 homology arm (segment A), an SbfI and SalI site for the insertion of the segment B (which contains the critical exon), and an XhoI site for the cloning of the 3 homology arm (segment C). The SfiI or NotI sites enable convenient linearization of the targeting vector prior to the electroporation of ES cells. The 5 homology arm A should have a size of 1 to 5 kb (Fig. 4A), the loxP-flanked segment B should comprise 0.5 to 2 kb and include one or more exons of the target gene to inactivate the target gene by Cre-mediated deletion. The third genomic segment C represents the 3 -homology region with a size of 2 to 5 kb. The size of segment C should be at least equal to the length of segment B to provide sufficient sequence space for HR downstream of the second loxP site. The primers for the PCR amplification of the three segments must include the restriction sites chosen for cloning of the respective genomic fragment into pEasyfloxII-DTA. As part of the vector design, the absence of these sites within the amplified segments must be confirmed. After cloning of the homology arms into the three restriction sites of the plasmid, the targeting vector contains the 5 -homology region, the loxP-flanked exchange cassette, and the 3 -homology arm (Fig. 4C).

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Figure 4 Targeting vectors for conditional knockout alleles. Genomic DNA or a BAC clone (A) is used for the PCR amplification of homology arms, which are then cloned into the pEasyFlox-II-DTA plasmid backbone (B). The resulting targeting vector (C) is used for the mutagenesis of the target gene by homologous recombination, leading to a neomycin-positive mutant allele (D). Finally, the neomycin cassette is excised using FLP recombinase to obtain the conditional allele (E). Exons of the target gene are shown as numbered rectangles.

Recombination of the targeting vector with the wild-type allele can be easily identified by Southern blot analysis as shown in the example in Figure 4E. As compared to the wild-type, digestion with restriction enzyme 1 (“Re1”, detected by the 5 probe) leads to an increased band size of ∼2 kb, and digestion with restriction enzyme 2 (“Re2”, detected by the 3 probe) leads to a band of reduced size. The presence of the loxP site can be confirmed by double digestion with Re2 and XhoI, using the 3 probe for detection (Fig. 4E). Finally, the pgk-neo cassette must be excised using FLP recombinase to obtain the functional (neo-free) allele for conditional knockout (Fig. 4E). This recombination event should be confirmed by Southern blot analysis using the 5 - and 3 -probes for hybridization.

Design and Generation of Gene-Targeting Vectors

Targeting vectors for insertion of subtle mutations The strategy for insertion of subtle mutations follows the guidelines for conditional gene knockout described in the preceding section Targeting vectors for conditional knockout alleles. The pEasyfloxII-DTA plasmid can be used as a vector backbone (Fig. 5B), but the cloning strategy is modified. The size of the homology arms follows the rules described above in Targeting vectors for conditional knockout alleles, including the necessity of a unique restriction site at the 5 end of segment A. The desired mutation can be inserted into a selected position of the target gene by the use of a PCR approach for site-directed mutagenesis (Cormack, 2001; Elion et al., 2007) to amplify segment B that harbors

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Figure 5 Targeting vectors for insertion of subtle mutations. Genomic DNA or a BAC clone (A) is used for the PCR amplification of homology arms. A subtle mutation and a silent mutation creating the restriction site Re2 are introduced by the use of a PCR approach for site-directed mutagenesis. The homology arms are then cloned into the plasmid backbone (B). The resulting targeting vector (C) is integrated into the target gene locus by homologous recombination, leading to a neomycinpositive mutant allele (D). Finally, the neomycin cassette is excised using FLP recombinase to obtain the mutant allele harboring the subtle mutation (E). Exons of the target gene are shown as numbered rectangles.

the subtle mutation (Fig. 5A). If possible, a new restriction site should be included by nearby silent nucleotide changes to enable the Southern blot detection of the mutant allele (Fig. 5D, E). The primers for the PCR amplification of the three segments must include the restriction sites chosen for cloning of the respective genomic fragment into pEasyfloxII-DTA. Segment B should be inserted into the SalI and XhoI sites to remove the intervening loxP site. Upon completion of vector cloning (Fig. 5C), the homology regions of the targeting construct should be sequenced to confirm the insertion of the desired mutation and the integrity of the critical exon’s reading frame, and to exclude other undesired mutations. Analogous to the conditional targeting vector (Targeting vectors for conditional knockout alleles), HR alleles generated with a subtle mutation-targeting vector can be initially identified through Southern blot analysis by the presence of the pgk-neo cassette (Fig. 5D). Finally, the pgk-neo cassette must be excised using FLP recombinase to obtain the subtle mutant (neo-free) allele (Fig. 5E), which should be confirmed by Southern analysis using the 5 and 3 hybridization probes. CONCLUDING REMARKS Until recently, scientists requiring a KO or conditional KO mouse for phenotypic studies were forced to generate these mutants themselves, starting with the planning and cloning

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of targeting vectors for mutagenesis in ES cells. This situation has dramatically changed in recent years with the activity of large-scale mutagenesis projects that provide premade targeting vectors, ES cell clones, and mutant lines to the scientific community. These central resources support scientists to focus on scientific questions instead of spending time on mutagenesis procedures. However, since resolving one scientific question raises new questions, targeting vectors will also be required in the future to generate customized mutant alleles.

LITERATURE CITED Branda, C.S. and Dymecki, S.M. 2004. Talking about a revolution: The impact of site-specific recombinases on genetic analyses in mice. Dev. Cell 6:7-28. Capecchi, M.R. 1989. The new mouse genetics: Altering the genome by gene targeting. Trends Genet. 5:70-76. Capecchi, M.R. 2005. Gene targeting in mice: Functional analysis of the mammalian genome for the twenty-first century. Nat. Rev. Genet. 6:507-512. Chan, W., Costantino, N., Li, R., Lee, S.C., Su, Q., Melvin, D., Court, D.L., and Liu, P. 2007. A recombineering based approach for high-throughput conditional knockout targeting vector construction. Nucleic Acids Res. 35:e64. Cormack, B. 2001. Directed mutagenesis using the polymerase chain reaction.Curr. Protoc. Mol. Biol. 37:8.5.1-8.5.10. Eggan, K., Akutsu, H., Loring, J., Jackson-Grusby, L., Klemm, M., Rideout, W.M. 3rd, Yanagimachi, R., and Jaenisch, R. 2001. Hybrid vigor, fetal overgrowth, and viability of mice derived by nuclear cloning and tetraploid embryo complementation. Proc. Natl. Acad. Sci. U.S.A. 98:6209-6214. Elion, E.A., Marina, P., and Yu, L. 2007. Constructing recombinant DNA molecules by PCR. Curr. Protoc. Mol. Biol. 78:3.17.1-3.17.12. Evans, M.J., Carlton, M.B., and Russ, A.P. 1997. Gene trapping and functional genomics. Trends Genet. 13:370-374. Friedel, R.H., Seisenberger, C., Kaloff, C., and Wurst, W. 2007. EUCOMM–the European conditional mouse mutagenesis program. Brief. Funct. Genomic Proteomic 6:180-185. Fu, J., Teucher, M., Anastassiadis, K., Skarnes, W., and Stewart, A.F. 2010. A recombineering pipeline to make conditional targeting constructs. Methods Enzymol. 477:125-144. Gomez-Rodriguez, J., Washington, V., Cheng, J., Dutra, A., Pak, E., Liu, P., McVicar, D.W., and Schwartzberg, P.L. 2008. Advantages of q-PCR as a method of screening for gene targeting in mammalian cells using conventional and whole BAC-based constructs. Nucleic Acids Res. 36:e117. Hansen, J., Floss, T., Van Sloun, P., Fuchtbauer, E.M., Vauti, F., Arnold, H.H., Schnutgen, F., Wurst, W., von Melchner, H., and Ruiz, P. 2003. A large-scale, gene-driven mutagenesis approach for the functional analysis of the mouse genome. Proc. Natl. Acad. Sci. U.S.A. 100:9918-9922. Hasty, P., Abuin, A., and Bradley, A. 2000. Gene targeting, principles, and practice in mammalian cells. In Gene Targeting: A Practical Approach, 2nd ed. (A.L. Joyner, ed.) pp. 1-35. Oxford University Press, Oxford. Hitz, C., Steuber-Buchberger, P., Delic, S., Wurst, W., and Kuhn, R. 2009. Generation of shRNA transgenic mice. Methods Mol. Biol. 530:101-129. Kranz, A., Fu, J., Duerschke, K., Weidlich, S., Naumann, R., Stewart, A.F., and Anastassiadis, K. 2010. An improved Flp deleter mouse in C57Bl/6 based on Flpo recombinase. Genesis 48:512-520. K¨uhn, R. and Wurst, W. 2005. Mouse mutagenesis and gene function. In Encyclopedia of Genetics, Genomics, Proteomics and Bioinformatics, p. 18. John Wiley & Sons, Hoboken, N.J. Kwan, K.M. 2002. Conditional alleles in mice: Practical considerations for tissue-specific knockouts. Genesis 32:49-62. Lee, S.C., Wang, W., and Liu, P. 2009. Construction of gene-targeting vectors by recombineering. Methods Mol. Biol. 530:15-27. Mansour, S.L., Thomas, K.R., Deng, C.X., and Capecchi, M.R. 1990. Introduction of a lacZ reporter gene into the mouse int-2 locus by homologous recombination. Proc. Natl. Acad. Sci. U.S.A. 87:7688-7692. Nagy, A., Gertsenstein, M., Vintersten, K., and Behringer, R. 2003. Manipulating the Mouse Embryo, third edition ed. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York. Design and Generation of Gene-Targeting Vectors

Pettitt, S.J., Liang, Q., Rairdan, X.Y., Moran, J.L., Prosser, H.M., Beier, D.R., Lloyd, K.C., Bradley, A., and Skarnes, W.C. 2009. Agouti C57BL/6N embryonic stem cells for mouse genetic resources. Nat. Methods 6:493-495.

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Rajewsky, K., Gu, H., Kuhn, R., Betz, U.A., Muller, W., Roes, J., and Schwenk, F. 1996. Conditional gene targeting. J. Clin. Invest. 98:600-603. Schnutgen, F., Doerflinger, N., Calleja, C., Wendling, O., Chambon, P., and Ghyselinck, N.B. 2003. A directional strategy for monitoring Cre-mediated recombination at the cellular level in the mouse. Nat. Biotechnol. 21:562-565. Schnutgen, F., De-Zolt, S., Van Sloun, P., Hollatz, M., Floss, T., Hansen, J., Altschmied, J., Seisenberger, C., Ghyselinck, N.B., Ruiz, P., Chambon, P., Wurst, W., and von Melchner, H. 2005. Genomewide production of multipurpose alleles for the functional analysis of the mouse genome. Proc. Natl. Acad. Sci. U.S.A. 102:7221-7226. Southern, E. 2006. Southern blotting. Nat. Protoc. 1:518-525. te Riele, H., Maandag, E.R., and Berns, A. 1992. Highly efficient gene targeting in embryonic stem cells through homologous recombination with isogenic DNA constructs. Proc. Natl. Acad. Sci. U.S.A. 89:5128-5132. Testa, G., Zhang, Y., Vintersten, K., Benes, V., Pijnappel, W.W., Chambers, I., Smith, A.J., Smith, A.G., and Stewart, A.F. 2003. Engineering the mouse genome with bacterial artificial chromosomes to create multipurpose alleles. Nat. Biotechnol. 21:443-447. Testa, G., Vintersten, K., Zhang, Y., Benes, V., Muyrers, J.P., and Stewart, A.F. 2004. BAC engineering for the generation of ES cell-targeting constructs and mouse transgenes. Methods Mol. Biol. 256:123-139. Thomas, K.R. and Capecchi, M.R. 1987. Site-directed mutagenesis by gene targeting in mouse embryoderived stem cells. Cell 51:503-512. Torres, R.M. and K¨uhn, R. 1997. Laboratory Protocols for Conditional Gene Targeting. Oxford University Press, Oxford. Valenzuela, D.M., Murphy, A.J., Frendewey, D., Gale, N.W., Economides, A.N., Auerbach, W., Poueymirou, W.T., Adams, N.C., Rojas, J., Yasenchak, J., Chernomorsky, R., Boucher, M., Elsasser, A.L., Esau, L., Zheng, J., Griffiths, J.A., Wang, X., Su, H., Xue, Y., Dominguez, M.G., Noguera, I., Torres, R., Macdonald, L.E., Stewart, A.F., DeChiara, T.M., and Yancopoulos, G.D. 2003. High-throughput engineering of the mouse genome coupled with high-resolution expression analysis. Nat. Biotechnol. 21:652-659. Wu, S., Ying, G., Wu, Q., and Capecchi, M.R. 2008. A protocol for constructing gene targeting vectors: Generating knockout mice for the cadherin family and beyond. Nat. Protoc. 3:1056-1076. Yagi, T., Ikawa, Y., Yoshida, K., Shigetani, Y., Takeda, N., Mabuchi, I., Yamamoto, T., and Aizawa, S. 1990. Homologous recombination at c-fyn locus of mouse embryonic stem cells with use of diphtheria toxin A-fragment gene in negative selection. Proc. Natl. Acad. Sci. U.S.A. 87:9918-9922.

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Mouse Strains and Genetic Nomenclature Jean-Louis Gu´enet1 and Fernando J. Benavides2 1

D´epartement de Biologie du D´eveloppement, Institut Pasteur, Paris, France The University of Texas M.D. Anderson Cancer Center, Science Park-Research Division, Smithville, Texas 2

ABSTRACT In this article we describe the main characteristics and peculiarities of the different strains and stocks of laboratory animals from the genetic point of view. We explain how they are produced and maintained as well as their advantages and disadvantages in the context of animal experiments. We also provide some guidance to make the best possible choice when establishing an experimental C 2011 by John Wiley & Sons, Inc. protocol. Curr. Protoc. Mouse Biol. 1:213-238  Keywords: inbred strains r congenic strains r recombinant inbred strains r recombinant congenic strains r outbred stocks

THE DIFFERENT CATEGORIES OF LABORATORY STRAINS AND STOCKS From the geneticist’s point of view, laboratory rodents belong to one of two categories. The first category consists of genetically uniform and highly standardized populations, where all animals have exactly the same genetic constitution. The second category consists of genetically heterogeneous populations segregating for a variety of alleles. Inbred strains and their derivatives belong to the first category. These are artificial populations analogous (but not identical) to cloned animals of the two sexes. They are different from human populations, but have the enormous advantage of being genetically uniform and expected to remain so, generation after generation, provided they are bred with an appropriate protocol. Inbred strains are numerous, and, altogether, they represent a wide sample of allelic forms at several loci. They also have derivatives such as congenic strains, recombinant inbred strains, recombinant congenic strains, and consomic strains that represent interesting tools for geneticists, as discussed in this article. In contrast to inbred strains, randombred and outbred stocks are genetically heterogeneous, and in this respect they are more similar to human populations. Randombred stocks, as the name indicates, are bred with no specific protocol: in other words, progenitors at generation G are mated randomly to produce generation G + 1. The genetic constitution of these stocks is generally unknown

and may change with time, sometimes rapidly, depending on the size of the breeding nucleus. Outbred stocks, on the other hand, are bred according to a specific protocol and, although they segregate for a variety of alleles at a number of loci, the frequency of these alleles in the population fluctuates within limits that are determined by the breeding system. The decision to use an inbred strain rather than a randombred or outbred stock in an experimental protocol depends on the biological question that wants to be addressed. The reasons for choosing one over the other will become obvious in the following sections, once the genetic constitution and main characteristics of the two categories are explained.

INBRED STRAINS AND THEIR DERIVATIVES Inbred Strains According to the definition of the International Committee on Standardized Nomenclature for Mice and the Rat Genome, “a strain can be regarded as inbred if it has been produced by mating systematically brother to sister for at least 20 consecutive generations, and individuals of the strain can be traced to a single ancestral pair at the twentieth or subsequent generation.” Once past the 20th generation of full brother-to-sister (full-sib) mating, the genome of each animal within the inbred strain will, at least theoretically, have no more than 2% of residual heterozygosity and members of the same strain can then be regarded as

Current Protocols in Mouse Biology 1: 213-238, March 2011 Published online March 2011 in Wiley Online Library (wileyonlinelibrary.com). DOI: 10.1002/9780470942390.mo100181 C 2011 John Wiley & Sons, Inc. Copyright 

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Figure 1 This drawing schematically represents the breeding system that is commonly used to produce an inbred strain: mating a male and a female from the same litter (brother × sister) in successive generations. Theoretical computation would indicate that exceptional matings between parent and offspring would not affect the progress towards homozygosity provided that the parent that is selected for mating is always the youngest. A male, for example, can be used for mating with one of its daughters but not with a female offspring born from this cross. Each generation of inbreeding is symbolized by the uppercase letter F, followed by the number of generations. When this number is not known, a question mark is often used; for example, F ? + 27 would indicate that the number of brother × sister matings was not known when the strain was acquired but 27 generations of unrelaxed inbreeding have been added since this time. B, brother; S, sister.

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genetically identical (Silver, 1995; Davisson, 1996; Fig. 1). In practice, most of the mouse and rat inbred strains that are commonly used in research nowadays have undergone several tens of generations of inbreeding, and some have even been bred exclusively with this system since the beginning of the last century, which means that they now have passed 300 generations, and sometimes more. This definition of an inbred strain calls for a few comments. As has already been mentioned, mice and rats of the same inbred strain are genetically identical or, as geneticists would say, they are isogenic. Because of strict inbreeding, they have also become homozygous at all loci of their genome that were segregating in the

founder ancestors (i.e., the ancestral breeding pair). After a few tens of generations, one of the alleles that had been segregating at a given locus has become fixed in the two parents, while the others (up to three) have been lost. This process of allele loss (or fixation) is easy to understand considering that, when by chance an allele that was present at generation F is not represented in at least one member of the breeding pair that is mated to produce the generation F + 1, it is then permanently lost. In other words, as inbreeding progresses, alleles are constantly lost and none are ever introduced (with the exception of rare de novo mutations). The sorting of the alleles at each generation depends mainly on chance; if the inbreeding protocol could

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be reset, with the same founder animals, it would certainly lead to a strain with a different genetic constitution after the same 20 generations. This means that an inbred strain represents a unique and fortuitous assortment of alleles. To get an idea of the genetic profile of an inbred strain, one could imagine a totally virtual scenario where an X-bearing spermatozoon fertilizes an oocyte. Then, suppose that the female pronucleus is removed before it merges with the male pronucleus, and finally, that the chromosomes that were brought into the oocyte by the spermatozoon are duplicated to form the nucleus of an embryo (one-cell stage). This new (n + n = 2n) and totally artificial embryo would be a female, with the two chromosomes of each pair absolutely identical. This is precisely what the genome of all members of an inbred strain looks like, with the exception, of course, of the sex chromosomes. During the process of inbreeding, the progression toward homozygosity is not linear. It is relatively fast during the first few generations, where a great number of genes become homozygous, then it slows down, and after 20 generations of unrelaxed inbreeding it is no more than about 1% to 2% of the loci that are still segregating. A mathematical series, based on Fibonacci’s numbers, is traditionally used to model the progress toward homozygosity while the number of sib matings increases; it is, however, only an approximation (Fig. 2).

Since complete homozygosity is virtually reached, at all loci, after a few tens of brother to sister matings, it may then come to mind that it is no longer necessary to use such a stringent breeding protocol to propagate an inbred strain. In fact, this would be a dangerous decision because mutations constantly occur and the interruption of inbreeding would allow an increasing number of new mutations (polymorphisms) to accumulate insidiously in the population. Even if the spontaneous mutation rate is very low (around 10−6 per locus per gamete, on average), the vast number of genes in a mammalian genome (around 25,000 protein-coding genes, according to recent estimates) would make this source of polymorphisms non-negligible. Accordingly, and for this reason mainly, inbreeding must never be relaxed. However, and as the International Committee on Standardized Nomenclature decided, other breeding schemes than brother × sister matings may occasionally be used, for example parent × offspring matings with the youngest of the parents being used only once (i.e., the offspring that is mated to its parent at generation Fn may be mated to one of its offspring at generation Fn + 1). As previously mentioned, the fact that all members of an inbred strain are isogenic is an enormous advantage because scientists working with the same inbred strain, but in different laboratories or at different time periods,

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can perform experiments where the fluctuations in the experimental results, by definition, will not be the consequence of possible differences in the genetic constitution of the animals. Being isogenic also provides the great advantage that one can define, in detail and comprehensively, the phenotypic characteristics of an inbred strain by gathering experimental data concerning this strain from several sources. For example, The Jackson Laboratory has developed a program to establish a collection of baseline phenotypic data on the most popular inbred strains of mice through a coordinated international effort. Information collected in this program (The Mouse Phenome Database; Paigen and Eppig, 2000) is freely available to the research community through the Internet (http://phenome.jax.org/). The development of this database, which is regularly updated, is possible only because inbred mice are isogenic (Fig. 3). Around 450 different mouse and 200 rat inbred strains are available worldwide, and a dozen among these strains, in both species, have become very popular. The genetic characteristics of inbred strains are considered artificial since their genetic constitution (isogenicity and homozygosity) has no natural equivalent in any species, including man. They are also artificial because we now know, from historical records confirmed by molecular data collected at the DNA level, that although the classical laboratory inbred strains of mice derive from a very limited number of ancestral progenitors, they do not derive from a single subspecies of the Mus genus but from at least two: Mus musculus domesticus and Mus musculus musculus (Bonhomme et al., 1987; Wade et al., 2002; Frazer et al., 2007). In this respect, the genome of inbred strains can be regarded as a patchwork of chromosomal segments stemming from these two subspecies in proportions that vary from one strain to another. This characteristic is unique to the laboratory mouse and does not exist in the laboratory rat, where all strains are derived from the same species, Rattus norvegicus. In the mouse, this is particularly interesting because it contributes to an in-

crease in genetic polymorphisms, making each strain quite different from the next. Studies on the genetic determinism of complex traits will benefit from this situation, as discussed later (Fig. 4). Being isogenic, mice of the same inbred strain are also histocompatible (also known as syngeneic). This means that they permanently accept tissue grafts from any mouse of the same strain (and sex). Mouse geneticists have extensively used this peculiarity since it allows studying the fate of cells with an immunological function in different contexts (cellular cooperation). It has also been extensively used for the serial transplantation of malignant cells. As previously mentioned, the main characteristics of the most commonly used inbred strains can be retrieved from the Internet. Taking a look at these descriptions is always of help when selecting a strain for an experimental protocol. Another important issue to consider is the phylogenic (or ancestral) relationships of the different strains, as recently established using molecular markers (mostly SNPs) (Beck et al., 2000; Frazer et al., 2007). This information is available both for mouse and rat inbred strains and is very valuable for studying the genetic determinism of phenotypic difference between strains. Indeed, selecting the less related parental strains when setting up a cross offers a greater chance to get a high resolution in the genetic analysis (Fig. 5). In the above discussion of the characteristics of mouse and rat inbred strains, it was mentioned that mutations were constantly occurring during inbreeding, generating new polymorphisms, and it was stated that inbreeding should not be relaxed to prevent an increase of these new polymorphisms. However, while a proportion of these new mutant alleles are effectively (and randomly) eliminated by inbreeding, another proportion may also become progressively fixed in the homozygous state, replacing the original allele: this is one aspect of what geneticists call genetic drift. It is a very slow process but it is totally unavoidable and

Figure 3 (figure appears on previous page) This table was captured from a window from the Mouse Phenome Database. It represents the complete set of data for blood cholesterol performed on both male and female mice of 43 different inbred strains in Dr. B. Paigen’s laboratory at the Jackson Laboratory. These data (in mg/deciliter) correspond to baseline data from mice aged 7 to 9 weeks. Checking these data before embarking on a research project related to cholesterol metabolism is definitely of great help (http://phenome. jax.org/db/qp?rtn=views/measplot&brieflook=9904). Arrowheads indicate exceptionally high or low values.

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Mus m. musculus

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Modern laboratory inbred strains - Mus laboratorius

Figure 4 Historical data, confirmed by sequence data, indicate that modern laboratory inbred strains derive from a small number of ancestors belonging to several different subspecies of the genus Mus. Today’s classical laboratory inbred strains should be regarded as recombinant strains derived (in unequal percentages) from three parental components: Mus musculus domesticus, Mus musculus musculus, and Mus musculus castaneus. For this reason it would probably be more appropriate to designate them as Mus “laboratorius”! This heterogeneous and unnatural genetic constitution is detectable at the genomic/sequence level by variations in the density of polymorphisms (in particular SNPs), with sharp edges, as represented on the picture by color differences for some chromosomal regions. The pattern of DNA polymorphisms distribution along the different chromosomal regions varies according to the strain and can be used for the purpose of mapping complex (QTL) traits.

contributes progressively to strain divergence when, for example, the same strain is propagated independently in different places. Genetic drift is a slow process because the spontaneous mutation rate is very low, and only one-fourth of the new mutant alleles become fixed. However, as mentioned, there are many genes in a mammalian genome and substrain divergence is a serious issue that should certainly not be underestimated. There are many examples in the literature where substrains, although stemming from the same original inbred strains, have acquired new and unique phenotypic characteristics as a consequence of

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genetic drift (Bulfield et al., 1984; Specht and Schoepfer, 2001; Stevens et al., 2007). Cryopreservation of embryos is an efficient way to minimize the effects of the genetic drift, which is often used in the world largest genetic repositories. Inbreeding of mice and rats was initiated almost exactly one century ago. Dr. Helen D. King is associated to the first inbred strain of rat, while Clarence C. Little, the founder and first Director of the famous Jackson Laboratory, and Leonell C. Strong are both associated with some of the most commonly used inbred strains of mice: C57BL and DBA

Figure 5 (figure appears on following page) (A) This picture represents a mouse family tree. The 102 inbred strains represented on this picture have been genotyped for a set of 1,638 informative SNP markers, evenly distributed over the whole genome (spaced on average <1.5 Mb). Applying a neighbor-joining method to the data, the authors constructed a mouse strain family tree that could be organized into seven groups. The length and angle of the branches have been optimized for printing and do not reflect the actual evolutionary distances between strains. This family tree is in good agreement with most other existing genealogies. Adapted from Petkov et al. (2004). (B) This picture represents a phylogenetic family tree of 93 rat inbred strains. It was developed with the same technique as above (heuristic search for maximum parsimony). Adapted from Mashimo et al. (2006).

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(Little), and C3H and CBA (Strong). For those readers interested in the historical aspects of inbred strains, a book titled “Origins of Inbred Mice” (Morse, 1978) is accessible on the Mouse Genome Informatics (MGI) Web site (http://www.informatics.jax.org/morsebook/). It is a rich and most interesting source of information, with a beautiful chapter by Dr. L. C. Strong himself! Most classical laboratory inbred strains are quite closely related because they are derived from a handful of progenitors: probably a single female that existed ∼200 years ago and whose mtDNA has been transmitted virtually unchanged to modern laboratory mice, with no more than two or three males of two different subspecies whose Y chromosome is still present in the same modern laboratory inbred strains (Ferris et al., 1982; Yonekawa et al., 1982; Bishop et al., 1985; Tucker et al., 1992). In most cases, this low level of polymorphism between strains is not a problem, and comparing two classical laboratory inbred strains sometimes makes the genetic analysis rather simple. In other instances, however, it is a more serious issue, and this is well exemplified in the observation that, unlike most wild mice, virtually all the classical inbred strains are susceptible to infection with flaviviruses, which impedes their use for investigating the genetics of resistance to the flaviviridae family. This phenotypic characteristic depends on a particular gene on Chromosome 5 (Oas1b) that is present (by chance?) in the same allelic form across all laboratory strains except one (Mashimo et al., 2002). To overcome the drawbacks associated with the (relative) lack of polymorphisms in the classical inbred strains, several scientists have recently developed novel inbred strains from progenitors belonging to the same Mus genus (Mus musculus musculus, Mus musculus molossinus, and Mus spretus, mainly) directly trapped in the wild state in diverse geographical locations (Bonhomme and Gu´enet, 1996). These novel strains have the drawback of not being as tame as most laboratory strains, but

they represent a very rich source of polymorphisms for laboratory investigation and have already been extensively used (Gu´enet and Bonhomme, 2003; Dejager et al., 2009). In most mammalian species, inbreeding is often associated with a decline in fitness and reproductive performances occurring after a few generations of sib matings. This is known as inbreeding depression and is thought to be a consequence of some deleterious recessive alleles becoming homozygous, or of some unfavorable allelic combination at different loci (Charlesworth and Willis, 2009). In mouse species, inbreeding depression is not common and is generally associated with the existence of the so-called t-haplotypes in the original breeders. t-haplotypes behave as recessive lethal alleles, which are transmitted by males in a non-Mendelian manner, and, accordingly, are quite common in wild populations of Mus musculus domesticus mice. On the contrary, mouse strains that are derived from closed populations are often partly inbred, and fullsib mating generally does not result in any depression in their vitality. Since the occurrence of a phase of inbreeding depression is unpredictable, new inbreeding projects should involve a sufficiently large number of lines to compensate for the possible extinction of a proportion of these lines during the first few generations (Fig. 6). Interestingly, very few of the inbred strains developed in our own laboratory from wild-trapped progenitors have been affected by inbreeding depression, irrespective of their phylogenetic origin (Mus m. musculus or Mus spretus). We believe that this is a consequence of the fact that the wild mice we trapped and used as progenitors were all from the same geographical location and probably shared several ancestors and accordingly were already moderately inbred. Nowadays, mouse inbred strains are commercially available from a number of vendors established in several countries worldwide, and it is generally not a problem to purchase top-quality animals even in large quantities.

Figure 6 (figure appears on previous page) This picture schematically represents the different breeding schemes that can be used for keeping an inbred strain with three cages. (A) The first schemes consist of mating one male and one female in each of the three cages, at each generation, with the breeders being brother (B) and sister (S) born in the same cage. This system cannot be recommended because in case of infertility or death one line will be lost, and in case of full success (unlikely!), one would end up with three loosely related substrains instead of one single strain. (B) This breeding scheme is the most reliable. It consists of selecting the progeny at generation F, which would allow making the largest number of pairs of breeders at generation F+1. It is very safe, but, unfortunately, it is not always applicable. (C) This system is an intermediate between A and B, and is the most popular in practice.

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These mice are often considered expensive, but it must be kept in mind that they are bred with high standards and are, in most instances, carefully monitored for their health status and genetic quality. The only problem is that mice or rats from a given strain, bred and commercialized by a given vendor, sometimes differ slightly from mice of the same strain bred elsewhere, and the scientific literature is full of examples of such variations (Stevens et al., 2007). This means that clear and precise references should always be provided in the scientific reports to avoid the publication of misleading results (Sundberg and Schofield, 2010). When explaining the progress toward full homozygosity during inbreeding, scientists often consider the genome as a little bag full of independent “molecular beads”: the genes. In reality, we know that the beads in question are linked and arranged on linear structures, the chromosomes, and the evolution towards homozygosity involves blocks or “chunks” of chromosomes rather than individual beads/genes. This explains why inbred strains carrying the same ancestral phenotypic marker share the same short segment of flanking DNA (haplotype) on both sides of the marker. For example, strains homozygous for the ancestral albino (Tyrc ) allele (A; AKR; BALB/c; and SJL) are all homozygous for a stretch of “ancestral” DNA flanking the albino mutation.

Interstrain F1 Hybrids

Mouse Strains and Genetic Nomenclature

Resulting from the cross of two inbred strains, F1 hybrids are heterozygous at all loci for which the parental strains have different alleles; however, they are genetically uniform (isogenic). Pairs of the same sex are strictly analogous to monozygotic twins. They are also histocompatible and permanently accept tissue transplantations from either parental strain, from their littermates, and from their offspring; however, the parental strains will not accept a graft from the F1 hybrids. They also exhibit the legendary hybrid vigor, the opposite of inbreeding depression, making them a material of choice in many experimental protocols (for example, DNA pronuclear microinjections). Considering all their many advantages, we could even say that F1 mice or rats would probably be the optimal choice in many experimental protocols. However, a major drawback is that, when intercrossed, their progeny (referred as F2) are genetically heterogeneous, since the alleles at all polymorphic loci start segregating. This is common, for example, in the protocols aimed at the production of genet-

ically engineered animals, where F1 hybrids are often used because of their high production of pre-implantation embryos or for the creation of strong chimeras. Interstrain hybrids can also be used to generate genetically heterogeneous populations. This is the case when, for example, F1 hybrids between strain A and strain B (abbreviated ABFl) are crossed with Fl hybrids between strain C and strain D (CDFl) to generate a 4-way heterogeneous stock. In this case the basic ingredients of such a genetically heterogeneous stock (i.e., the original inbred strains A, B, C, and D) are perfectly identified, and similar, but not identical stocks, can be produced, at will, when necessary. Genetically heterogeneous stocks with an even more complex structure (for example 8-way crosses stemming from eight different and unrelated inbred strains) have also been bred on a large scale for research in quantitative genetics (Jackson et al., 1999; Threadgill et al., 2002; Li et al., 2005). We will come back to this point when discussing the nonisogenic populations of laboratory animals, at the end of the present unit.

Co-Isogenic and Congenic Strains When a mutation occurs in the breeding nucleus of a highly inbred strain, and if we assume that this new mutation is fixed (probability = 0.25), then the inbred strain in question differs from the original strain at only one specific locus. If the new mutation is viable and does not impair the fertility, one can propagate the new strain by crossing brother and sister mutant mice or, much better, by mating, at each generation, a non-mutant mouse of the original inbred strain to an animal of the new mutant strain. These two strains are said to be co-isogenic strains or, sometimes, segregating inbred strains, and less frequently mutant strains. Co-isogenic strains are extremely useful for gene annotation because they allow comparing the phenotypes of two allelic forms of a given gene under optimal conditions (i.e., with no influence of the genetic background). A large number of such strains are stored worldwide, in particular, in the major genetic repositories. Some inbred strains, like the famous C57BL/6, have several co-isogenic “companion” strains segregating for a variety of allelic forms involved, for example, in the determinism of coat color. Other mutations with detrimental effects on the development or metabolism are also very interesting models because they can help in the analysis of the pathophysiology,

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providing both the experimental animal and its control. Using such strains it is possible, for example, to attempt phenotypic rescues by grafting normal cells into a co-isogenic mutant mouse as a preliminary study for the design and development of possible therapies. Co-isogenic strains, when developed in parallel to the background strain, may accumulate other genetic differences over time as a consequence of genetic drift, so, to minimize the effects of this drift, they must be periodically back-crossed to the parental strain, or be cryopreserved. Co-isogenic strains have two major drawbacks that are inherent to their origin and seriously limit their use: (1) they appear mainly as a consequence of a rare and fortuitous event (a mutation), and (2), although they can appear in any inbred strain, it is in general not the one that we would have been primarily interested in. For these two reasons, the use of co-isogenic strains is rather limited although, for example, mice of the C57BL/6Tyrc albino co-isogenic strain have become popular for the production of easily recognizable C57BL/6- +/+ ⇔ C57BL6-Tyrc /Tyrc chimeric mice from C75BL/6 ES cells. It is important to keep in mind that genetically engineered mice and rats can also be considered co-isogenic strains when the genetic modification is done in a way such that the targeted locus or transgene is the only difference from the wild-type animals. In the case of transgenic animals, this can be achieved by doing the pronuclear DNA microinjection (carrying the transgene) using embryos derived from an inbred strain (e.g., FVB/N). Transgenic lines must be developed independently from each founder animal (microinjected embryos) and are normally kept by backcrossing the transgenic carriers (heterozygous Tg/0) with wild-type animals from the inbred (background) strain and by selecting, at each generation, the new carriers (typically by PCR genotyping). One important difference between transgenic and co-isogenic strains is that, in most instances, the structure of the transgenic insertion can change with time in terms of copy number and DNA methylation, and sometimes even completely lose the foreign DNA, leaving behind a microchromosomal rearrangement (a deletion in general). In the case of targeted mutations (knock-outs and knock-ins), we can achieve co-isogenic strains by using ES cells and host blastocyst from the same inbred strain. For example, when C57BL/6-derived ES cells are

injected into albino C57BL/6 blastocysts, the chimeric mice are easily identified because their coats exhibit white and black patches. These chimeras can then be crossed with albino C57BL/6 mice to test for germline transmission (validated by the appearance of EScell derived black offspring; Schuster-Gossler et al., 2001). Congenic strains are an alternative to coisogenic strains with the advantage that any allele of the genome may be moved (technically “introgressed”) to any inbred background; the disadvantage is that the situation is not as pure from the genetic point of view as it is in the case of co-isogenics. Congenic strains are produced by crossing two strains: the first strain carries the gene (allele) or chromosome region of interest (i.e., spontaneous, induced, or targeted mutations, as well as transgenes), and is referred as the donor strain; the second strain is referred to as the recipient strain or the background strain. The F1 offspring generated by crossing the abovementioned two strains are back-crossed to the background strain, and the offspring that carry the allele of interest (i.e., the one originating from the donor strain) are crossed again to the background strain and so forth, typically for ten or more successive generations (Fig. 7). During this succession of back-crosses, the chromosomes of the background strain progressively replace those of the donor strain, except for the one that carries the allele of interest. For this particular chromosome, the segment containing the target allele is reduced in size only when a recombination event occurs that exchanges a piece of chromosome of the donor strain for the homologous segment of the background strain. Since the frequency of that sort of event depends on the size of the segment, one understands then that the chromosome carrying the target allele is gradually eroded on both sides, generation after generation, but in a nonlinear manner. The chromosomal segments that are flanking the selected locus have a tendency to remain associated to this locus, generation after generation, and this causes the basic difference between congenic and co-isogenic strains to materialize: while co-isogenic strains differ from the background strain strictly at a single locus, congenic strains differ by a short chromosomal segment flanking the target locus, with the size of this segment being progressively reduced during the successive back-cross generations. Since, on average, at each generation, an equivalent proportion of the background strain

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Mouse Strains and Genetic Nomenclature

replaces one half of the genome of the donor strain, the progress in genome substitution is given by the formula 1/2N where N is the number of back-cross generations. This means that, theoretically, after 10 back-cross generations only 1/210 (∼ 1/1,000) of the donor genome remains in the congenic strain. It is clear that this assumption is, again, purely statistical and the actual percentage of donor genome is subject to important variations at each generation. In addition, and as we already pointed out, this estimation stands only for the chromosomes that do not carry the allele of interest. In the latter case, the reduction in size is a much slower process. The use of polymorphic and easy-to-score DNA markers has allowed a much more rapid and rigorous process of congenic strains development: the so-called marker-assisted breeding, also referred to as the speed congenics methodology. The principle that underlies the speed congenics process is based on the fact that one can select the breeders, at each generation of back-crossing, based on the percent-

age of donor genome they have, by using either microsatellites or SNPs discriminating the two parental strains. Obviously, the mouse with the lowest percentage of donor DNA is the one to select as a breeder, at each generation, for setting the next back-cross (Fig. 8). Doing this, one can seriously reduce the number of generations necessary to reach full congenicity (for example, from N10 to N5), and the strain development time, approximately by half. At this point, it is important to note that, although a large number of molecular markers are necessary to perform an efficient and reliable genotyping during the first back-cross generations (in general 80/90 evenly distributed over the whole genetic map, for the N2 generation), this number decreases rapidly since, once a marker is typed “homozygous” for the allelic form of the background strain, it is no longer necessary to genotype the offspring of the future generations for this marker; it is permanently fixed (Fig. 9; Markel et al., 1997; Wakeland et al., 1997). As explained in Figure 10, the use of molecular markers is

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Figure 8 After each back-cross generation, an average of 50% of the genomic DNA of the donor strain is replaced by the equivalent proportion of genomic DNA of the background strain. However, selecting at each back-cross generation, the breeder with the lowest percentage of introgressed (donor) DNA greatly accelerates the establishment of a congenic strain. This picture represents the breeder (boxed) that was recognized as the most interesting (“best breeder”) after genotyping because of the lowest percentage of donor genome (shown in red). The mutation or region of interest is indicated by an arrowhead.

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Figure 9 Selecting the best breeders at each back-cross generation can save a lot of time when establishing a congenic strain. It is important to note that genotyping requires many polymorphic DNA markers only for the first back-cross progenies. Once a marker is characterized homozygous, it is no longer necessary to type it in the forthcoming generations. The y axis represents the number of mice with a percentage of homozygous loci lower or equal to the value indicated on the x axis. Adapted from Wakeland et al. (1997). Mouse Strains and Genetic Nomenclature

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D7Mi327

D7Mit25

D7Mit58

Targeted locus

D7Mit86

F1

N2

N4

N6

Figure 10 The genomic DNA flanking the target (introgressed) gene or transgene can be estimated very precisely with a battery of molecular markers, generation after generation. When, by chance, a cross-over reduces the flanking DNA on one side or the other, the mouse then becomes a privileged breeder. The genotyping of the flanking regions with the aim of selecting the “best breeder” can be integrated in the overall genotyping protocol. Briefly, it is recommended to select and type many more markers in the flanking regions than in other regions of the genome. The probability of finding a mouse with one crossing-over on each side of the flanking region is extremely low. For this reason it is recommended to proceed in two discrete steps, sequentially breeding mice with cross-overs in a flanking region of the targeted gene. If we consider a locus A, which is c Morgans distant from the selected (or targeted) locus T, on the same chromosome, the probability that there has been no recombination between the two loci A and T is e−c per generation and therefore e−nc after n generations (Johnson, 1981). After 10 generations of back-crossing, there is a 37% chance that no recombination occurred between loci A and T if they are distant 10 centimorgans (cM). This will rise to 61% if A and T are 5 cM apart and up to 90% if the two loci are 1 cM apart. If we consider that there are approximately 27,000 genes in the mouse genome and that the genetic map in this species spans ∼1,520 cM on average, this would mean that in 90% of the cases two congenic strains differ by ∼17 genes on both sides of the introgressed locus T. Of course, this is far from being negligible even if it should be weighted by the fact that no more than 20% of the genes are polymorphic between any two inbred strains.

Mouse Strains and Genetic Nomenclature

also useful for the selection of breeders with the smallest amount of “flanking” or “hitchhiking” DNA, to avoid the “flanking gene” concern (Wolfer et al., 2002; Chen et al., 2004). This requires the breeding of a large number of offspring, but these mice or rats can be genotyped at an early age and euthanized if considered unnecessary for future matings. It is increasingly recognized that the genetic background (i.e., all genomic sequences other than the gene of interest) can have profound influences on the phenotype of an animal model. It has been shown that mutations (spontaneous and induced), transgenes, and targeted alleles (knock-outs and knock-ins) that are “moved” onto a different background can show a change in phenotype (Threadgill et al., 1995; Linder, 2001). One of the first

cases involved the classical diabetes (Leprdb ) mutation that presented transient diabetes on a C57BL/6 background but overt diabetes on C57BLKS (Hummel et al., 1972). In order to stay away from confounding or unreliable experimental results, particularly with the increasing number of mouse and rat strains, attention to the genetic background is crucial. Everything described so far about how to establish a congenic strain corresponds to the classical strategy that can be applied in virtually any laboratory and for any species. In this strategy, the geneticist chooses the most “interesting” breeders for the intended purpose (gene introgression) and mates them with an inbred partner of the background strain, then nature does the rest. In this context, the length of pregnancy and the time to reach

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sexual maturity limits the progress towards full congenicity. However, one can substantially accelerate the production of congenic strains by combining the efforts of geneticists and embryologists. One can choose, for example, 3-week-old females as heterozygous (carrier) breeders, superovulate them, collect their oocytes (often called eggs), and perform in vitro fertilization with sperm from the background strain. The fertilized eggs (zygotes) can then be implanted in pseudo-pregnant females and, when these females deliver their progeny, one can proceed with another round of selection with molecular markers. With an efficient protocol, the time to implement a new backcross generation can be reduced to 8 weeks, and a new congenic strain can then be established in no more than 10 months (superspeed congenics). In this matter, Japanese scientists have established a new record by injecting round spermatid nuclei from immature males (only 17 days old) into mature oocytes in vitro. With this new technique called ROSI (for ROund Spermatid Injection), they developed a full-congenic strain (N3 mice genotyped with 86 DNA markers) in only 106 days, a real high-speed congenic strategy (Ogonuki et al., 2009). Congenic strains have been extensively used since the early days of mouse genetics and still are. They are particularly suited for the genetic analysis of phenotypes that are controlled by several genes, and it is precisely by developing such strains that George D. Snell, from The Jackson Laboratory, could elucidate the genetic determinism of histocompatibility (Snell, 1948). G. D. Snell was awarded the Nobel Prize in 1980 for this achievement. As we already mentioned, tissue transplantations performed between mice belonging to two different inbred strains are rejected, while the same transplantations performed between any two mice of the same inbred strains (and the same sex) are permanently accepted. The problem is that, in the case of tissue transplantations, the rejection (a phenotype) is controlled by several loci, each of them triggering independently the same phenotype. To clarify the situation, Snell bred a series of strains, with nearly the same C57BL/10 genetic background, but congenic for a single Mendelian unit inducing tissue incompatibility. To simplify the analysis of the phenotype and to save time, Snell injected tumor cells into mice segregating for the histocompatibility gene (all symbolized H) “selecting,” at each genera-

tion, only the mice that survived and accordingly were “resistant” to the (tumoral) tissue transplantation. He called these congenic mice congenic resistant (CR) and developed a very clever protocol to characterize each of these strains, thus avoiding duplications (CR strains congenic for the same gene just by chance). By doing this, Snell succeeded in making a comprehensive inventory of the H genes segregating among the laboratory strains. This strategy could be adapted with almost no change to the genetic analysis of any trait that is under polygenic control, for example, the resistance to infectious diseases. When a CR strain has been established, there is still a lot of work to do to finally characterize the gene that is involved in the phenotype, given that the chromosomal fragment is sometimes quite large, but CR strains are certainly of great help in these investigations. A few more comments will help us to complete the description of the congenic strains. The first refers to the fact that a pair of congenic strains can be perfectly established even if the donor strain is not inbred. For example, if a mouse is identified with an interesting characteristic segregating in a non-inbred population, it is always possible to derive one or more congenic strains for this trait following the same protocol described above. Another interesting possibility is to develop reciprocal congenic strains by introgressing a specific locus of strain A into the background strain B and, reciprocally, the homologous locus of strain B in the background strain A. At the end of the experiment, one has a total of four strains: the two parental inbred strains A and B on the one hand, and the reciprocal congenic strains AB and BA on the other. Then, one can compare the F1 between strain A and the congenic strain BA with the reciprocal F1 hybrid between strain B and the congenic strain AB (Fig. 11). That sort of experiment, making use of F1, has the advantage of eliminating the side effects of some possible epistatic interaction with the genetic background and is likely to provide more reliable answers. Finally, a comment is warranted on the use of congenic strains as tools for the analysis of quantitative (or complex) traits. When we discussed the experiments by Snell about the genetic analysis of histocompatibility, we mentioned that the derivation of CR strains made the individual identification of several H genes possible. Of course, this identification exclusively concerns the genes that are in a

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Reciprocal congenic strains

Reciprocal congenic F1

Figure 11 Reciprocal congenics allow making comparisons with a high degree of standardization because epistatic interactions may be detected by this procedure. For example, if a given allele in the background of a congenic strain interferes with the expression of the target gene, this might be detected when comparing the two reciprocal congenic strains but would presumably be corrected when comparing the two congenic F1s.

different allelic form in the congenic partners; those that are non-polymorphic remain undetected. This may appear a truism but, keeping in mind that the classical inbred strains of laboratory mice were all derived from a small pool of ancestral progenitors, it is clear that the experiments by Snell made possible the discovery of only a small proportion of all the H genes of the mouse species. Many other loci remained undetected and it is likely that the derivation of new CR strains from wild mouse specimens would certainly be very rewarding. This comment applies, of course, to all situations where many genes (and many alleles) are involved in the determinism of a complex or quantitative trait.

Consomic Strains

Mouse Strains and Genetic Nomenclature

Consomic strains also (improperly) designated chromosome substitution strains (CSS) are a variation of the congenic strains concept in which the introgressed DNA is a complete chromosome, rather than a piece of chromosome flanking a given gene (Fig. 12; Nadeau et al., 2000). These strains have been very useful for the rapid mapping of phenotypic traits to a single chromosome. They are also useful for the detection of chromosomal regions (the so-called Quantitative Traits Locus; QTL) having an influence in the determinism of a particular phenotype (for example the resistance or susceptibility to carcinogenesis).

Only a few sets of consomic mouse and rat strains are available, but it is likely that other sets will be developed in the near future to accompany the development of investigations in multifactorial inheritance (Gregorov´a et al., 2008; Mattson et al., 2008). Using a marker-assisted protocol, consomic strains appear (theoretically) easy to produce. However, one must keep in mind that tiny pieces of chromosomes of the donor strain might escape the marker-assisted selection process if, by chance, they are not identified by a marker. There is also no guarantee that the telomeric region is transferred intact since there is, in most instances, no distal marker to check this. Finally, we must say here that our attempt to develop a set of inter-specific consomic mouse strains using SEG/Pas (donor strain, Mus spretus) and C57BL/6 (background strain) was mostly unsuccessful. We found that, although we could breed most of the “heterozygous” consomics (with a complete chromosome of Mus spretus being introgressed in an otherwise C57BL/6 background), we could not or very rarely breed “homozygous” for a complete chromosome of SEG origin. We suspect that this was the consequence of deleterious epistatic interactions between genes (or alleles) that have been separated by evolution for a long time (over 1.5 million years), but alternative explanations are also possible.

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Background strain A

Strain A-Chr2B

Strain A-Chr4B

Strain A-Chr3B

Strain A-Chr1B

Donor strain B

Figure 12 This picture represents a set of four consomic strains flanked by the parental strains A and B. In each strain, a complete chromosome pair has been replaced by the homologous chromosome of the other inbred strain via a series of marker-assisted back-crosses. A complete panel of consomic strains consists of 21 strains, each derived from the same donor and host but having a different chromosome (Chr 1-19, X or Y) of the host replaced by its counterpart from the donor. A reciprocal panel can be produced by inverting the donor and host strains, respectively.

Recombinant Inbred Strains and Recombinant Congenic Strains Recombinant inbred strains (RIS) are developed by first crossing two parental inbred strains to generate hybrid F1s and then intercrossing the F1 to generate F2s. Randomly chosen F2 animals are then brothersister mated for twenty or more generations to develop a group of related inbred strains (Bailey, 1971). RIS go by sets (also referred to as panels): a collection of RIS derived from the same parental strains. For example, the C57BL/6 × DBA/2 (BXD) is, at the moment, the largest mouse RI panel with 77 strains. These are true inbred strains, meaning that they are homozygous at all loci but have the additional characteristic that each RIS has a unique fixed combination of the parental alleles in a 50%:50% proportion (on average). For example, each strain of the set of 33 AXBBXA strains, derived from the initial cross of a C57BL/6 mouse with a A/J mouse, carries either the B6 allele or the A allele at each locus of its genome, and by typing all of these allelic forms, one can establish a strain distribution pattern (SDP) for each of the strains, which lists the collection of alleles inherited from either the parental strain A or the parental

strain B6 (Fig. 13). Of course this SDP is fixed forever in each strain, and new data are constantly added to it, allowing correlations to be made between genotypes and phenotypes simply by scanning, generally with the help of a simple computer program, the co-segregation of new phenotype (or genotype) with the existing SDP. RIS have proven very helpful when used for gene mapping, in particular for the rapid regional assignment of microsatellites on a given chromosome, when these markers were cloned by the thousands for the establishment of high-density genetic maps. They have also been used for the mapping of chromosomal regions (QTLs) involved in the genetic determinism of some behavioral characteristics (taster/non-taster for a chemical compound, alcohol intake, etc.) or of some immunological responses, and they will very likely still be of help in many other experiments where the phenotype is measured on a group of animals rather than on individuals (Zou et al., 2005). However, since most of these RIS are all derived from the classical laboratory inbred strains, and given that the number of strains in the different sets is relatively small, QTL detection and their resolution remain limited.

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Parental strain A

R.I. strain AXB1

R.I. strain AXB3

R.I. strain AXB8

R.I. strain AXB12

Parental strain B

Figure 13 This diagram represents a set of four recombinant inbred (R.I.) strains flanked by the parental strains A and B. Individual RI strains have a unique combination of loci derived by recombination of the alleles present in the original parental strains. Since RI strains are inbred and each strain has a unique genotype, RI strains have a number of advantages over F2 or back-cross mouse populations as tools for mapping genes or quantitative trait loci (QTL).

Mouse Strains and Genetic Nomenclature

A new kind of recombinant inbred panel, which is a variation of the recombinant inbred strain concept, is now being developed and will undoubtedly represent a true breakthrough for the future of quantitative genetics in mice: it is known as the Collaborative Cross (Churchill et al., 2004; Chesler et al., 2008). The Collaborative Cross is a randomized cross of eight inbred mouse strains that have been carefully selected by a panel of mouse geneticists (The Complex Trait Consortium) after extensive discussions. These strains are A/J, C57BL/6J, 129S1/SvImJ, NOD/LtJ, NZO, CAST/Ei, PWK/Ph, and WSB/Ei, with the last three strains being recently derived from wild progenitors of Mus musculus subspecies. The eight strains are first crossed pairwise, to make all (8 × 7 = 56) possible G1 parents, then all 8 genomes are brought together in a series of F1, and the offspring of this cross are inbred for several generations (Fig. 14). Several hundreds of new inbred strains, ideally up to 1,000 (recombinant for variable proportions of the original eight parental strains), will then be available in a few years (expected in 2012/2013). These strains will be able to detect biologically relevant correlations among thousands of measured traits, and the 1,000 strains, considered together, will represent 135,000 recombination events, which is an enormous and

unprecedented power of resolution. Such a panel would indeed represent a valuable community resource; the only points that are still to be solved are those related to the facilities to host all these strains, their distribution in the community, and the funding of the project . . . as always! Recombinant congenic strains (RCS) are very similar to RIS in their genomic structure except that the proportion of the parental alleles in a given strain is not 50%:50% but 75%:25% or 87.5%:12.5%, depending on the set (Demant and Hart, 1986). This is achieved by inbreeding mice of the first or second back-cross generation to one of the parental inbred strains (the background strain). RCS are helpful for identifying genes associated with polygenic inheritance, especially when the number of genes is quite high. RCS with a small percentage of introgressed genome in a background strain have a greater power of resolution, and their use increases the likelihood of having zero or only one single locus governing the studied phenotype (QTL) being isolated in a given RC strain. For example, RCS have been very helpful for unraveling the genetic determinism of colon cancer in the mouse (Demant et al., 2003). Interspecific recombinant congenic strains (IRCS) have also been developed from the parental

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A A

B

C

A

B

C

The Collaborative Cross D E

F

G

H

F

G

H

G0

G1

G2

G3

G4

G

20

B

The Collaborative Cross D

E

Figure 14 (A) The Collaborative Cross is a randomized cross of eight unrelated mouse inbred strains designed by members of the Complex Trait Consortium. The lines are first crossed pairwise to make all 56 possible G1 parents. A set of possible 4-way crosses is performed, keeping Y-chromosome and mitochondrial balance. Finally, all 8 genomes are brought together in G2:F1 and the offspring of this cross are inbred. The Collaborative Cross is a community resource that was initially designed for the purpose of mapping complex traits. (B) The initial previsions were to breed around 1000 inbred strains where all the alleles of the initial inbred strains would be associated in a wide and unique variety of combinations. Only one strain is represented in this illustration; other strains would be similar but with a different pattern of parental strain distribution. Mouse Strains and Genetic Nomenclature

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strain C57BL/6JPas and SEG/Pas (Mus spretus; Burgio et al., 2007). This set of strains has proven particularly useful for the analysis of the genetic determinism of some anatomical traits (Burgio et al., 2009).

OUTBRED AND RANDOMBRED STOCKS

Mouse Strains and Genetic Nomenclature

Outbred and randombred stocks are populations of laboratory animals radically different from those considered above in the sense that they are genetically heterogeneous, or heterogenic as we may say to keep the same sort of terminology. According to the official definition, outbred mouse or rat stocks are “closed populations (for at least four generations) of genetically variable animals that are bred to maintain maximum heterozygosity.” Compared with inbred strains, F1 hybrids, or congenic strains, the genetic constitution of a given animal, taken randomly from an outbred stock, is not known a priori and must be defined when necessary. Outbred mice or rats represent the bulk of laboratory animals sold by commercial vendors for the purpose of experimentation. These animals are usually bred according to a system that minimizes (or, more exactly, reduces) inbreeding, and accordingly contributes to the maintenance of a certain amount of heterozygosity in the population (Hartl, 2001). A classical breeding scheme for these populations would consist, for example, of the mating in room C and D of n males originating from room A with the equivalent number of females taken from room B, with n being as great as possible. For the production of the next generation (G+1), the breeding scheme would be similar with n males from room C being mated with n females of room D, and so on. Doing this, generation after generation, the polymorphic alleles that were segregating in the population at generation G have the greatest chance to be still represented at generation G+1 in roughly the same proportion, and the greater the samples of breeders used for the production of G+1, the smaller the variations in frequency at each generation (Poiley, 1960). The degree of genetic heterogeneity in outbred colonies depends greatly on their history. It can be very low, for example as a consequence of genetic drift (or bottleneck effect), when the pool of breeders has been accidentally or intentionally reduced to a few individuals (this is common when a new breeding facility is created and a small group of breeders is imported). On the contrary, the genetic heterogeneity can be much higher when the stock

has been recently outcrossed. Some commercial breeders probably monitor the polymorphisms segregating in their stocks with DNA markers, but the methodology they use and the results they get are not always made public. Genetically heterogeneous, outbred, and randombred stocks have a greater fertility index than inbred strains, and accordingly they are sold at a much cheaper price per unit. Because they are heterogeneous populations, like human populations, outbred mice are often considered the most appropriate category of laboratory animals to use in genetics, toxicology, and pharmacology research. However, several geneticists have disputed this point of view and it has even been considered that, in many studies, outbred mice were used inappropriately, wasting animals’ lives and resources on suboptimal experiments (Chia et al., 2005; Festing, 2010). In fact, any outbred stock can be replaced by a “synthetic” population obtained by intercrossing classical inbred strains. As we already said, crossing two inbred strains to produce an F1 progeny and then crossing two independent F1 generates a 4-way polymorphic population. This population is heterogenic, in the sense that individuals are genetically different, and in addition the population often carries a greater number of allelic forms, which is generally considered an advantage compared to a classical outbred population. Recently, however, researchers have considered that outbred stocks might be useful to refine the identification of QTLs because these heterogeneous stocks accumulate many breakpoints over time that split their chromosomes into “fine-grained mosaics,” facilitating the high-resolution mapping of complex traits (Mott et al., 2000; Flint et al., 2005; Yalcin et al., 2010). This advantage of the outbred stocks will probably not persist, given the advent of strains derived from the Collaborative Cross. Finally, randombred stocks are of very limited interest for geneticists. These stocks are bred with no specific rules, paying almost no attention to the genetic diversity in the population. Since they are in general of relatively small size, they drift rapidly towards a moderately inbred but still undefined population.

NOMENCLATURE RULES FOR MOUSE AND RAT STRAINS Nomenclature rules represent a very important issue that must be considered with care, because they are no less than the code used for the correct designation of laboratory

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animal strains and stocks. These rules were established several years ago by the International Committee on Standardized Genetic Nomenclature for Mice and Rats and are constantly updated to fit with the progress in the genetics of these two species. They are extensively described—with examples—on the Mouse Genome Informatics (MGI) Web page (http://www.informatics.jax.org/mgihome/ nomen/strains.shtml) as “Guidelines for Nomenclature of Mouse and Rat Strains (revision September 2010.)” For this reason, we will only address the essentials of these rules and make a few comments. An inbred strain should be designated by a unique brief symbol made up of uppercase, roman letters, or a combination of letters and digits beginning with a letter (e.g., C57BL). Some pre-existing strains, like the group of 129 strains, do not follow this convention. Substrains are given the symbol of the original strain they are derived from, followed by a forward slash and a substrain designation. This substrain designation is, in most instances, the code of the individual or laboratory originating the strain. For example, A/He is the substrain of the A inbred strain developed by Dr. W. Heston. Similarly, 129S2/SvPasCrl is the symbol of the 129 substrain that was originally developed by Dr. L.C. Stevens, from The Jackson Laboratory, then introduced to the Pasteur Institute of Paris, and finally transferred for breeding to Charles River Laboratories in November 1996. Congenic strains are designated by a symbol consisting of three parts. The full or abbreviated symbol of the recipient strain is separated by a period from an abbreviated symbol of the donor strain, this being the strain in which the allele or mutation originated, which may or may not be its immediate source in constructing the congenic strain. In cases where the chromosome from which the mutation arose is unknown (e.g., the donor is not inbred), the symbol Cg is used to denote congenic. The use of the donor strain symbol, or Cg, is essential to distinguish congenic from coisogenic strains. A hyphen then separates the strain name from the symbol (in italics) of the differential allele(s) introgressed from the donor strain. For example, NOD.CB17Prkdcscid is a symbol for a strain congenic for the Scid mutation (Prkdcscid ), initially discovered in the CB17 strain, and introgressed into a NOD background. For outbred stocks, the common strain root is preceded by the Laboratory Code of the

institution holding the stock. For example, Tac:ICR is the ICR outbred stock maintained by Taconic Farms, Inc. Mouse and rat gene symbols are italicized and begin with an uppercase letter followed by all lowercase letters, except for recessive mutations, which begin with a lowercase letter. Human gene symbols are also italicized but are written in all uppercase letters. Protein symbols for mouse and rat follow that of human and are all uppercase letters. Table 1 provides various examples of different genetic nomenclature. In addition to this, the Jackson Laboratory supporting Web page provides a useful tutorial on basic mouse nomenclature rules and guidelines describing what is in the name of a mouse strain (http://jaxmice.jax.org/support/nomenclature/ tutorial.html). Another important point is that many scientific journals require the use of standardized nomenclature. A complete list of these journals can be found on the MGI Web site, Journals Enforcing Standard Nomenclature (http:// www.Informatics.jax.org/mgihome/nomen/ journals.shtml). In this context, the use of strain designations C3H, C57, Balb/c in place of C3H/HeNHsd, C57BL/6NCrl, or BABL/cByJ, would be flagged for corrections. As we already mentioned, when discussing the causes and mechanisms of genetic drift, some substrains are homozygous for point mutations, small deletions, or insertions of recent origin. Many of these mutations can be considered “quiet” in the sense that they do not produce any obvious phenotype (Stevens et al., 2007). However, in some instances, the phenotype of these mutations is expressed only in a specific context, and in the case where the substrain designation is not rigorously applied, the published results may be misleading. For example, mice of the C57BL/6JOlaHsd substrain have a phenotype comparable to all other C57BL/6 substrains, but they display complete resistance to the neurotoxic effects of 1-methyl-4-phenyl-1,2,3,6tetrahydropyridine (MPTP) because they are homozygous for some disruptions in the gene encoding synuclein alpha (Snca) (Specht and Schoepfer, 2001). It is then clear that if this particular substrain is used for research on Parkinson disease, a genetic disease involving synuclein activity, the results collected may be different from those collected using another C57BL/6 substrain. A similar comment applies to the substrain C3H/HeJ, which is homozygous for a mutation inactivating the Tolllike receptor 4 gene (Tlr4Lps-d ), making mice

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Table 1 Examples of Nomenclature Symbols in Mouse and Rat

Symbol

Definition

Gene locus Myo5a

Gene symbol for MyosinVA - Chr 9

Tyr

Gene symbol for Tyrosinase - Chr 7

Hexa

Hexosaminidase A - Chr 9

Col1a1

Collagen, type I, alpha 1- Chr 11

Lrp4

Low density lipoprotein receptor-related protein 4 - Chr 2

Mutations Tyrc Tyr

albinism - formerly c

c-Btlr

Tyrc-19J

albinism - a chemically (ENU) induced allele in Dr. Bruce Beutler’s laboratory albinism - a spontaneous reoccurrence of an albino allele at Jackson Laboratory (ranked N◦ 19)

Knock-out Hexatm1Grv tm1Bst

Col1a1

hexosaminidase A; targeted mutation 1, Roy A Gravel collagen, type I, alpha 1; targeted mutation 1, Paul Bornstein

Gene trapped HexaGt(AF0619)Wtsi

hexosaminidase A; gene trap AF0619, Wellcome Trust Sanger Institute

Transgene Tg(Col1a1-cre)1Kry

transgene insertion 1, Gerard Karsenty - Location unknown

Other mutations

Mouse Strains and Genetic Nomenclature

Fmn1ld-Is(17;In2)1Gso

Formin 1; limb deformity insertion 1 - W. Generoso Chromosomal transposition with inversion - complex

Lrp4dan

Low density lipoprotein receptor-related protein 4; digitation anormale - Chr 2

Col1a1tm2(tetO-Pou5f1)Jae

collagen, type I, alpha 1; targeted mutation 2, Rudolf Jaenisch knock-in

of this substrain hypo-responsive to bacterial lipopolysaccharide and more susceptible to infection by Gram-negative bacteria (Rosenstreich and Glode, 1975; Poltorak et al., 1998). These two examples, selected among the many others already published, emphasize the absolute necessity to comply with the nomenclature rules in the scientific publications (Sundberg and Schofield, 2010). This is especially true if we consider that mutations of the kind that we have described are constantly discovered, introducing subtle differences in the genotype/phenotypes of these highly standardized strains. It is also very important to emphasize that in no way may a substrain be considered “better” than another based on the sort of peculiarities that we have described; in fact all substrains are interesting and have the same

basic qualities (isogenicity and homozygosity). The only point that must be mentioned is that this has to be very precisely defined in the publications. Assigning official symbols prior to publication of articles prevents erroneous symbols from being propagated throughout the literature. To assist authors with their nomenclature concerns, the MGI nomenclature group can be contacted via e-mail (nomen@ informatics.jax.org) or via the Submission/ Registration forms located on the MGI Nomenclature Home Page.

CONCLUSIONS In writing this article, our intention has been to help scientists, technicians, and students, especially the newcomers, in selecting the best

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kind of laboratory rodents for their experiments. For this reason, we have provided a detailed description of the genetic makeup of all the strains and stocks that are or will become shortly available. Time has come now to provide an answer to the highly pragmatic question: which strain or stock is the best choice for my study? By and large, one can say that inbred strains should be considered first, in most instances, because animals of that kind are highly standardized populations and commercially available almost everywhere. They are also interesting because a very large amount of information of all kinds has been accumulated over the years and stored in public databases, making the choice somewhat easier. Another interesting aspect is that one can generate F1 animals by crossing two parental strains, and get isogenic populations, which are heterozygous at many loci and accordingly exhibit hybrid vigor. By using several different inbred strains of independent phylogenetic origin, one can “synthesize” genetically heterogeneous stocks (4-way or 8-way crosses), with many different alleles segregating in the population. Since the parental strains are known, such stocks can be produced at will and used in virtually any laboratory. Congenic strains can now be bred upon request, with very efficient methodologies. They are indicated under all circumstances where the effect of a given allele (or transgene) on the phenotype is to be studied per se, with no interference from the genetic background. Hybrid F1s are always interesting when the experiment consists of a comparison between groups of treated mice with untreated controls. However, in this case, it may be wise to test a few mice belonging to several F1 hybrid strains before making the experiment. F1 mice, in general, exhibit relatively homogeneous responses to biological assays, which allow more sensitive statistical analysis. The numerous inbred strains that will be derived from the Collaborative Cross certainly have a promising future, especially when their genome is entirely sequenced because this would greatly speed the identification of small genomic regions with an impact on specific phenotypes. Outbred mice are cheaper than all other types of mice, and this aspect certainly contributes to making them popular, at least to some extent. They can be used in many experiments where the genetic component is not important (vasectomized males for inducing

psevdopregnancy in female mice, recipient females for embryo transfer, etc.), and they can also be used to breed strong and vigorous F1 mice as recipients for transplantations (for example, BALB/c × outbred stock would accept most hybridomas). However, a statement concerning outbred stocks that appears to be controversial nowadays is that they are a better choice for many experiments because they are genetically heterogeneous, like humans. An excellent review by Dr. Michael Festing is accessible via the Internet (http://www.isogenic. info/html/animal models in research.html), and we strongly advise all scientists performing biological assays or other types of experiments to read Dr. Festing’s comments. Reading his recommendations will help to make the best possible choice of experimental animals and thus the best experimental design.

WARNING! In a number of countries, the use of animals for experimental purposes is strictly regulated by laws, and can be performed only by persons with special permission or under their responsibility.

LITERATURE CITED Bailey, D.W. 1971. Recombinant inbred strains, an aid to finding identity, linkage, and function of histocompatibility and other genes. Transplantation 11:325-327. Beck, J.A., Lloyd, S., Hafezparast, M., LennonPierce, M., Eppig, J.T., Festing, M.F., and Fisher, E.M. 2000. Genealogies of mouse inbred strains. Nat. Genet. 24:23-25. Bishop, C.E., Boursot, P., Baron, B., Bonhomme, F., and Hatat, D. 1985. Most classical Mus musculus domesticus laboratory mouse strains carry a Mus musculus musculus Y chromosome. Nature 315:70-72. Bonhomme, F. and Gu´enet, J.-L. 1996. The laboratory mouse and its wild relatives. In Genetic Variants and Strains of the Laboratory Mouse (M.F. Lyon, S. Rastan. and S.D.M. Brown, eds.) pp. 1577-1596. Oxford University Press, Oxford. Bonhomme, F., Gu´enet, J.-L., Dod, B., Moriwaki, K., and Bulfield, G. 1987. The polyphyletic origin of laboratory inbred mice and their rate of evolution. Biol. J. Linnean Soc. 30:51-58. Bulfield, G., Siller, W.G., Wight, P.A., and Moore, K.J. 1984. X chromosome-linked muscular dystrophy (mdx) in the mouse. Proc. Natl. Acad. Sci. U.S.A. 81:1189-1192. Burgio, G., Szatanik, M., Gu´enet, J.-L., Arnau, M.R., Panthier, J.J., and Montagutelli, X. 2007. Interspecific recombinant congenic strains between C57BL/6 and mice of the Mus spretus species: A powerful tool to dissect genetic control of complex traits. Genetics 177:2321-2333.

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Burgio, G., Baylac, M., Heyer, E., and Montagutelli, X. 2009. Genetic analysis of skull shape variation and morphological integration in the mouse using interspecific recombinant congenic strains between C57BL/6 and mice of the Mus spretus species. Evolution 63:2668-2686. Charlesworth, D. and Willis, J.H. 2009. The genetics of inbreeding depression. Nat. Rev. Genet. 10:783-796. Chen, S., Kadomatsu, K., Kondo, M., Toyama, Y., Toshimori, K., Ueno, S., Miyake, Y., and Muramatsu, T. 2004. Effects of flanking genes on the phenotypes of mice deficient in basigin/CD147. Biochem. Biophys. Res. Commun. 324:147-153. Chesler, E.J., Miller, D.R., Branstetter, L.R., Galloway, L.D., Jackson, B.L., Philip, V.M., Voy, B.H., Culiat, C.T., Threadgill, D.W., Williams, R.W., Churchill, G.A., Johnson, D.K., and Manly, K.F. 2008. The Collaborative Cross at Oak Ridge National Laboratory: Developing a powerful resource for system genetics. Mamm. Genome. 19:382-389. Chia, R., Achilli, F., Festing, M.F., and Fisher E.M.C. 2005. The origins and uses of mouse outbred stocks. Nat. Genet. 37:1181-1186. Churchill, G.A., Airey, D.C., Allayee, H., Angel, J.M., Attie, A.D., Beatty, J., Beavis, W.D., Belknap, J.K., Bennett, B., Berrettini, W., Bleich, A., Bogue, M., Broman, K.W., Buck, K.J., Buckler, E., Burmeister, M., Chesler, E.J., Cheverud, J.M., Clapcote, S., Cook, M.N., Cox, R.D., Crabbe, J.C., Crusio, W.E., Darvasi, A., Deschepper, C.F., Doerge, R.W., Farber, C.R., Forejt, J., Gaile, D., Garlow, S.J., Geiger, H., Gershenfeld, H., Gordon, T., Gu, J., Gu, W., de Haan, G., Hayes, N.L., Heller, C., Himmelbauer, H., Hitzemann, R., Hunter, K., Hsu, H.C., Iraqi, F.A., Ivandic, B., Jacob, H.J., Jansen, R.C., Jepsen, K.J., Johnson, D.K., Johnson, T.E., Kempermann, G., Kendziorski, C., Kotb, M., Kooy, R.F., Llamas, B., Lammert, F., Lassalle, J.M., Lowenstein, P.R., Lu, L., Lusis, A., Manly, K.F., Marcucio, R., Matthews, D., Medrano, J.F., Miller, D.R., Mittleman, G., Mock, B.A., Mogil, J.S., Montagutelli, X., Morahan, G., Morris, D.G., Mott, R., Nadeau, J.H., Nagase, H., Nowakowski, R.S., O’Hara, B.F., Osadchuk, A.V., Page, G.P., Paigen, B., Paigen, K., Palmer, A.A., Pan, H.J., Peltonen-Palotie, L., Peirce, J., Pomp, D., Pravenec, M., Prows, D.R., Qi, Z., Reeves, R.H., Roder, J., Rosen, G.D., Schadt, E.E., Schalkwyk, L.C., Seltzer, Z., Shimomura, K., Shou, S., Sillanpaa, M.J., Siracusa, L.D., Snoeck, H.W., Spearow, J.L., Svenson, K., Tarantino, L.M., Threadgill, D., Toth, L.A., Valdar, W., de Villena, F.P., Warden, C., Whatley, S., Williams, R.W., Wiltshire, T., Yi, N., Zhang, D., Zhang, M., Zou, F., and the Complex Trait Consortium. 2004. The Collaborative Cross: A community resource for the genetic analysis of complex traits. Nat. Genet. 36:11331137. Mouse Strains and Genetic Nomenclature

Davisson, M.T. 1996. Rules for nomenclature of inbred strains In Genetic Variants and Strains of the Laboratory Mouse (M.F. Lyon, S. Rastan,

and S.D.M. Brown, eds.) pp. 1532-1536. Oxford University Press, Oxford. Dejager, L., Libert, C., and Montagutelli, X. 2009. Thirty years of Mus spretus: A promising future. Trends Genet. 25:234-241. Demant, P. 2003. Cancer susceptibility in the mouse: Genetics, biology and implications for human cancer. Nat. Rev. Genet. 4:721-734. Demant, P. and Hart, A.A.M. 1986. Recombinant congenic strains: A new tool for analyzing genetic traits determined by more than one gene. Immunogenetics 24:416-422. Ferris, S.D., Sage, R.D., and Wilson, A.C. 1982. Evidence from mtDNA sequences that common laboratory strains of inbred mice are descended from a single female. Nature 14:163-165. Festing, M.F. 2010. Inbred strains should replace outbred stocks in toxicology, safety testing, and drug development. Toxicol. Pathol. 38:681690. Flint, J., Valdar, W., Shifman, S., and Mott, R. 2005. Strategies for mapping and cloning quantitative trait genes in rodents. Nat. Rev. Genet. 4:271286. Frazer, K.A., Eskin, E., Kang, H.M., Bogue, M.A., Hinds, D.A., Beilharz, E.J., Gupta, R.V., Montgomery, J., Morenzoni, M.M., Nilsen, G.B., Pethiyagoda, C.L., Stuve, L.L., Johnson, F.M., Daly, M.J., Wade, C.M., and Cox, D.R. 2007. A sequence-based variation map of 8.27 million SNPs in inbred mouse strains. Nature 448:1050-1053. Gregorov´a, S., Divina, P., Storchova, R., Trachtulec, Z., Fotopulosova, V., Svenson, K.L., Donahue, L.R., Paigen, B., and Forejt, J. 2008. Mouse consomic strains: Exploiting genetic divergence between Mus m. musculus and Mus m. domesticus subspecies. Genome Res. 18:509-515. Gu´enet, J.-L. and Bonhomme, F. 2003. Wild mice: An ever-increasing contribution to a popular mammalian model. Trends Genet. 19:24-31. Hartl, D.L. 2001. Genetic management of outbred laboratory rodent populations. Charles River Genetic Literature. http://www.criver. com/sitecollectiondocuments/rm gt r genetic management outbred rodent.pdf Hummel, K.P., Coleman, D.L., and Lane, P.W. 1972. The influence of genetic background on expression of mutations at the diabetes locus in the mouse. Biochem. Genet. 7:1-13. Jackson, A.U., Fornes, A., Galecki, A., Miller, R.A., and Burke, D.T. 1999. Multiple-trait quantitative trait loci analysis using a large mouse sibship. Genetics 151:785-795. Johnson, L.L. 1981. At how many histocompatibility loci do congenic mouse strains differ? Probability estimates and some implications. J. Hered. 72:27-31. Li, R., Lyons, M.A., Wittenburg, H., Paigen, B., and Churchill, G.A. 2005. Combining data from multiple inbred line crosses improves the power and resolution of quantitative trait loci mapping. Genetics 169:1699-1709.

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Linder, C.C. 2001. The influence of genetic background on spontaneous and genetically engineered mouse models of complex diseases. Lab Anim. 30:34-39.

Rosenstreich, D.L. and Glode, L.M. 1975. Difference in B cell mitogen responsiveness between closely related strains of mice. J. Immunol. 115:777-780.

Markel, P., Shu, P., Ebeling, C., Carlson, G.A., Nagle, D.L., Smutko, J.S., and Moore, K.J. 1997. Theoretical and empirical issues for marker- assisted breeding of congenic mouse strains. Nat. Genet. 17:280-284.

Schuster-Gossler, K., Lee, A.W., Lerner, C.P., Parker, H.J., Dyer, V.W., Scott, V.E., Gossler, A., and Conover, J.C. 2001. Use of coisogenic host blastocysts for efficient establishment of germline chimeras with C57BL/6J ES cell lines. Biotechniques 31:1022-1026.

Mashimo, T., Lucas, M., Simon-Chazottes, D., Frenkiel, M.P., Montagutelli, X., Ceccaldi, P.E., Deubel, V., Gu´enet, J.L., and Despres, P. 2002. A nonsense mutation in the gene encoding 2 5 -oligoadenylate synthetase/L1 isoform is associated with West Nile virus susceptibility in laboratory mice. Proc. Natl. Acad. Sci. U.S.A. 99:11311-11316. Mashimo, T., Voigt, B., Tsurumi, T., Naoi, K., Nakanishi, S., Yamasaki, K., Kuramoto, T., and Serikawa, T. 2006. A set of highly informative rat simple sequence length polymorphism (SSLP) markers and genetically defined rat strains. BMC Genet. 7:19. Mattson, D.L., Dwinell, M.R., Greene, A.S., Kwitek, A.E., Roman, R.J., Jacob, H.J., and Cowley, A.W. Jr. 2008. Chromosome substitution reveals the genetic basis of Dahl saltsensitive hypertension and renal disease. Am. J. Physiol. Renal Physiol. 295:837-842. Morse, H. C. III. 1978. Origins of Inbred Mice, Academic Press, San Diego. Mott, R., Talbot, C.J, Turri, M.G., Collins, A.C., and Flint, J. 2000. A method for fine mapping quantitative trait loci in outbred animal stocks. Proc. Natl. Acad. Sci. U.S.A. 97:12649-12654. Nadeau, J., Singer, J., Matin, A., and Lander, E. 2000. Analyzing complex genetic traits with chromosome substitution strains. Nat. Genet. 24:221-225. Ogonuki, N., Inoue, K., Hirose, M., Miura, I., Mochida, K., Sato, T., Mise, N., Mekada, K., Yoshiki, A., Abe, K., Kurihara, H., Wakana, S., and Ogura, A. 2009. A high-speed congenic strategy using first-wave male germ cells. PLoS ONE. 4:e4943. Paigen, K. and Eppig, J.T. 2000. A mouse phenome project. Mamm. Genome 11:715-717. Petkov, P.M., Ding, Y., Cassell, M.A., Zhang, W., Wagner, G., Sargent, E.E., Asquith, S., Crew, V., Johnson, K.A., Robinson, P., Scott, V.E., and Wiles, M.V. 2004. An efficient SNP system for mouse genome scanning and elucidating strain relationships. Genome Res. 14:18061811. Poiley, S.M. 1960. A systematic method of breeder rotation for non-inbred laboratory animals colonies. Proc. Anim. Care Panel 10:159. Poltorak, A., He, X., Smimova, I., Liu, M.Y., Van Huffel, C., Du, X., Birdwell, D., Alejos, E., Silva, M., Galanos, C., Freudenberg, M., Ricciardi-Castagnoli, P., Layton, B., and Beutler, B. 1998. Defective LPS signaling in C3H/HeJ and C57BL/10ScCr mice: Mutations in Tlr4 gene. Science 282:2085-2088.

Silver, L.M. 1995. Mouse Genetics. Concepts and applications. Oxford University Press, Oxford. Also see Internet Resources. Snell, G.D. 1948. Methods for the study of histocompatibility genes. J. Genet. 49:86-108. Specht, C.G. and Schoepfer, R. 2001. Deletion of the alphasynuclein locus in a subpopulation of C57BL/6J inbred mice. BMC Neurosci. 2:11. Stevens, J.C., Gareth T., Banks, G.T., Festing, M.F.W. and Fisher, E.M.C. 2007. Quiet mutations in inbred strains of mice. Trends Mol. Med. 13:512-519. Sundberg, J.P. and Schofield, P.N. 2010. Commentary: Mouse genetic nomenclature: Standardization of strain, gene, and protein symbols. Vet. Pathol. 47:1100-1104. Threadgill, D.W., Dlugosz, A.A., Hansen, L.A., Tennenbaum, T., Lichti, U., Yee, D., La Mantia, C., Mourton, T., Herrup, K., and Harris, R.C. 1995. Targeted disruption of mouse EGF receptor: Effect of genetic background on mutant phenotype. Science 269:230-234. Threadgill, D.W., Hunter, K.W., and Williams, R.W. 2002. Genetic dissection of complex and quantitative traits: from fantasy to reality via a community effort. Mamm. Genome 13:175-178. Tucker, P.K., Lee, B.K., Lundrigan, B.L., and Eicher, E.M. 1992. Geographic origin of the Y chromosomes in “old” inbred strains of mice. Mamm. Genome. 3:254-261. Wade, C.M., Kulbokas, E.J. 3rd, Kirby, A.W., Zody, M.C., Mullikin, J.C., Lander, E.S., LindbladToh, K., and Daly, M.J. 2002. The mosaic structure of variation in the laboratory mouse genome. Nature 420:574-578. Wakeland, E., Morel, L., Achey, K., Yui, M., and Longmate, J. 1997. Speed congenics: A classic technique in the fast lane (relatively speaking). Immunol. Today. 18:472-477. Wolfer, D.P., Crusio, W.E. and Lipp, H.P. 2002. Knockout mice: Simple solutions to the problems of genetic background and flanking genes. Trends Neurosci. 25:336-340. Yalcin, B., Nicod, J., Bhomra, A., Davidson, S., Cleak, J., Farinelli, L., Oster˚as, M., Whitley, A., Yuan, W., Gan, X., Goodson, M., Klenerman, P., Satpathy, A., Mathis, D., Benoist, C., Adams, D.J., Mott, R., and Flint, J. 2010. Commercially available outbred mice for genome-wide association studies. PLoS Genet. 6:e1001085. Yonekawa, H., Moriwaki, K., Gotoh, O., Miyashita, N., Migita, S., Bonhomme, F., Hjorth, J.P., Petras, M.L., and Tagashira, Y. 1982. Origins of

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laboratory mice deduced from restriction patterns of mitochondrial DNA. Differentiation 22:222-226. Zou, F., Gelfond, J.A., Airey, D.C., Lu, L., Manly, K.F., Williams, R.W., and Threadgill, D.W. 2005. Quantitative trait locus analysis using recombinant inbred intercrosses: Theoretical and empirical considerations. Genetics 170:12991311.

INTERNET RESOURCES http://www.informatics.jax.org/mgihome/nomen/ strains.shtml In 2001, the International Committee on Standardized Nomenclature for Mice (Chairperson: Dr. Janan T. Eppig) and the Rat Genome and Nomenclature Committee (Chairperson: Dr. G¨oran Levan) agreed to establish a joint set of rules for strain nomenclature, applicable to strains of both species. These guidelines are updated annually by the international nomenclature committees. The official Web site for these guidelines may be found at the above URL.

http://phenome.jax.org/ Official Web site of The Mouse Phenome Project, an international collaboration representing five countries in both the academic and corporate sectors. Its aim is to establish a collection of baseline phenotypic data on commonly used and genetically diverse inbred mouse strains through a coordinated effort. http://www.informatics.jax.org/external/festing/ search form.cgi An annotated list of mouse and rat inbred strain. http://www.isogenic.info/html/animal models in research.html Dr. M. F. W. Festing’s Web site about the best use of animal models. http://www.informatics.jax.org/morsebook/ Electronic version of the book by Herbert C. Morse III, Origins of Inbred Mice, Academic Press. 1978. http://www.informatics.jax.org/silverbook/ Electronic version of the book by Lee M. Silver. 1995. Mouse Genetics: Concepts and Applications, Oxford University Press.

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Mouse Breeding and Colony Management Abdelkader Ayadi,1 Gis`ele Ferrand,2 Isabelle Goncalves da Cruz,1 and Xavier Warot2 1 2

Institut Clinique de la Souris (ICS), Illkirch, France Ecole Polytechnique F´ed´erale de Lausanne, Lausanne, Switzerland

ABSTRACT The possibility to genetically modify the mouse genome has enabled the creation of numerous lines of genetically engineered mouse models (GEMMs). As a result, the demand for housing space in research facilities is increasing. Knowledge of the basis of mouse reproduction and of the methods to handle colonies of GEMMs is therefore mandatory to efficiently populate facilities. The mouse has a short generation period, produces large progenies, and can breed all year round. However, environmental parameters (bedding, diet, cage type, temperature, hygrometry, light, noise, and sanitary status) strongly influence the breeding efficiency and experimental data, and must be tightly controlled. Efficient GEMM colony management requires adequate recording of breeding and proper identification and genotyping of animals. Various mating types and breeding schemes can be used, depending on the type of studies conducted. The recent development of assisted reproduction methods helps circumvent some of the issues faced with those lines especC 2011 by John Wiley & Sons, Inc. ially difficult to breed. Curr. Protoc. Mouse Biol. 1:239-264  Keywords: mouse r reproduction r breeding r efficiency r mating r colony management r assisted reproduction

INTRODUCTION The mouse has been used as a research model for many years. The laboratory mouse was established in the early years of the twentieth century by A. Lathrop, W. Castle, and C.C. Little (for a history of the laboratory mouse, see Davisson and Linder, 2004). With the development of international initiatives to systematically knock-out all the genes of the mouse (The International Mouse Knockout Consortium, 2007), the number of genetically engineered mouse models (GEMMs) has grown rapidly over the last years. More and more research groups have access to mouse models and are also able to set up matings to generate their model of interest. A good understanding of the mouse as an animal and as a model is necessary to produce scientifically relevant data. This article summarizes the relevant information to breed and manage a mouse colony. In the first part, key physiological data regarding mouse reproduction is presented. The quantification of reproduction efficiency and the calculation of the production efficiency index (PEI) of a colony are also explained. The environmental and sanitary parameters affecting the breeding efficiency are detailed in the second and third sections of this article, respectively. The microenvironment and

the macroenvironment of the mouse in a facility are also described. In addition, monitoring the health status and its influence on breeding efficiency is exemplified. In the fourth part of this article, focus is placed on the management of a mouse colony, with a number of practical aspects. The basic tools required to managing a colony, i.e., breeding records, identification, and genotyping, and the importance of the genetic background, are presented. Different mating types and breeding schemes that can be applied are also exemplified. Tips and tricks for troubleshooting issues with breeding are highlighted. Finally, assisted reproduction methods and the way they can be applied to circumvent breeding issues are reviewed.

BASICS OF MOUSE REPRODUCTION The mouse is a mammal that has a long history of use in scientific studies. Mice are easy to handle and house, breed all year round, have a short generation period, and produce relatively large progenies. Mice tolerate inbreeding rather well compared to other species. The genetics of the mouse has been extensively studied and is now well documented; as such, the mouse genome can be easily manipulated.

Current Protocols in Mouse Biology 1: 239-264, March 2011 Published online March 2011 in Wiley Online Library (wileyonlinelibrary.com). DOI: 10.1002/9780470942390.mo100214 C 2011 John Wiley & Sons, Inc. Copyright 

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Physiological Data Mice belong to the Muridae family, which encompasses numerous species and genera. The laboratory mouse belongs to the genus Mus, species musculus. The strains used today were derived from a limited number of strains that were genetically diverse, namely M.m. musculus, M.m. castaneus, M.m. domesticus, and the hybrid strain M.m. molossinus. In the eighteenth century, mouse fanciers in Asia had intercrossed and domesticated many varieties as pets. These lines were imported into Europe and the United States, and mating programs were set up with a limited number of founder fancy mice, giving rise to the modern classical strains. The classical strains used in the laboratories can be divided into two groups: (1) the inbred strains, which have been established by inbreeding of siblings for >20 generations. All individuals from an inbred strain are genetically identical and their genomes are homozygous. (2) The outbred stocks, which are a population of genetically diverse individuals; their genomes are heterozygous.

Inbred strains and outbred stocks are widely used and exhibit different breeding performances (see Quantification of Reproduction Efficiency: Production Efficiency Index; also see Gu´enet and Benavides, 2011). Although genetically diverse, the physiological and breeding data of the different strains and stocks are similar and are listed in Tables 1 and 2. The following factors are of particular importance for the line maintenance and colony management and will be taken into account when setting up a mating strategy (see Colony Management). (1) The litter size is variable and highly dependent on the strain or stock used. (2) The mouse is a species with postpartum estrus, meaning that estrus with ovulation and corpus luteum production occurs immediately following the birth of the young. (3) The reproductive life span starts between 7 and 8 weeks and lasts for ∼8 months (The Jackson Laboratory, 2009). (4) The mating type and the breeding scheme will be set up according to the specifics of the strain studied.

Table 1 Physiological Data of the Laboratory Mousea

Longevity

1.5–3 years

Body temperature

35.5◦ –39.0◦ C

Breathing rate

140–250 breaths/min

Heart rate

325–780 pulse/min

Adult body weight

20–40 g

Male sexual maturity

50 days

Female sexual maturity

50–60 days

Body weight upon weaning

10–12 g

Weaning time

18–21 days

a Adapted from Havenaar et al. (1993).

Table 2 Breeding Data of the Laboratory Mousea

Mouse Breeding and Colony Management

Gestation

18.5–21 days

Estrous cycle

4 days

Birth weight

1g

Eyes open

11 days

Haircoat

8 days

Weaning

18–21 days

Productive breeding life

8 months

Postpartum estrus

Yes

Litter size

2 to 12 pups

a Adapted from Havenaar et al. (1993) and Hardy (2004).

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Table 3 Production Index Efficiency (PEI) for the Most Commonly Used Inbred Mouse Strainsa

Strain

Production efficiency index (PEI)b

C57BL/6

0.50

C3H

0.80

BALB/C

0.80

129S2/SVPas

0.80

DBA/2

0.64

FVB

0.80

CD1 (outbred)

2.00

a Adapted from White (2007). b PEI values are an average of several production sites and colonies.

Quantification of Reproduction Efficiency: Production Efficiency Index (PEI) Impaired breeding performance or reduced fertility in genetically engineered mice (GEM) can result in the loss of valuable research time and can be a serious issue for planning a mouse research project. It is therefore crucial to assess the reproduction efficiency in the specific facility. This helps to define mice production goals and schedule of the project. The following indices can be tracked to assess reproductive performance of the breeding colonies: (1) time between mating and first litter; (2) interval between litters; (3) litter size: number of pups born per litter; (4) weaning rate: numbers of pups weaned per litter (also called wean-to-born ratio). Breeding efficiency can be affected by a variety of specific events such as breeding failures (males and/or females), loss of newborn pups (poor mothering or cannibalism), or loss of breeders. To take into account all of these events, a convenient way to easily monitor the overall productivity of a colony is the production efficiency index (PEI), which corresponds to the number of weaned pups divided by the total number of females in a given time period (Festing and Peters, 1999). In general, the PEI is used by institutions and commercial breeders as follows: PEI = number weaned pups/female/week Reproductive performance varies widely with the genetic background of the strain. Traditional inbred mouse strains have defined reproductive parameters that are considered characteristic of the strain (Silver, 1995). Table 3 shows the PEI values obtained with the most commonly used strains of mice.

The indexes given in Table 3 should be used only as a guideline. As discussed below (see Factors Affecting Breeding Efficiency: Environmental Parameters), the breeding performance of mice relies on their environment. PEI is therefore specific not only to the strain, but also to the housing conditions under which the colony is being maintained. PEI of a breeding colony should therefore be monitored in any facility, taking into account the local specificities related to the breeding environment. Breeding performance variations between the wild-type inbred/outbred strain and the genetically modified mice of the same background strain should also be taken into consideration. Genetic mutations can alter the reproductive profile through different mechanisms and not only those which are directly related to breeding (Matzuk and Lamb, 2008; Naz et al., 2009). Regardless of whether inbreds, outbreds, or GEMMs are being produced, PEI is a specific production index that is a valuable tool to be used for planning production and to fulfill research demands (Festing and Peters, 1999).

FACTORS AFFECTING BREEDING EFFICIENCY: ENVIRONMENTAL PARAMETERS Control of the housing conditions is a key factor for ensuring efficient reproduction in a vivarium. One can distinguish between the microenvironment and the macroenvironment of the mice. The microenvironment of a mouse is the physical environment immediately surrounding it, i.e., the primary enclosure (cage) with its own temperature, humidity, and gaseous and particulate composition of the air. The physical environment of the secondary

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enclosure such as a room constitutes the macroenvironment. Most of the environmental parameters adequate for the proper housing of laboratory mice are detailed in the Guide for the Care and Use of Laboratory Animals (National Research Council, 2010). The authors have summarized the effects of the environment on mouse breeding and have drawn attention to the way one can modify them when evaluating breeding issues.

Microenvironment: The Cage Unit There are different ways of housing mice in a facility, including open cages, static microisolation cages, individually ventilated cages (IVCs), and isolators. The most commonly used housing system consists of IVCs grouped on racks. They act as a barrier-atthe-cage-level to keep out or keep in microorganisms. IVCs were developed as a solution to the problem of protecting the mice from pathogenic organisms and at the same time enabling open access to animals for researchers.

Cage

Mouse Breeding and Colony Management

The cage floor area is an important parameter for the breeding of mice. Cage space recommendations have been published and are now part of the European Union regulation (Voipio et al., 2010). It is crucial for efficient breeding to avoid overcrowded cages as this will result in loss of offspring due to physical overcrowding. Changes in cage air, temperature, light levels, and noise levels are critical environmental parameters for proper housing and breeding of mice (see below). Whereas those parameters can be easily monitored at the macroenvironment level, their monitoring at the microenvironment level is much more difficult. As an example, recording the light intensity at the cage level in the authors’ vivarium revealed huge differences from one cage to another, depending on the location of the cage on the rack (Ayadi et al., unpub. observ.). The actual microenvironment parameters are mostly unknown, making it difficult to define the best parameters for efficient breeding. Nevertheless, in case of breeding difficulties, changing the cage location on a rack or in the room could be evaluated. Depending on the type of IVCs, the change rate of the cage air can be controlled more or less accurately. Cage air change rate is important to supply sufficient oxygen for the animals and to eliminate heat, excess humidity, carbon dioxide, and any potential noxious compounds

that may accumulate in the cage (such as ammonia). IVCs with separate air handling units will provide a more consistent and monitored environment than cages ventilated through the general heating, ventilating, and air conditioning (HVAC) system.

Bedding Bedding, food, and water are critical for a mouse breeding colony. Various types of bedding (alpha cellulose, aspen, corn cob, poplar, etc.) can be used in the cage and will influence the wellness of the animals. The bedding type will have a direct effect on the microenvironment (e.g., by affecting the ammonia level in the cage). It has been shown that the lowest ammonia concentration has been observed in cages that house mice on hardwood bedding or a combination of corn cob and alpha cellulose (Smith et al., 2004).

Diet Nutrition has a major influence on reproduction and providing an adequate diet is important to ensure efficient breeding. The diet should be provided ad libitum with all the essential nutrients in sufficient amounts. Extensive studies have been carried out to determine the needs of the mouse to sustain gestation and lactation. Breeding diet formulations have been developed by manufacturers, which mainly consist of a diet with increased protein and fat levels. Proteins can be of animal or vegetal origin. Those specific breeding formulations have proven to promote good breeding performance in the authors’ facilities. Enriched diets can be used for strains difficult to breed. The presence of phytoestrogens can be of particular concern, due to the interaction with the natural estrogens and possible disruption of the cycling of the females. Food sanitization—sterilization or pasteurization—is essential to the prevention of infectious diseases in animal facilities. Sterilization is a process that frees the diet from any living organisms and also destroys all spores. Pasteurization is a partial sterilization that destroys organisms but not the spores. The most commonly used methods of sanitization (sterilization or pasteurization) are steam and γ irradiation. Irradiation is an expensive method compared to autoclaving, which is cost effective; however, irradiation ensures a higher microbiologic stability of the diet in the facility as well as less nutrient loss, with no effect on the hardness of the food materials.

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Water should be provided ad libitum as well. Guidelines recommend providing animals with clean, fresh drinking water. Laboratory animal drinking water standards have not been defined. Thus, the water quality varies considerably from one location to another, together with the treatment applied to the water in the vivarium (such as acidification, chlorination, filtration, ionization, or autoclaving; Lemken, 2010). However, in the authors’ experience, the exchange of animals between two different institutions located in different countries does not result in any evidence of potable water quality influencing the breeding efficiency.

Nest Nesting material is often considered as a type of enrichment and has proven to be of importance for efficient reproduction. The nest is an efficient buffer against variations of the environmental parameters in the cage, variations that can be deleterious to the neonate. Paperderived materials such as tissues, towels, or paper strips can be used as nesting material. Again, on top of a standard bedding material, extra enrichment material can be added for strains difficult to breed, for example, little mouse houses together with tissues.

Macroenvironment: Ventilation, Temperature, Hygrometry, Noise, and Light The control of physical parameters, such as temperature, hygrometry, ventilation, light, and noise, is a critical point for the well-being of the animals housed in the vivarium and for the efficiency of breeding. Mice are homoeothermic mammals, which require, for their well-being, maintenance of their body temperature in their thermoneutral zone. The usual recommendation for the tem-

perature of the housing rooms is 22◦ C ±2◦ C, which is comfortable for humans. It has been shown that the ambient temperature selected by mice in a temperature gradient experiment is much higher than this temperature (Voipio et al., 2010). Mice must therefore make adjustments to maintain their body temperature in their thermoneutral range, which can affect their breeding performance. Increasing the temperature of the housing room can help increase breeding efficiency, and is of particular interest in housing rooms where transgenesis is performed and pseudopregnant implanted females are housed. Hygrometry is tightly linked to temperature and should always be in the midrange, i.e., relative value 55% ±10%. As mentioned in the Microenvironment section above, cage air change rate is crucial. The ventilation system supplies sufficient oxygen and eliminates heat and carbon dioxide and any potential noxious compounds. The cage air change rate is defined at the macroenvironment and microenvironment levels. Typical values for mice rooms are between 13 and 15 air changes per hour. The air flow can be a source of discomfort for the animals, which may influence the breeding. The light intensity can affect the physiology and behavior of the animals (CastelhanoCarlos and Baumans, 2010). The most important environmental parameter is regulating the temporal pattern of animal behavior and physiology, the circadian rhythms, and stimulating and synchronizing the breeding cycles. Mice are nocturnal animals, being active mostly during the dark. Light cycles should therefore be regular and carefully monitored in the facility, with special attention to the dark phase, which should not be interrupted by exposure to light during this cycle. The light cycles should be carefully verified if changes in the breeding performance are observed.

Table 4 Micro- and Macro-Environmental Parameters and Breeding

Bedding material Diet composition Nesting and enrichment material Cage position in the room / on the racks Room temperature and hygrometry Light cycle and photoperiod Noise levels Sanitary status

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Attention must be paid to the noise in the vivarium (Castelhano-Carlos and Baumans, 2010). Having a constant but not uniform noise level in the housing rooms reduces the stress of the animals when mice are exposed to the sounds that occur during changing of cages or cleaning of the rooms. However, the most important point is that the greatest sensitivity to mice is to frequencies that are inaudible to humans. Mice hear sounds above 64 kHz, compared to 20 kHz, the maximum a human being can hear. Mice emit a peculiar set of ultrasounds during mating, which is being studied by scientists (Castelhano-Carlos and Baumans, 2010). Thus, ultrasound can interfere with the reproductive behavior. Although not apparent, noise should be considered when evaluating the causes of decreased breeding performance. A summary of the environmental parameters influencing the breeding efficiency is given in Table 4.

FACTORS AFFECTING BREEDING EFFICIENCY: SANITARY STATUS

Mouse Breeding and Colony Management

Numerous factors affect breeding and among them pathogenic microorganisms can directly or indirectly have an effect on reproduction. The presence or absence of pathogenic microorganisms in a mouse population defines the sanitary status of the population. Health monitoring is therefore an integral part of any mouse breeding program. There is no universal definition of mouse sanitary status and there is no standard list of microorganisms to monitor. Recommendations have been established by scientific societies such as the Federation of Laboratory Animal Science Associations (FELASA) in Europe and various commercial breeders in the United States. A list of microorganisms to screen for and the frequency of the screening has been defined by FELASA recommendations (Nicklas et al., 2002; Fig. 1). Different health statuses can be found in a facility, starting with extremely “clean” (absence of microorganisms, opportunistic or pathogenic, in the mouse population) to “dirty” (presence of numerous different types of microorganisms). The various sanitary statuses are summarized in Figure 2. The main criterion for the definition of sanitary status is the presence of a particular pathogenic microorganism in the mouse. In most cases, breeding facilities are classified by the following health statuses:

1. Specific pathogen free (SPF) status. 2. Conventional status, meaning that some pathogens are present in the breeding facility, without any restriction regarding the type of pathogens.

Health Monitoring Strategies The health status of a breeding facility is monitored by a program that employs microbiologic screening most of the time using “sentinel” animals, which identify all of the microorganisms present in the facility and are representative of the population of animals of the facility. Apart from the sentinels, in the case of specific questions, alternate techniques can be implemented (e.g., PCR to detect for the presence of microorganisms on equipment in the facility).

Why monitoring? The presence of an infected mouse in a vivarium is a problem, with the following various characteristics: (1) the ethical aspect (the animal may suffer needlessly); (2) the sanitary aspect (the animal can spread an infection throughout the facility); (3) the research aspect (the experiments can be biased or modified by the infection); and (4) the breeding aspect (an infected mouse presents a high risk of impeding breeding). It is therefore important to regularly monitor the presence of potentially pathogenic microorganisms. Furthermore, some mouse pathogens are zoonotic and it is essential to monitor the health status from an occupational health perspective. The most important viruses transmitted from mice to humans are the Haantan virus (Korean hemorrhagic fever) and the LCMV (lymphocytic choriomeningitis virus).

Why health monitoring? For a barrier unit, it is important to define two categories: (1) a list of agents to monitor (e.g., FELASA list and additional agents); and (2) a list of exclusions (i.e., the microorganisms that should not be present in the facility). The FELASA list (see above) is a good starting point to define the health monitoring program. This “official list” is currently in the process of re-evaluation. Microorganisms can be added to the FELASA screening list depending on the research being done and the needs of the facility (e.g., norovirus, which affects the immune system and may be detrimental for immunocompromised mouse lines;

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Date of issue: Location: Housing (barrier, non-barrier, IVC, isolator): Species: mouse Strain: Species and strains present within the unit: Test frequency

Latest test date

Latest results

Testing laboratory

Test Historical results method (ⱕ18 months)

Viruses Mouse hepatitis virus Mouse rotavirus (EDIM) Parvoviruses Minute virus of mice Mouse parvovirus Sendai virus Theiler’s murine encephalomyelitis virus Ectromelia virus Lymphocytic choriomeningitis virus Mouse adenovirus type 1 (FL) Mouse adenovirus type 2 (K87) Mouse cytomegalovirus Reovirus type 3 Additional organisms tested: Bacteria, mycoplasma, and fungi Citrobacter rodentium Clostridium piliforme (Tyzzer’s disease) Corynebacterium kutscheri Mycoplasma spp. Pasteurellaceae Salmonella spp. Streptococci ␤ hemolytic (not group D) Streptococcus pneumoniae Helicobacter spp. Streptobacillus moniliformis Additonal organisms tested: Parasites Ectoparasites: Species designation Entoparasites: Species designation Pathological lesions observed Data are expressed as number positive/number tested Positive findings in other species in the same unit: Abbreviations used in this report: CULT, culture; ELISA, enzyme linked immunosorbent assay; HIST, histopathology; IFA, immunofluorescence assay; MICR, microscopy; NT, not tested; PATH, gross pathology; PCR, polymerase chain reaction.

Figure 1

Health monitoring in accordance with FELASA recommendations (adapted from Nicklas et al., 2002).

“Dirty”

“Clean” Containment

Isolator maintained

Terminology

Germ-free

Barrier maintained Defined flora

Specific pathogen free (SPF)

No containment Virus antibody free

Conventional

Figure 2 Common terminology for microbiological status of laboratory rodents (adapted from National Research Council, 1991).

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the opportunistic bacteria Pneumocystis carinnii, which affects some immunocompromised lines). Some opportunistic agents can be used as indicators to assess the efficiency of the barrier and to monitor the working procedures. Zoonotic agents such as Haantan virus, LCMV, Encephalitzoon cuniculi (rare), and Streptobacillus moniliformis (rat bite fever, rare) should be taken into consideration and added to the monitoring list, particularly in the case of pregnant employees.

How to perform the monitoring?

Mouse Breeding and Colony Management

The most common way of monitoring the health status is through a dirty bedding sentinel program. Briefly, na¨ıve adult (at least 8 weeks old) mice are challenged with dirty bedding from all the cages of the facility (collected while cleaning the cages). Dirty bedding is defined as bedding with feces (as most of the pathogens are transmitted through an oral-fecal cycle). As a result of exposure to pathogens, the na¨ıve animals will host the pathogens and produce antibodies. After a defined time of exposure, a serological analysis will be done, and the presence of specific antibodies will be an indicator of the presence of a specific pathogen. The dirty bedding sentinel program efficiency is highly related to the type of housing used in the facility. When working with open cages, all airborne pathogens can easily be transmitted through airborne transmission and transmitted to other cages by the activity of the animals and personnel. Putting the sentinel cage at the bottom row location of a rack will increase the exposure of sentinels to the dirtiest room environment. When using IVCs, airborne pathogens are less likely to be transmitted to sentinels. The sampling in IVCs can be improved by adding food and water bottle contents taken from an occupied cage. The optimum number of cages monitored with sentinel cages has not been determined. Having too many cages monitored by one cage of sentinels will result in diluting the agents to be detected and no immune response to a particular agent will result. Recommendations on how to calculate the best sample size (i.e., how many cages to sample to find a disease regarding its prevalence) can be found in the FELASA guidelines. However, this method of calculating the sample size is relatively difficult to use when working with IVCs. The authors therefore recommend having at least one cage of sentinel animals per side of ventilated rack. If several research groups share the

same rack, then it is worth having one cage of sentinels per group. The frequency of the monitoring depends on the needs of the breeding program. Maintenance of an SPF sanitary status will require regular and in-depth monitoring. Specific agents may be monitored with special attention if they represent a risk for the colonies and/or the type of research performed in the facility. The authors recommend monitoring quarterly for the most prevalent microbiologic agents and annually for the atypical ones. As an example, when working in a breeding facility with a conventional sanitary status, testing of viruses and parasites quarterly and testing of bacteria once or twice a year are recommended. With such monitoring, the pathogens present in the facility can be identified and documented. Several methods of analysis, direct or indirect, can be used to identify the microbial agents present in a breeding facility: 1. Direct and microscopic observation of parasites such as pinworms (presence in the caecum); 2. Culture for bacteria identification; 3. Serological analysis (detection of antibodies) for virus identification. The caveat of this method lies in the fact that only certain strains of mice develop antibodies against viruses, leading to false-negative results. For example, the parvovirus will trigger an immune response in NMRI mice but not in C57BL/6 animals; 4. PCR for detection of an agent, bacteria, or virus. Finally, one key point to keep in mind is that the results of the health monitoring program provide a retrospective image of the microorganisms present in the facility. For example, if sentinel mice are challenged between January 1st and March 31st , the results of the analysis will be obtained by the end of April. The overview of what happened between January 1 and March 31 will be therefore known only at the end of April. Any positive findings correspond to an event that took place sometime during the last 3 months, making it difficult to find the origin of the positive findings and to define the extent of the contamination of the breeding facility, as well as to determine the best strategy to eradicate the pathogens.

Pathogens Affecting Breeding Efficiency All pathogens that affect animal health will have an influence on breeding. The pathogens can have an effect on the pregnant females

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Table 5 Non-Exhaustive List of Agents that Can Affect Breeding

Viruses Epizootic diarrhea of infant mice (EDIM): Infects young pups (<12 days), which stop suckling Murine hepatitis virus (MHV): Neonatal diarrhea, which can cause death Bacteria Helicobacter (hepaticus or bilis): Responsible of rectocolitis, which can lead to rectal prolapse and is life threatening for the mother Pasteurella pneumotropica: Infects vagina or uterus Staphylococcus aureus: Is frequently responsible of preputial gland abscesses in young C57BL/6J or C3H males Parasites Pinworms (Syphacia muris, Syphacia obvelata, or Aspiculuris tetraptera): Results in diarrhea and eventually rectal prolapse in heavily infested animals

(e.g., an infection with pinworms will deprive the females of proteins and vitamins necessary for the progeny) or on the offspring (e.g., lethal viral neonatal diarrhea). Table 5 provides a few examples of agents that can affect breeding.

COLONY MANAGEMENT As a result of genetic engineering methodologies, the number of genetically modified mouse lines is rapidly expanding, resulting in increased demands for housing space in research facilities. Associated research costs are increasing in the same manner. Effective handling of colonies is therefore essential in order to use space efficiently and controlling production costs. Regardless of these economic considerations, breeding colony management must achieve the goals of any particular experimental study with respect to the 3Rs principle by using only animals that are necessary (Russell and Burch, 1959). The 3Rs (Reduce, Replace, Refine) is a landmark principle aimed to improve animal welfare and reduce suffering, to minimize the numbers of animals used without compromising the output and quality of the scientific study, and to replace animal models with non-animal ones wherever possible while still achieving the scientific objectives. Described below are some efficient and reliable techniques to ensure safe and economical GEMMs management.

Breeding Records Keeping careful and accurate records is an important aspect for maintaining breeding colonies by minimizing animal reduction (the 3Rs) and containing costs. The breeding data records ensure that production meets the

expected research goals. Regular analysis of records that monitor the production process and trends helps to detect abnormal events. If required, breeding colonies can be adjusted to meet the production objectives. When working with GEMMs, experimental or clinical observation data records are of utmost importance to track genetic and phenotypic variation, and for welfare assessment. These observations and other animal-specific events should be kept as a separate log. The minimal requirement to track data is the cage card. Information recorded on cage cards must contain parameters that enable one to know history, pedigree, purposes, and performance. The cage card should list mouse room identification or localization, name of the principal investigator and caretaker, cage number, strain, line number, animal IDs, gender, date of birth, genotype and basic pedigree (or parentage), and generation number. In addition, complete breeding records with breeding setup, successive litters with dates of birth, number of pups born, number of pups weaned, sex, and genotype should also be recorded. These data insure productive breeding colonies. See Figure 3 for an example of a breeding card with breeding records.

Identification and Genotyping Practical colony management relies on efficient and reliable identification and genotyping methods. Individual identification is necessary to trace a specific animal within a cage to track its history for breeding performance, in order pedigree purposes, mating schemes, and phenotyping studies. In a similar way, accurate

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A

Breeding cage card

Line ID:

PI:

Set up date:

Room no.

Retirement date:

Cage no.

genotype

ID

DOB

ID

No. of Pups born Litter no. DOB total

B

genotype

DOB

No.of Pups weaned total ID listing Remarks

Mouse colony log

Mouse ID

DOB

Sex

Genotype Line ID source

Cage no. source

Parentage Sire Dam

Genetic background

Cage no.

Note

Figure 3 Example of a cage card with breeding records and of a colony log to manage a GEMM mouse colony. (A) Important information such as line name, owner of the line, animal ID, gender, date of birth, genotype, and parentage should be indicated on the breeding card. For an efficient breeding program, breeding set-up and retirement dates should be known. (B) A separate mouse colony log with the data of all animals of the colony (animals used for breeding, experimentation, and stock) is necessary to manage the colony in an effective and safe way. Table 6 Different Mouse Identification Methods Depending on Age of Micea,b

Age Technique Temporary Permanent

<1 week 1–2 weeks Weaning time Post-weaning (3–4 weeks) time

Mark on tail

Yes

Yes

Yes

Yes

Coat marking

No

No

Yes

Yes

Ear notching

No

No

Yes

Yes

Toe clipping

Yes

No

No

No

Ear tag

No

No

Yes

Yes

Tattooing

Yes

Yes

Yes

Yes

Transponder

Yes

Yes

Yes

Yes

a Adapted from the BVAAWF/FRAME/RSPCA/UFAW, Joint Working Group on Refinement (2003). b The choice and use of the method must be approved by the institutional animal use and care committee and

should be compliant with national laws and regulations (e.g., toe clipping is allowed for mice up to 12 days in Switzerland).

genotyping is also a critical part of research and breeding programs. Mouse Breeding and Colony Management

Mouse identification methods Several identification methods have been used to identify individual mice such as ear tagging, ear punching, toe clipping, toe

or tail tattooing, and microchip implantation (Table 6). The choice of the most appropriate method should be based on (1) the research purposes (nature and duration of the experiment); (2) the reliability of the identification needed (temporary versus permanent); (3) the age of the animal; (4) the technical expertise;

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10

1

20

2 40

4

30

3

5

1

4

50

10

40

6

2

4

60

20

40

7

3

4

70

30

40

8

1

2

80

10

20

9

2

3

90

20

30

Figure 4 Ear notching in mice. Example of a numbering system for identifying mice. The scheme represents the dorsal view of the head of a mouse, with the position of notches within the ear. Units correspond to notches in the right ear, tens to notches in the left ear. Numbers 5 to 9 and 50 to 90 are created by a combination of two notches, as indicated. By combining notches for units and tens, one is able to number animals starting at 1 up to 99.

and (5) the number of characters in the identification code system (continuous numbering schemes). Whatever the method, it must be approved by the institutional animal use and care committee. Ear notching One of the most common identification methods is ear notching using a code system that links the number and location of ear notches to generate specific numbers (Fig. 4). This method is inexpensive, but problems with reading the code are not unusual. It is not uncommon for the holes or notches to close over a period of time. Therefore, it is important to check the markings regularly to be certain the animal can still be identified accurately. If housed in groups, mice tend to fight and they may rip the ears of others and cause ID information to be lost. Toe clipping Toe clipping is another widely used method to identify mice before weaning, because it is a reliable, easy, and fast technique to mark animals. This method however is controversial because of its potentially negative impact on animal welfare. Current recommendations and guidelines indicate to use toe clipping as a

last alternative to identify animals as early as 1 day after birth up to 7 days of age (National Research Council, 2010; BVAAWF/FRAME/ RSPCA/UFAW, Joint Working Group on Refinement, 2003). Recent publications indicate that toe clipping has few adverse effects on animal welfare and behavior and may be an acceptable method for marking mice (Castelhano-Carlos et al., 2010; Schaefer et al., 2010), particularly if tissue removal from toes is used for genotyping. Tattooing Mice can also be identified by tattooing the tail using an electric tattooing device (such as a manual-AIMS system) or tattooing the paw by injecting ink subcutaneously using a fine-gauge needle (Ketchum system). However, tattoo identification requires training, longer times to mark animals, and can be tedious depending on the numbering code system and the recommended anesthesia to minimize animal handling and distress. Moreover, it might be difficult to read the identity in young pigmented mice. New tools and devices are available on the market to simplify the tattooing process. One of these tools is an automated tattoo device that marks consistent alpha-numeric IDs onto the tails of the animals

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(http://www.somarkinnovations.com). The automated tail tattoo is permanent and easily legible. Ear tagging Ear tagging is another easy way to identify mice with coded metal tags that are clipped to the base of the ear to reduce the chance of tear-out. Animals can however only be tagged during the weaning age, after which it can be a drawback. A new type of ear tag with a 2-D barcode on its surface has been recently developed (http://www.zonotid.com). This new ear tag is less likely to be pulled out and is very lightweight, which is more comfortable for the animals. Such eartags cannot be misread like the metal ones as they are scanned with a barcode reader. Microchip implantation Transponders or microchips using radio frequency identification (RFID) systems provide a fast and reliable method to identify mice. The microchips are implanted subcutaneously between the shoulders and can be implanted only a few days after birth due to the very small size (1 × 6–mm, 7.15-mg weight; Castelhano-Carlos et al., 2010). This method allows for identification of animals without handling or removing the animals from the cage. This method requires specific readers since there is no standardization on microchip frequencies used. It is somewhat costly compared to the other identification methods, but microchips are an easy, secure, and durable method of identification. Data registration can be linked to any breeding colony management database, which helps to ensure accurate collection of research data. An additional advantage is the transmission of body temperature without distressing the animal, which may be very useful (Le Calvez et al., 2006; van Gassen et al., 2008). One should keep in mind that implanted microchips may be damaged during imaging studies depending upon the imaging technique used. To avoid any issues, microchip specifications should first be checked with the manufacturer.

Genotyping

Mouse Breeding and Colony Management

The genotype is determined by the analysis of DNA extracted from young pup tissues by polymerase chain reaction (PCR) most of the time. Sufficient DNA for PCR analysis can be obtained from a variety of tissue sources including blood, ear punch tissue samples, tail tip biopsies (tail snips), toe sample, hair samples, stool, and oral swabs. The choice of the

sampling method should take into account: (1) the source of tissue, (2) the age of the animal, and (3) the genotyping method (the sample should provide enough high-quality DNA for the method of genotyping chosen). Most often GEMMs are identified and biopsied simultaneously for genotyping before weaning age by PCR, which provides fast and reliable results with a minimal quantity of DNA. It is important to get the genotyping results before the weaning step to avoid overcrowding and to save cage space. Some specific genetic models can be genotyped only by Southern blot analysis, which is technically more demanding and requires a large amount of DNA. Whatever technique is used, if genotypes are unclear, they must be repeated with the appropriate controls. Furthermore, during tissue sampling, one should avoid mixing samples as this can lead to false genotyping results. In addition, instruments used for sampling must be decontaminated or replaced between samples to avoid potential microbiological and DNA contamination. A poor-quality tissue sampling method or inaccurate genotyping may jeopardize the breeding program and research goals.

Genetic background Whether breeding or phenotyping is the purpose for managing a mouse colony, the genetic background should be taken into consideration. The genetic background represents strain characteristics that are influenced by a set of genes (allelic gene variation, various gene expression levels and patterns). Each strain has a unique set of characteristics. This is well documented for GEMMs, where phenotypes observed are very much dependent on the mouse strain harboring the mutation (Doetschman, 2009). The type of strain and/or stock used, its origin, and breeding strategies must be recorded. Monitoring of the genetic background is highly recommended for proper analysis of the phenotyping data. There are different strains and/or stocks available and various breeding strategies to take into account the genetic background parameter (Table 7).

Nomenclature Genetic engineering technologies have markedly increased the number of mouse strains available for research. A myriad of GEMM lines carrying spontaneous or induced mutations, transgenes, or targeted mutant alleles in various genetic backgrounds exist. To

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Table 7 Definitions of Various Strains and Stocks

Strain/stock

Definition

Inbred

Strain maintained by breeding of siblings for at least 20 generations. Mice are genetically identical. Highly desirable due to their high degree of homogeneity.

Hybrid

Mice are called F1 hybrids, resulting from mating between two distinct inbred strains. F1 hybrids as a population are genetically identical (except for sex chromosomes).

Outbred

Stock or population with genetic variation among individual mice. Outbred stock is maintained by a rotational breeding scheme to minimize the loss of heterogeneity.

Coisogenic

Strain differing from the parental inbred strain at one locus (such as a targeted gene modification, bred onto mice on the same genetic background that the ES cells were derived from).

Congenic

Inbred strain that contains a DNA segment from another strain. Mice are genetically identical except at a single locus and its surrounding region. Strain generated by backcrossing.

Consomic

Inbred strain that contains a full chromosome from another strain. Strain generated by backcrossing.

ensure that scientists can identify the kind of mouse acquired and to keep relevant information associated with the name of each line, a standardized nomenclature system has been established to correctly name genes, alleles, and mouse strains (R¨ulicke et al., 2007; Montoliu and Whitelaw, 2010). Having complete and proper information helps scientists to choose the most appropriate research model and will facilitate precise communication of scientific findings. Rules and guidelines for mouse nomenclature are set by the International Committee for Standardized Genetic Nomenclature in Mice (http://www.informatics.jax.org/mgihome/ nomen/inc.shtml). The Mouse Genome Informatics Database (MGD) serves as the central repository and is the authoritative source of official names for mouse genes, alleles, and strains. Nomenclature follows the rules and guidelines established. The most up-to-date rules and guidelines for genes, genetic mutations, strains, and chromosome aberrations are available at the MGI Mouse Nomenclature Home Page (http://www.informatics. jax.org/mgihome/nomen/#rag). The MGI Nomenclature Committee provides assistance in assigning a unique identifier to a gene or an allele ([email protected]). Several publications have summarized the principles to correctly design mouse strains (inbred, hybrid, and outbred), genes, alleles, and mutations. Examples of a gene-targeted and a transgenic line are given in Table 8.

For a gene-targeted line, the components of the standard designation start with the genetic background of the mouse strain. Embryonic stem cells (ES cells) that are available for gene targeting by homologous recombination may be derived from different strains (mainly 129 and C57BL/6 substrains) and the resulting chimera may be backcrossed to the same or to another mouse strain. The description of the name should clearly indicate whether it is an inbred, mixed, congenic, or coisogenic strain along with the history of the strain. The correct targeted gene symbol followed by tm (targeted mutation), an allele number, and the code of the laboratory or the investigator who produced the mouse line (all superscripted and in italics) follow the genetic background. Laboratory or investigator codes (usually three to four letters, first letter uppercase, followed by all lowercase) can be obtained from the Institute for Laboratory Animal Research (ILAR) Web page (http://dels.nas.edu/global/ilar/Lab-Codes). For a transgenic line, the name is made up of several similar components, with the background strain first, followed by the symbol Tg (transgenic line). Description of the construct is detailed between brackets, including the promoter, regulatory elements, and the transgene expressed. Next to this description comes the identifier of the transgenic founder followed by the laboratory code of the originating laboratory. Italics are not used for transgenes.

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Table 8 Standard Nomenclature of Strains and Various GEMMs with Detailed Descriptions

Mouse strain

Strain type

Description

C57BL6/J

Inbred

Breeding of siblings for at least 20 generations

Crl:CD1

Outbred

Name of outbred start with the laboratory code of the institution holding the stock followed by a colon and the common root stock name. Here the outbred is named CD1 stock of Charles River Laboratories (CRL).

B6;FVB-Tg(Ppm1a CreERT2 )43.3Ics

Mixed inbred

Transgenic mice (Tg) in a mixed genetic background made initially using FVB/N oocytes and backcrossed to C57BL/6N. The transgene is a construct with the promoter of the protein phosphatase 1A, magnesium dependent, alpha isoform gene (Ppm1a) driving ubiquitous expression of the inducible Cre recombinase Cre-ERT2 . Founder line 43.3 was established. This strain is produced and available from the Institut Clinique de la Souris (Ics). (http://www.ics-mci.fr/mousecre/results/synthesis?subline id=14).

B6;129-Tlr2tm1Kir

Mixed inbred

Mixed inbred knock-out strain initially produced in a 129 ES cell line and backcrossed less than five times to C57BL/6J mice (B6), as indicated by the semi-colon. This strain harbors the first targeted mutation (tm1a) of the toll-like receptor 2 (Tlr2) made in Carsten Kirschning’s laboratory (Kir) (Wooten et al., 2002).

B6.129-Tlr2tm1Kir /J

Congenic

Inbred toll-like receptor 2 (Tlr2) knock-out produced by Carsten Kirschning’s laboratory, backcrossed more than five times (note that the semi-colon is replaced by a period), and maintained by The Jackson Laboratory (J), stock number 004650.

B6.129-Ncoa6tm1.1Ics /Ics B6.129-Ncoa6tm1.2Ics /Ics

Congenic

Inbred Ncoa6 knock-out and floxed mutant mice. The nuclear receptor cofactor Ncoa6 gene (also called RAP-250) has been targeted to generate the constitutive (knock-out ) and the conditional (floxed) mutant mice. Due to the Cre/LoxP system, the same targeting event in ES cells allows the generation of two germline transmissible alleles after mating with a Cre transgenic mouse strain. The regular tm designation rules are applied with parental designation followed by a decimal point and serial number. In this example, Ncoa6 has been targeted in 129S2/SvPas ES cells by the Institut Clinique de la Souris (Ics). Ncoa6tm1.1Ics designates the null allele of Ncoa6 gene, whereas Ncoa6tm1.2Ics designates the conditional (floxed) allele. Mice have been backcrossed more than 9 times to C57BL/6J mice and the lines are maintained by the Ics (http://www.ics-mci.fr/nr zoo.html).

FVB/N-Tg(Acta2RAC1*G12V)33Pjgc/J

Congenic

Inbred transgenic mice (Tg) containing the constitutively active glycine to valine mutation at amino acid 12 (G12V) of the human RAC1 gene under the control of mouse smooth muscle alpha actin (Acta2) promoter. This transgene was microinjected into FVB/N oocytes. Founder line 33 was subsequently established. The mice were then crossed to FVB/N by the Pascal Goldschmidt-Clermont’s laboratory (Pjgc). This line is maintaining by The Jackson Laboratory (J), stock number 010634.

C57BL/6NTacTcf7tm1a(EUCOMM)Wtsi /Ics

Coisogenic

Inbred knock-out mouse strain in C57BL/6NTac genetic background derived from a JM8.N4 ES cell carrying the targeted mutation (tm1a) of the Tcf7 gene produced by the Wellcome Trust Sanger Institute (Wtsi) for the EUCOMM program. This line was produced through blastocyst injection of the ES clone (http://www.eucomm.org) and is maintained by the Institut Clinique de la Souris (Ics).

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Table 9 Examples of Different Mating Typesa

Pros

Cons

Applications

Monogamous permanent mating

Genealogical records; Female breeding records; Post-partum estrus

Space and time consuming; Male present at littering

Mutations, transgene; Project start

Permanent trios

Genealogical records; Post-partum estrus; Olfactory stimulation

Competition between litters; Male present at littering

Idem monogamous; Intermediate production level

Permanent harem mating

Post-partum estrus; Olfactory stimulation; Synchronization of littering

Fights; No genealogical records; Cage size; Competition between litters

Large-scale production

Boxing-out mating

Olfactory stimulation; Synchronization of littering; Genealogical records; Less competition between litters

No post-partum estrus; Large-scale Fights; production; Cage size Timed births

a Adapted from Hardy et al. (2004).

For both of these lines, the strain name may contain the symbol of the laboratory (ILAR code) maintaining the strain.

actly what the needs and goals of the experiment are. An overview of the different systems can be found in Donnelly et al. (2010).

Laboratory information management system (LIMS)

Mating Types

Record keeping is key for successful mouse colony management and welfare assessment. With the broad availability of GEMMs, a laboratory management information system (LIMS), rather than hand-written notebooks or spread-sheet applications, should be used regardless of the size of the colonies and the facility. Data sharing and communication are facilitated between technicians, colony managers, and investigators. This is of utmost importance for barrier level facilities with restricted access. The use of a controlled vocabulary avoids any misunderstanding about colony management. Furthermore, as most of the national legislations or institutional animal care and use committees now require a yearly accounting of animals used in a facility, with a LIMS one can easily establish the listing of animals, their number, their purposes, and functional categories (experimentation, breeding, stock). A number of software applications designed to manage mouse colony and phenotyping data are commercially or freely available. The main criterion for selecting a LIMS is to know ex-

The physiology of mouse reproduction allows different types of mating (summarized in Table 9). The choice of a given mating type depends on several parameters, among others the number of breeders, the number of animals to produce in a given time frame, the genealogical records needed, and the space available.

Breeding Strategies When working with GEMMs, it is important to consider the most appropriate genetic background, especially if phenotyping studies are planned. Depending on the type of research, some inbred strains are more suitable than others. Whatever the source of the GEMM strain, complete strain information is necessary in order to use appropriate breeding strategies.

Establishment of a new GEMM line from ES cell–derived chimera or transgenic founder When coisogenicity is required, a germ line competent chimera must be crossed with an inbred strain of the same strain from which the

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Table 10 Breeding Schemes with Genotype Expected Ratios

Breeder genotypes

Genotype ratio of offspring

Heterozygote × wild-type

50% heterozygote; Can be applied for colony expansion and maintenance 50% wild-type (backcrossing) or production of a cohort with dominant or semidominant mutant allele if homozygote mice are not viable or fertile

Heterozygote × heterozygote

25% homozygote; Can be applied for colony expansion and maintenance, 50% heterozygote; production of a cohort, or most appropriate breeding 25% wild-type scheme when working with mixed or undefined genetic background

Heterozygote × homozygote

50% homozygote; Can be applied for production of a cohort when working 50% heterozygote with inbred or congenic GEMM or optimization of the production of mutants with space constraints

Homozygote × homozygote

100% homozygote Can be applied for colony maintenance for no more than ten generations, production of a cohort when working with inbred or congenic GEMM, optimization of the production of mutants with space constraints, or homozygote mutant mice should be fertile and viable

Comments

ES cell was derived. Most of the ES cells used for targeted transgenesis are derived from 129 substrains and C57BL/6 substrains (Auerbach et al., 2000; Keskintepe et al., 2007). Similarly, the transgenic founder must be mated with the same inbred strain from which the embryos used for pronuclear injections were obtained. FVB/N and C57BL/6 animals are the most widely used inbred strains for efficient transgenesis (Auerbach et al., 2003). Once germ line transmission of the targeted mutation or the randomly integrated transgene is characterized, the new GEMM line must be maintained and expanded by breeding with the appropriate inbred strain to suit research needs.

(2) the genotype and controls needed; (3) the characteristics of the mutation; (4) the breeding schemes (heterozygote × heterozygote, homozygote × heterozygote, homozygote × homozygote; see Table 10); (5) the reproductive parameters; (6) the ratio of mice weaned to born; (7) the colony productivity (PEI); and (8) the ratio of non-productive breeders (Silver, 1995). To produce an estimate for an original established GEMM, one might use the strain characteristics of the parental strain used to generate the GEMM. Figure 6 shows how to properly size a mouse colony with two different cases studies.

Breeding schemes

Backcross: Breeding to congenicity

Table 10 summarizes the breeding schemes with genotype expected ratios. An example of a complex breeding scheme for the generation of a conditional knock-out of a gene of interest in a given tissue with the Cre/LoxP system is given in Figure 5 (for more insight on the Cre/LoxP system, see Birling et al., 2009; Gofflot et al., 2011).

Sizing the GEMM breeding colony

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From scientific and welfare standpoints, it is necessary to accurately calculate the number of breedings necessary to generate the number of animals needed for statistically relevant experimental studies. This calculation also helps to save space and costs. Sizing the breeding colonies should be based on: (1) the number, sex, and age range of mice required;

In many circumstances, such as (1) the non-availability of a given substrain, (2) the need to move a mutation from an unknown genetic background to a known one, (3) the non-suitability of an inbred strain for a given research field, or (4) the search for gene modifiers, it may be necessary to transfer the mutation by breeding to a different and defined genetic background. Figure 7 illustrates the breeding strategy called backcross to generate congenic lines. Transfer of the mutation from a given genetic background (donor strain) begins with a cross of the donor strain to the desired recipient inbred strain. This first cross is named outcross. The pups from this outcross are considered as the F1 generation and 50% of their genome originates from the donor strain and 50% from the host strain. After Current Protocols in Mouse Biology

Cre hemizygous mouse

TSP

Cre

tissue-specific promoter

50%

Floxed homozygous mouse

Ex1

Ex2

Ex3

Ex1

Ex2

Ex3

LoxP

LoxP

50%

Ex1

Ex2

Ex3

Ex1

Ex2

Ex3

25%

TSP

Cre

Ex1

Ex2

Ex3

Ex1

Ex2

Ex3

25%

Ex1

Ex2

Ex3

Ex1

Ex2

Ex3

25%

Ex1

Ex2

Ex3

Ex1

Ex2

Ex3

Ex1

Ex2

Ex3

Ex1

Ex2

Ex3

25% TSP

Cre

TSP

Cre

Ex1

Ex2

Ex3

Ex1

Ex2

Ex3

Ex1

Ex2

Ex3

Ex1

Ex2

Ex3

Figure 5 Complex breeding strategy: generation of a conditional, tissue-specific, gene knock-out in mice through a classical Cre-loxP strategy. The floxed mouse is obtained by introducing loxP sites around an essential exon of the gene of interest and using classical homologous recombination in ES cells. The Cre-expressing mouse, in which the Cre recombinase is under the control of a tissue-specific promoter (TSP), is usually a transgenic line produced by pronuclear microinjection. Both lines are established and maintained independently, the floxed line at the homozygous stage and the Cre line at the hemizygous stage. In a first step, the Cre line and the floxed homozygous line are intercrossed. Fifty percent of the offspring harbor the Cre transgene (highlighted in light orange on the figure) and a floxed allele of the gene of interest. Those animals are then once again crossed with the floxed homozygous line resulting in 25% of the offspring being homozygous for the floxed gene and hemizygous for the Cre. In those animals, excision of the floxed exon occurs only in Cre-expressing cells or tissue, while the gene of interest remains functional in other cells (highlighted in light blue on the figure). Such two-step breeding strategies allows the generation of experimental (Cre hemizygous, floxed gene homozygous) and appropriate control (floxed gene homozygous) animals at the same time with an intermediate frequency. Numbers correspond to the theoretical percentage of animals of a given genotype.

genotyping, F1 mice harboring the mutation or the transgene are bred back to the host inbred strain. This second cross and the subsequent ones are called backcrosses. Each generation after the F1 mice are denoted with an N number. The N2 generation retains only 25% of the donor strain. By N10, mice are classified as congenic as only 0.1% of the original donor genome remains. At this point, the GEMM line is considered identical to the recipient inbred strain, apart from the area around the gene modification which is still from the donor strain (an average size of 20 cM) (Silver, 1995). Congenic strain development by traditional breeding is simple but

the process is time consuming as it requires ∼3 years.

Speed congenic development To speed up the establishment of congenic strains, the marker-assisted breeding strategy, also known as “speed congenics” can be used to reduce the time required for deriving congenic strains (Markel et al., 1997; Wakeland et al., 1997). The breeding strategy is similar to the traditional method. However, heterozygous animals with the highest percentage of recipient genome are selected at each generation. The relative genome distribution is determined by PCR screening for polymorphic

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Colony characteristics Breeding scheme:

Heterozygote ⫻ Heterozygote (Het ⫻ Het)

Average liter size

5

Weaned to born ratio

80%

PEI

0.6

% of productive mating

85%

Case study no. 1: Number of breeding cages required to produce a phenotyping cohort of ten males and ten females homozygote and littermate controls Factors

Formulation

Calculation

Het ⫻ Het breeding scheme: 25% (1/4) of pups will be homozygote (genotype required)

Pups to produce ⫽ total number of experimental mice needed with the genotype required ⫻ 4

(10 ⫹ 10) ⫻ 4

Weaning-to-born ratio: 80% of pups born will be weaned

Total number of pups to produce ⫽ total number of experimental mice needed ⫻ 4/0.8

[(10 ⫹ 10) ⫻ 4]/0.8

Total number of mice to produce

100

Theoretical number of female breeders required (i.e., if 100% of the matings are productive)

Total number of mice to be produced/average litter size

100/5

Total number of female breeders required, taking into account the ratio of productive mating

Theoretical number of female breeders required/ratio of productive mating

20/0.85

Number of breeding pairs

24

Number of breeding trios

12

Case study no. 2: Number of breeding cages/week required to produce every 2 weeks five male homozygotes Factors

Formulation

Calculation

Het ⫻ Het breeding scheme: 12.5% (1/8) of pups will be male homozygotes for the genotype required

Pups to produce ⫽ total number of experimental mice with the genotype required ⫻ 8

5⫻8

Age requirement: 2-week age range

Pups to produce/week ⫽ pups to produce/week age range

(5 ⫻ 8)/2

Total number of mice to produce per week

20

Theoretical number of female breeders required/week

Total number of mice to be produced per week/PEI

20/0.6

Total number of female breeders required, taking into account the ratio of productive mating

Theoretical number of female breeders required/week/ratio of productive mating

(20/0.6)/0.85

Number of breeding pairs to maintain per week Number of breeding trios to maintain per week

39 18

Figure 6 Sizing a mouse colony appropriately: Two case studies showing how to calculate the number of breeding cages required for a GEMM colony. Calculations are based on the colony characteristics and reproductive performance, number of animals, age, genotype, and gender required (adapted from The Jackson Laboratory, 2009).

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DNA markers that span the entire genome. This technique reduces congenic strain development to ∼12 to 16 months depending on the robustness and intensiveness of the polymorphic analysis between the gene donor and recipient strains. A high-speed congenics strategy using early-stage male spermatid

injected into oocytes reduces the time to 6 months (Ogonuki et al., 2009).

Maintenance of a GEMM Line Appropriate maintenance strategies depend on (1) the type of strain (mixed, coisogenic, congenic), (2) the mutation inheritance mode, Current Protocols in Mouse Biology

donor strain 129S2/SvPas-Genetm/⫹

recipient strain C57BL/6J

X

outcross

X identification of Genetm/⫹ offspring: F1 generation

backcross 1 C57BL/6J

X identification of Genetm/⫹ offspring: N2 generation

backcross 2 C57BL/6J

X identification of Genetm/⫹ offspring: N3 generation

backcross 3 C57BL/6J

backcross 10 congenic GEMM B6.129-Genetm/⫹

identification of Genetm/⫹ offspring: N10 generation

Figure 7 Backcross breeding scheme to generate a congenic GEMM. In this example, a targeted mutation (Genetm ) is generated from 129S2/SvPas ES cells (genetic background represented here by a brown circle). To transfer the mutation onto the recipient C57BL/6J background (represented here by a black circle), a germline competent chimera (i.e., heterozygote carrier for a targeted mutation Genetm /+) is first crossed with the inbred C57BL/6J strain (outcross). The F1 progeny is screened for the mutation: 50% of the genome of the F1 offspring comes from the 129S2/SvPas strain and 50% from the C57BL/6J strain (depicted by brown and black hemispheres in the circle). Then, F1 heterozygous Genetm /+ are backcrossed to C57BL/6J. Heterozygous progeny of the F1 and subsequent generations are backcrossed to C57BL/6J until the tenth generation. N10 offspring are considered congenic, i.e., genetically identical to the C57BL/6J inbred strain except for the targeted gene locus and its surrounding region (represented here by the residual brown area within the black circle of N10).

(3) the genotype and the phenotype associated with the mutation (viability, fertility, maternal behavior, life span, penetrance), and (4) the necessary control mice require. Traditionally, the majority of GEMMs are derived from hybrid strains (e.g., B6;129 for targeted mutation, and B6SJL or B6D2 for transgenics). In this situation, continuous breeding between heterozygotes may be used. For phenotyping purposes, controls must be littermates from as many breeders as possible. However, for long-term maintenance, this mating scheme is not suitable, as animals will

differ genetically (except for the mutation) by inbreeding depression, i.e., decrease of vigor and/or yield (Wolfer et al., 2002). The second and best way to maintain GEMM lines is to backcross the mutation to an appropriate inbred strain depending on the research area of interest to develop a congenic GEMM strain. In this way heterozygote animals on a standardized genetic background are readily available to generate mutant and control animals. With such a congenic strain, it is acceptable to derive homozygous mutants and maintain the colony by intercrossing

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homozygotes as a separate stock. Mutant mouse production from a homozygous stock is more effective since unnecessary animals (wild-type and heterozygous mice) are not produced. For phenotyping studies, control animals can be selected from the parental inbred strain used to generate the congenic line. Homozygous and wild-type mice should be bred in the same facility. However, this homozygote × homozygote breeding scheme is not recommended for more than ten generations as it may lead to genetic drift with genetic differences between the congenics and the parental recipient inbred strain (Crusio et al., 2009). To prevent such a substrain divergence, backcrossing the colony strain to the parental inbred strain every five to ten generations is recommended (Crusio et al., 2009; The Jackson Laboratory, 2009). Fortunately, most new GEMMs are now coisogenic due to the development of robust C57BL/6 ES cells lines (Auerbach et al., 2000; Keskintepe, 2007; Pettitt et al., 2009). The C57BL/6 strain is one of the best characterized inbred strains of mice and is the reference strain for the mouse genome sequence (Mouse Genome Sequencing Consortium, 2002; International Mouse Knockout Consortium, 2007).

Long-term GEMM maintenance

Mouse Breeding and Colony Management

It may be preferred to maintain a strain for which there is no immediate need for expansion and phenotyping. The first option to maintain the line in an efficient and costeffective way is to have two or three active mating pairs (for a line with standard fecundity) with the appropriate breeding strategies depending on the type of strains (see Table 10). Keeping two distinct generations is advised until fertility of the newest generation is proven. If the strain is maintained by incross (homozygote × homozygote), remember to backcross the strain to prevent any development of a substrain. Monitoring of breeding records helps to track the strain history, to keep the line productive (see Management Tips and Troubleshooting), and to detect any deviance related to a genetic drift. The second option is to cryopreserve the line. Mouse embryos, sperm, or oocytes can be frozen (reviewed in Landel, 2010). Archiving is an effective alternative to managing colonies and enables resource savings as follows: (1) reduces resources needed such as animal housing space, husbandry, labor, and costs; (2) prevents strain loss due to disease or disaster; (3) facilitates mouse shipments as it is more con-

venient and secure to ship frozen samples than live animals; and (4) facilitates mice importation by avoiding the quarantine period. Cryopreservation is gaining popular recognition due to recent developments, with a fast, economical, and reliable method to cryopreserve and to recover animals from sperm of the C57BL/6 strain, from which most of the GEMMs are maintained (Ostermeier et al., 2008). Several public mouse strain repositories are available worldwide (http://www.fimre.org) and offer, to the international scientific community, a free mouse archiving service (after an evaluation process) and access to a large range of mouse strains (Hagn et al., 2007; Wilkinson et al. 2010).

Management Tips and Troubleshooting Several factors and aspects must be considered for efficient breeding programs including husbandry, animal care, and welfare considerations. Summarized below are some key elements for a successful program. Facility organization and operations considerations 1. Use cage card with a code color to distinguish each strain in a housing room; 2. Keep animals from a given colony altogether in the same housing room (breeders, weanings, and stock); 3. Use pre-printed cards to avoid transcription mistakes; 4. Avoid paper recording and use LIMS data recording systems to save time and to accurately record animal data; 5. Have adequately trained animal caretakers and technicians; 6. Have a dedicated animal technical team for each housing room or colony. Avoid staff turnover; 7. Maintain stable environmental parameters (hygrometry, light, temperature, see Factors Affecting Breeding Efficiency: Environmental Parameters) and record these parameters; 8. Be aware of any building renovations or construction operations. Efficient breeding programs 1. Limit noise and traffic in the mouse room as much as possible; 2. Handle cages and housing racks as gently as possible; 3. Select healthy animals without clinical abnormalities for breeding; 4. Obtain mice to expand the colony from an approved vendor or known source;

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Table 11 Breeding Troubleshooting Checklist

Question

Comment

Age and welfare of mice? Breeding life history? Source of mice? Background strain? Litter size consistent with the background?

Breeding performance is strain-specific

What is the breeding scheme and Genetic drift/sub strain divergence generation/sibling number of crosses? Genotype of breeders?

Transgene or mutation specific effect

Are the females getting pregnant? Are pups born?

Fetal resorption occurs in case of stress generated by human activities and noise

Are pups nursing? What is the genotype of surviving pups?

Transgene or mutation specific effect

Is there cannibalism?

Cannibalism is associated with environmental stress (noise, vibrations, etc.). Cannibalism is not unusual with C57BL/6 strains, especially for the first litter.

What about the other colonies in housing room? What is the overall health condition?

Overall colony productivity and health status of the mouse room are helpful information to identify the reason(s) of breeding issue.

5. Use mice during their optimal breeding life (from 6 weeks up to 8 months of age depending on strains); 6. Mate mice early (6 to 8 weeks of age); 7. Replace breeding mice before their reproductive performance declines as reproductive capacity diminishes with age; 8. Replace breeders if no litters or weaned pups are produced within 2 months; 9. Use proven breeder males whenever possible or keep them experienced (once a month) before starting a breeding program; 10. To set up matings, especially for cohort production of age-matched animals, isolate males a couple of days before mating in a new cage, and then introduce the females to the male cage; 11. Check for vaginal plug during the morning to verify if mating has occurred; 12. Leave male continuously with female(s) to take advantage of the post-partum estrus. Females can get pregnant within 24 hr after delivery; 13. Avoid handling or disturbing pregnant females several days before and after giving birth;

14. Provide breeding mice with nesting material to optimize maternal environment; 15. Keep a piece of soiled nest and bedding when changing the cage to minimize environment disturbance for parents and offspring; 16. Keep accurate records (pedigree, genotype); 17. Monitor the PEI closely to detect early any deviance and to take corrective measures. Troubleshooting breeding problems Reproductive issues are not unusual when working with mouse mutants, and it is challenging to keep up efficient breeding programs as they rely on a combination of parameters, including environmental and husbandry conditions (see Factors Affecting Breeding Efficiency: Environmental Parameters), health status (see Factors Affecting Breeding Efficiency: Sanitary Status), and genetic modifications (see Colony Management). For this purpose, a check list of questions (Table 11) is helpful to identify the parameters affecting the breeding efficiency.

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SPECIFIC BREEDING METHODS: ASSISTED REPRODUCTION TECHNIQUES Many techniques are available to support GEMMs colony management by improving mouse production or dealing with breeding issues related to their genetic modifications. Some of them are technically demanding and require dedicated equipment and trained staff. The most commonly used techniques for breeding GEMMs are described below. For a comprehensive description, see Nagy et al. (2003).

Timed Matings It may be necessary to know the time of mating to monitor pregnancies for research on embryonic development, reproductive phenotyping, or for dealing with breeding issues related to reproductive performance. This is commonly done by placing a male and female(s) together just prior to the dark phase at the end of the working day. The morning of the following day, the presence of a white and waxy copulatory plug in the vagina of the female indicates that mating occurred. Because mating usually occurs near the midpoint of the dark phase, the day of mating indicated by the presence of the vaginal plug is referred to as day 0.5 of embryonic development. Presence of the plug should be checked early in the morning before its disappearance. A vaginal plug does not necessarily mean pregnancy. This can be confirmed at a later time by palpation after the 10.5 days stage of pregnancy. Because females are only receptive to males while they are in estrus (i.e., ovulating; the female mouse has a 4- to 5-day cycle), only

20% to 25% of the breedings set up will mate. This must be taken into account when planning timed pregnancies. If it is necessary to maximize the number of pregnant females at the same stage, then only females in proestrus cycling stage in the afternoon should be placed with male(s). The stage of the estrus cycle is determined by examining the external genitalia of the female. Roughly, up to 90% of females in proestrus stage will go into estrus and mate (Champlin et al., 1973). Note that females that are group-housed may have their estrus cycle suppressed because of the LeeBoot effect (Lee and van der Boot, 1955). Estrus cycles can be resumed simultaneously by exposure to male urine by placing dirty bedding in the cage for at least 3 days (Whitten, 1959).

Fostering Fostering mouse pups is a method commonly used to save valuable GEMMs when mothers are not able to care for their pups for various reasons including nurturing defects, lactation defects, or cannibalism. Basically, pups to be fostered are removed from their birth dam (donor strain) near the time of birth and are placed into the cage of another lactating female (recipient strain). This means that a colony of foster mothers should be maintained in the breeding facility. Typically, the recipient strain used is an outbred mouse stock such as CD1 or Swiss, as they are good mothers. The white coat color of these foster mothers helps to identify the fostered GEMM pups, which usually have a different coat color. The foster mother should be suckling its own litter that is about the same age as the pups to be

Table 12 Step-by-Step Protocol for Efficient Fostering

Mouse Breeding and Colony Management

1

Transfer the lactating foster mother into a holding cage.

2

Remove the appropriate number of pups according to the number of pups to be fostered. Pay attention not to exceed the size of the recipient litter. Keep intact the nesting material.

3

Have the foster mother urinate on the pups to be fostered.

4

Transfer the pups and mix them with the recipient pups (if applicable). Wipe them with dirty bedding and leave nesting material to give them the same scent as the recipient’s pups.

5

Bring back the foster mother to her cage (separation time should be as short as possible).

6

Return cage to the rack, leave it, and check it carefully after 1 hr to be sure the foster mother has accepted the new pups.

7

Check periodically on the following days that the mother is taking care of the pups.

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fostered. Additionally, the number of pups to be fostered should not exceed the number of pups of the natural litter of the recipient mother to not affect the mother’s milk supply production. A straightforward protocol is described in Table 12.

In Vitro Fertilization (IVF) IVF involves fertilization of oocytes with capacitated sperm in a Petri dish. IVF is by far the most used assisted reproductive technique, as it allows the production of large numbers of embryos for several purposes. First, IVF is used to rescue hypofertile lines by overcoming breeding failures, regardless of the cause of limitations, including age, illness, or mutation phenotype. Second, IVF is also used to support GEMM breeding programs for fast colony establishment or expansion since sperm from one single animal can be used to fertilize a large number of oocytes from multiple females. IVF is used routinely to generate agematched phenotyping cohorts at the Institut Clinique de la Souris. This application combined with sperm freezing is a very convenient GEMM colony management tool as one can rapidly and efficiently set up a colony (typically in one shot starting from archived material). An IVF success rate is strain specific, but due to the recent methodological advances, the IVF success rate with thawed sperm from several inbred strains (including C57BL/6 strains, the most popular inbred strain for GEMMs) has been enormously improved (Ostermeier et al., 2008). Additionally, IVF can also shorten the timelines for (1) backcrossing onto an inbred strain (Ogonuki et al., 2009), and (2) importation of strains. Finally, the IVF is advantageous for cryopreservation. Two-cell embryos produced by IVF may be directly frozen to archive the GEMM strain. This process, known as Cryo-IVF, shortens the timeline for safely archiving a line with several hundred embryos and minimizes the workload and housing space required for cryopreservation of embryos. Several protocols are published and differ either with the type of fertilization media, the method to collect and handle sperm and oocytes, or the timetable. An IVF protocol adapted from Sztein et al. (2000) is available at the following link: http://www.emmanet.org and is used at the Institut Clinique de la Souris. The IVF method by itself appears straightforward, but IVF requires highly trained and organized personnel, significant technical efforts,

and specific handling of sperm and oocytes to set up successful sessions.

Intra-Cytoplasmic Sperm Injection (ICSI) The ICSI is an assisted IVF, which consists of the injection of a single spermatozoon into the cytoplasm of an oocyte. The main advantage of this technique is the use of spermatozoa that can be non-motile or even dead. The only requirement is the preservation of the genetic integrity. Therefore, ICSI is an attractive method to rescue lines with male infertility due to sperm hypomotility. ICSI should also be considered when the thawing of frozen sperm results in low motility rates, which has been known to be the case for some specific inbred strains (Kawase et al., 2001; Szczygiel et al., 2002). This method is technically challenging and requires specific skills and costly equipment. ICSI is not a routine method and must be used only when other methods are ineffective.

Ovarian Transplantation (OT) OT is an alternative method to save or rescue colonies from valuable females that are unproductive for various reasons including age, poor health, and phenotype-related issues with pregnancy, parturition, or mothering. Ovaries from newborn pups or adults are transplanted into histocompatible or immunodeficient recipient ovarectomized females. After a period of recovery, transplanted females are mated with males to produce progeny. OT is helpful and may be the only solution to maintain valuable GEMM lines such as transgenic female founders or other mutants having a shortened life span. Additionally, ovaries can be cryopreserved, thus providing valuable addition to other cryopreservation techniques. The transplantation technique is also used to recover mouse strains from cryopreserved ovaries (Sztein et al., 1998). OT is technically simple but requires surgery skills, aseptic conditions to perform the transplantation, appropriate analgesia, and postoperative care to insure the recovery of the animal. A detailed procedure is described by Nagy et al. (2003).

CONCLUSION Several aspects have to be taken into account to breed and manage efficiently a mouse colony. The physiological data for mouse reproduction should be known, especially the gestation time (21 days), the average litter size (2 to 12 pups), and the reproductive life span (average 8 months). One has to keep in mind

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that these data differ between inbred strains and outbred stocks. Knowledge of the environmental parameters in the facility where the animals are housed is also important for optimizing the breeding, especially the housing microenvironment (cage type, bedding, diet and nesting material) and the macroenvironment (air renewal, temperature, hygrometry and light cycle). Good communication between the research groups and the animal facility management is crucial to identify any unwanted modification of one or more of these factors and to correct them as soon as possible in order to minimize the impact on breeding. Similarly, one should be aware of the sanitary status of the facility, as the presence of given bacteria or viruses can affect the breeding of the strain of interest. Prior to the start of managing a mouse colony, the mating type (duo, trio, harem type) and the breeding scheme (depending of the genotype of the desired experimental animals) must first be defined to follow and establish the way the animals will be identified and genotyped. The question of the genetic background of the line should also be raised, and backcrosses may be set up to work with the appropriate background for the study. All the breeding data should be recorded, at least at the level of the cage. Efficient colony management requires, in most cases, the use of an animal colony management software where all the data regarding the lines will be recorded and easily accessed. Such software may also be used as a communication tool between the research groups and the animal facility team (e.g., for specific work orders). In the case of breeding issues, assisted reproduction techniques can be used, going from simple techniques such as time mating and fostering to more advanced ones such as IVF or ICSI, the latter requiring specific technical skills. Breeding may seem like a trivial factor but different skills are necessary for efficient mouse breeding and colony management; therefore, none of the abovementioned should be neglected.

ACKNOWLEDGEMENTS

Mouse Breeding and Colony Management

Gis`ele Ferrand and Xavier Warot thank their colleagues at the Center of PhenoGenomics of the Ecole Polytechnique F´ed´erale de Lausanne for copious discussions, namely Friedrich Beermann, Rapha¨el Doenlen, Emilie Gesina, Marcel Gyger, and Charles Thomas. Abdelkader Ayadi and Xavier Warot wish to express their deep gratitude to Gary R. No-

vak (Baltimore, Maryland) for critical review and comments on the manuscript.

LITERATURE CITED Auerbach, W., Dunmore, J., Fairchild-Huntress, V., Fang, Q., Auerbach, A., Huszar, D., and Joyner, A. 2000. Establishment and chimera analysis of 129/SvEv- and C57BL/6-derived mouse embryonic stem cell lines. Biotechniques 29:10241032. Auerbach, A., Norinsky, R., Ho, W., Losos, K., Guo, Q., Chatterjee, S., and Joyner, A. 2003. Strain-dependent differences in the efficiency of transgenic mouse production. Transgenic. Res. 12:59-69. Birling, M.C., Gofflot, F., and Warot, X. 2009. Site-specific recombinases for manipulation of the mouse genome. In Transgenesis Techniques, Methods in Molecular Biology (E.J. Cartwright, ed.) pp. 245-263. Humana Press, Totowa, New Jersey. BVAAWF/FRAME/RSPCA/UFAW, Joint Working Group on Refinement. 2003. Refinement and reduction in production of genetically modified mice. Lab. Anim. 37:S1-S49. Castelhano-Carlos, M.J. and Baumans, V. 2010. Noise and light in the vivarium. ALN World 3:1217. Castelhano-Carlos, M., Sousa, N., Ohl, F., and Baumans, V. 2010. Identification methods in newborn C57BL/6 mice: A developmental and behavioural evaluation. Lab. Anim. 44:88-103. Champlin, A., Dorr, D., and Gates, A. 1973. Determining the stage of the estrous cycle in the mouse by the appearance of the vagina. Biol. Reprod. 8:491-494. Crusio, W., Goldowitz, D., Holmes, A., and Wolfer, D. 2009. Standards for the publication of mouse mutant studies. Genes Brain Behav. 8:1-4. Davisson, M.T. and Linder, C.L. 2004. Historical foundations. In The Laboratory Mouse (H.J. Hedrich and G. Bullock, eds.) pp. 15-24. Elsevier Academic Press, Boston, Massachusetts. Doetschman, T. 2009. Influence of genetic background on genetically engineered mouse phenotypes. Methods Mol. Biol. 530:423-433. Donnelly, C., McFarland, M., Ames, A., Sundberg, B., Springer, D., Blauth, P., and Bult, C. 2010. JAX Colony Management System (JCMS): An extensible colony and phenotype data management system. Mamm. Genome 21:205-215. Festing, M.F.W. and Peters, A.G. 1999. Animal production and breeding methods. In The UFAW Handbook on the Care and Management of Laboratory Animals, Vol. 1, 7th ed. (T. Poole, ed.) pp. 28-44. Blackwell Science Ltd, Oxford. Gofflot, F., Wendling, O., Chartoire, N., Birling, M.C., Warot, X., and Auwerx, J. 2011. Characterization and validation of Cre-driver mouse lines. Curr. Protoc. Mouse Biol. 1:1-15. Gu´enet, J-L. and Benavides, F.J. 2011. Mouse strains and genetic nomenclature. Curr. Protoc. Mouse Biol. 1:213-238.

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Hagn, M., Marschall, S., and Hrab´e de Angelis, M. 2007. EMMA-the European mouse mutant archive. Brief Funct. Genomic Proteomic 6:186192. Hardy, P. 2004. Gnotobiology and breeding techniques. In The Laboratory Mouse (H.J. Hedrich and G. Bullock, eds.) pp. 409-433. Elsevier Academic Press, Boston, Massachusetts. Havenaar, R., Meijer, J.C., Morton, D.B., RitskesHoitinga, J., and Zwart, P. 1993. Biology and husbandry of laboratory animals. In Principles of Laboratory Animal Science (L.F.M. van Zutphen, V. Baumans and A.C. Beynene, eds.) pp. 17-74. Elsevier Science Publishers, Maarssen, Netherlands. The International Mouse Knockout Consortium. 2007. A mouse for all reasons. Cell 128:9-13. The Jackson Laboratory. 2009. Breeding Strategies for Maintaining Colonies of Laboratory Mice Manual. Available online at: http://jaxmice.jax.org/manual/index.html. Kawase, Y., Iwata, T., Toyoda, Y., Wakayama, T., Yanagimachi, R., and Suzuki, H. 2001. Comparison of intracytoplasmic sperm injection for inbred and hybrid mice. Mol. Reprod. Dev. 60:7478. Keskintepe, L., Norris, K., Pacholczyk, G., Dederscheck, S., and Eroglu, A. 2007. Derivation and comparison of C57BL/6 embryonic stem cells to a widely used 129 embryonic stem cell line. Transgenic Res. 16:751-758. Landel, C. 2010. Cryopreservation of mouse gametes and embryos. Methods Enzymol. 476:85105. Le Calvez, S., Perron-Lepage, M., and Burnett, R. 2006. Subcutaneous microchip-associated tumours in B6C3F1 mice: A retrospective study to attempt to determine their histogenesis. Exp. Toxicol. Pathol. 57:255-265. Lee, S. and van der Boot, L.M. 1955. Spontaneous pseudopregnancy in mice. Acta Physiol. Pharmacol. Neerl. 4:442-443. Lemken, B. 2010. Rethinking global water quality standards. ALN World 3:12-16. Markel, P., Shu, P., Ebeling, C., Carlson, G., Nagle, D., Smutko, J., and Moore, K. 1997. Theoretical and empirical issues for marker-assisted breeding of congenic mouse strains. Nat. Genet. 17:280-284. Matzuk, M. and Lamb, D. 2008. The biology of infertility: Research advances and clinical challenges. Nat. Med. 14:1197-1213. Montoliu, L. and Whitelaw, C. 2010. Using standard nomenclature to adequately name transgenes, knockout gene alleles and any mutation associated to a genetically modified mouse strain. Transgenic Res. 19:587-594.

embryo. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York. National Research Council. 1991. Infectious diseases of mice and rats. Committee on infectious diseases in mice and rats. Institute of Laboratory Animal Resources, Commission of Life Sciences. National Academies Press, Washington D.C. National Research Council, 2010. Guide for the Care and Use of Laboratory Animals. National Academies Press, Washington D.C. Naz, R., Engle, A., and None, R. 2009. Gene knockouts that affect male fertility: Novel targets for contraception. Front. Biosci. 14:39944007. Nicklas, W., Baneux, P., Boot, R., Decelle, T., Deeny, A.A., Fumanelli, M., and Illgen-Wilcke, B. 2002. Recommendations for the health monitoring of rodent and rabbit colonies in breeding and experimental units. Lab. Anim. 36:2042. Ogonuki, N., Inoue, K., Hirose, M., Miura, I., Mochida, K., Sato, T., Mise, N., Mekada, K., Yoshiki, A., Abe, K., Kurihara, H., Wakana, S., and Ogura, A. 2009. A high-speed congenic strategy using first-wave male germ cells. PLoS One 4:e4943. Ostermeier, G., Wiles, M., Farley, J., and Taft, R. 2008. Conserving, distributing and managing genetically modified mouse lines by sperm cryopreservation. PLoS One 3:e2792. Pettitt, S, Liang, Q., Rairdan, X., Moran, J., Prosser, H., Beier, D., Lloyd, K., Bradley, A., and Skarnes, W. 2009. Agouti C57BL/6N embryonic stem cells for mouse genetic resources. Nat. Methods 6:493-495. R¨ulicke, T., Montagutelli, X., Pintado, B., Thon, R., Hedrich, H., and Group, F.W. 2007. FELASA guidelines for the production and nomenclature of transgenic rodents. Lab. Anim. 41:301-311. Russell, W. and Burch, R. 1959. The Principle of Humane Experimental Technique. Metheun & Co. Ltd, London. Schaefer, D., Asner, I., Seifert, B., B¨urki, K., and Cinelli, P. 2010. Analysis of physiological and behavioural parameters in mice after toe clipping as newborns. Lab. Anim. 44:7-13. Silver, L. 1995. Mouse Genetics: Concepts and Applications. Oxford University Press, New York. Smith, E., Stockwell, J.D., Schweitzer, I., Langley, S.H., and Smith, A.L. 2004. Evaluation of cage micro-environment of mice housed on various types of bedding materials. Contemp. Top. Lab. Anim. Sci. 43:12-17.

Mouse Genome Sequencing Consortium. 2002. Initial sequencing and comparative analysis of the mouse genome. Nature 420:520-562.

Szczygiel, M., Kusakabe, H., Yanagimachi, R., and Whittingham, D. 2002. Intracytoplasmic sperm injection is more efficient than in vitro fertilization for generating mouse embryos from cryopreserved spermatozoa. Biol. Reprod. 67:12781284.

Nagy, A., Gertsenstein, M., Vintersten, K., and Behringer, R. 2003. Manipulating the mouse

Sztein, J., Sweet, H., Farley, J., and Mobraaten, L. 1998. Cryopreservation and orthotopic

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transplantation of mouse ovaries: New approach in gamete banking. Biol. Reprod. 58:10711074.

Whitten, W.K. 1959. Occurrence of anoestrus in mice caged in groups. J. Endocrinol. 18:102107.

Sztein, J., Farley, J., and Mobraaten, L. 2000. In vitro fertilization with cryopreserved inbred mouse sperm. Biol. Reprod. 63:1774-1780.

Wilkinson, P., Sengerova, J., Matteoni, R., Chen, C., Soulat, G., Ureta-Vidal, A., Fessele, S., Hagn, M., Massimi, M., Pickford, K., Butler, R., Marschall, S., Mallon, A., Pickard, A., Raspa, M., Scavizzi, F., Fray, M., Larrigaldie, V., Leyritz, J., Birney, E., Tocchini-Valentini, G., Brown, S., Herault, Y., Montoliu, L., Hrab´e de Angelis, M., and Smedley, D. 2010. EMMAmouse mutant resources for the international scientific community. Nucleic Acids Res. 38:D570D576.

van Gassen, K., Hessel, E., Ramakers, G., Notenboom, R., Wolterink-Donselaar, I., Brakkee, J., Godschalk, T., Qiao, X., Spruijt, B., van Nieuwenhuizen, O., and de Graan, P. 2008. Characterization of febrile seizures and febrile seizure susceptibility in mouse inbred strains. Genes Brain Behav. 7:578-586. Voipio, H.-M., Tsai, P.-P., Brandstetter, H., Gyger, M., Hackbarth, H., Kornerup Hansen, A., and Krohn, T. 2010. Housing and care of laboratory animals. In The COST Manual of Laboratory Animal Care and Use, Refinement, Reduction, and Research (B. Howard, T. Nevalainen, and G. Perretta, eds.) pp. 29-73. CRC Press, Boca Raton, Fla.

Wolfer, D., Crusio, W., and Lipp, H. 2002. Knockout mice: Simple solutions to the problems of genetic background and flanking genes. Trends Neurosci. 25:336-340. Wooten, R.M., Ma, Y., Yoder, R.A., Brown, J.P., Wies, J.H., Zachary, J.F., Kirschning, C.J., and Weis, J.J. 2002. Toll-like receptor 2 is required for innate, but not acquired, host defense to Borrelia burgdorferi. J. Immunol. 168:348-355.

Wakeland, E., Morel, L., Achey, K., Yui, M., and Longmate, J. 1997. Speed congenics: A classic technique in the fast lane (relatively speaking). Immunol. Today 18:472-477.

Internet Ressources

White, W.J. 2007. Management and design: Breeding facilities. In The Mouse in Biomedical Research (J.G. Fox ed.) pp. 235-270. Academic Press, Amsterdam.

http://www.jax.org Web site of the Jackson Laboratory. A number of online useful resources on mouse reproduction freely available (books, manuals, posters).

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Overview of the Measurement of Lipids and Lipoproteins in Mice Anne Tailleux1,2,3,4 and Bart Staels1,2,3,4 1

Universit´e Lille Nord de France, Lille, France Inserm, U1011, Lille, France 3 UDSL, Lille, France 4 Institut Pasteur de Lille, Lille, France 2

ABSTRACT Experimental mouse models are widely used for preclinical research on dyslipidemia, atherosclerosis, and cardiometabolic diseases. This unit reports the most commonly used biochemical analysis methods available to determine the lipid and lipoprotein phenotype in the mouse. The discussed methods include: the qualitative and quantitative analysis of lipids, apolipoproteins, and lipoproteins (with a specific emphasis on species-specificities of mice and humans), and the activity assay of major enzymes involved in lipoprotein remodeling (LCAT, PLTP, and LPL). The unit also discusses the most frequently used functional tests to analyze lipid/lipoprotein metabolism in vivo, including triglyceride metabolism, reverse cholesterol transport, intestinal C 2011 by John Wiley & lipid absorption, and secretion. Curr. Protoc. Mouse Biol. 1:265-277  Sons, Inc. Keywords: lipids r lipoproteins r apolipoproteins r mouse

INTRODUCTION Lipoproteins are soluble complexes transporting proteins (apolipoproteins or apos) and lipids in the circulatory system. Lipoproteins are mainly synthesized in the liver and the intestine. In the circulatory system, lipoproteins are in a state of constant dynamic remodeling, changing in composition and physical structure as peripheral tissues take up their various components before return of the lipoprotein remnants to the liver. Lipoprotein components undergo enzymatic reactions, such as facilitated and spontaneous lipid transfers, transfer of soluble apolipoproteins, and conformational changes of the apolipoproteins in response to the compositional changes. Finally, lipoproteins are taken up and catabolized in the liver and peripheral tissues via receptor-mediated endocytosis and other mechanisms. The most abundant lipid constituents are triacylglycerols (TG), free cholesterol (FC), cholesteryl esters (CE), and phospholipids (PL) (especially phosphatidylcholine), though fat-soluble vitamins and antioxidants are also transported. Free (unesterified) fatty acids (FFAs) are bound in plasma to the protein albumin by hydrophobic forces. Lipoproteins can be classified based on chemical or physical parameters, such as size, density, or electrophoretic mobility. The main

lipoproteins are: chylomicrons (CM), verylow-density lipoproteins (VLDL), low-density lipoproteins (LDL), and high-density lipoproteins (HDL), based on the relative densities of the particles. An alternative nomenclature is based on the relative electrophoretic mobility on agarose gels. Thus, α-, pre-β-, and βlipoproteins correspond to HDL, VLDL, and LDL, respectively (see Table 1). Lipoprotein classes differ with respect to their clinical significance and association with cardiovascular disease risk. Indeed, LDL are proatherogenic, whereas HDL generally are antiatherogenic. Moreover, plasma TG concentrations represent an independent risk factor for cardiovascular diseases. The use of mice has several advantages over other species, including easy breeding, short generation time, the availability of extensive genetic information on numerous inbred strains, and ease of genetic manipulations [transgenesis, knock-out (KO), knock-in, and conditional KO]. Moreover, the small size of mice allows the analysis of the efficacy of pharmacological compounds using only low amounts of drugs before the need for largescale production, constituting interesting pharmacological models (Fievet et al., 2007). There are, however, also a number of drawbacks to the use of the mouse as an

Current Protocols in Mouse Biology 1: 265-277, June 2011 Published online June 2011 in Wiley Online Library (wileyonlinelibrary.com). DOI: 10.1002/9780470942390.mo110001 C 2011 John Wiley & Sons, Inc. Copyright 

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Table 1 Main Physicochemical Properties of Lipoproteinsa

Density

Electrophoretic mobility

Diameter (nm)

Chylomicrons

<0,94

Origin

100-1000

1,000,000

B48

b

0.94-1.006

Pre-β

30-80

7,500

B100, B48, Cs

VLDL b

Molecular Main weight (kDa) apolipoproteins

1.006-1.019

Slow pre-β

25-35

4,000

B100, E

b

1.019-1.063

β

15-25

2,000

B100

b

1.063-1.210

α

6-14

200-400

AI, AII

IDL

LDL

HDL

a Lipoproteins are classified according to different physicochemical characteristics that allow their separation, such as

hydrated density (ultracentrifugation), electrophoretic mobility (electrophoresis on cellulose acetate), size (gel-filtration chromatography), molecular weight (nondenaturing electrophoresis), as well as apolipoprotein composition (affinity chromatography). These properties are similar in mice and in humans, except for the presence of apoB48 in lipoproteins, which are produced by the liver only in the mouse. b Abbreviations: VLDL, very low-density lipoproteins; IDL, intermediary density lipoproteins; LDL, low-density lipoproteins; HDL, high-density lipoproteins.

Overview of the Measurement of Lipids and Lipoproteins in Mice

experimental model in the research of lipid and lipoprotein metabolism. First, lipid and lipoprotein metabolism is dissimilar between mice and humans. In the mouse, most of the cholesterol is carried by HDL instead of LDL (the major carrier of cholesterol in humans). This is partly due to the absence, in mice, of cholesteryl ester transfer protein (CETP), an enzyme exchanging TG/CE between lipoproteins. Second, plasmafree glycerol concentrations are very high in the mouse, as compared to humans, and impacts on plasma TG measurement depending on the method employed. Third, the structure of most apolipoproteins differs between mouse and humans, which requires the development of specific immunoreactants. Fourth, the regulation of genes encoding proteins involved in lipid and lipoprotein metabolism is not identical between humans and mice, and thus data obtained in the mouse are not always directly relevant to humans. Finally, the mouse is highly resistant to atherosclerosis and does not develop atherosclerotic lesions spontaneously. Indeed, most current mouse models of atherosclerosis are based on perturbations of lipoprotein metabolism through dietary and/or genetic manipulations (Tailleux et al., 2003; Fievet et al., 2007). This unit reports the main analysis methods available to determine the lipid and lipoprotein phenotype in mice. This encompasses the qualitative and quantitative analysis of lipids and lipoproteins with emphasis on species differences, the activity assay of the main enzymes involved in lipoprotein remodeling, and the main functional tests available to assess

lipid/lipoprotein metabolism in vivo to more dynamically explore lipid metabolism.

QUANTITATIVE AND QUALITATIVE ANALYSIS OF CIRCULATING LIPIDS AND LIPOPROTEINS Blood sample collection and processing Circulating lipid and/or lipoprotein analysis is generally performed on samples collected after a 4-hour fasting period during the day (8 am to 12 am, or 9 am to 1 pm). It is important to note that this procedure consists not only of taking the food pellets out of the cage, but also in changing the sawdust to avoid that pellets which could have fallen into the cage are eaten. Blood is collected by retro-orbital venipuncture in anesthetized animals, or by intracardiac puncture, or by tail vein cutting. The main drawback of the latter method is the low volume of blood collected, which restricts analyses. The number of repetitive sample collections, as well as the volume of samples, are tightly controlled by ethical committees in different countries. Blood can be collected in tubes containing an anticoagulant, such as EDTA, heparin, or citrate (to obtain plasma), or none (to obtain serum). In all cases, blood samples are immediately placed on ice and rapidly centrifuged to sediment the erythrocytes (generally 15 min at 2100 × g, 4◦ C). After centrifugation, plasma/serum is carefully collected with a pipet, placed in a tube, and stored at 4◦ C until analysis (if performed within 3 days). Samples can also be stored at −20◦ C.

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However, it is recommended to avoid freezing hypertriglyceridemic samples. It is preferred to store the samples at +4◦ C for analysis within 3 days. Lipid and lipoprotein analyses can be performed both on plasma and serum samples.

Lipid and lipoprotein analysis Numerous quantitative and qualitative lipid and lipoprotein analysis methods are performed in mice using the same techniques as those used for their analysis in humans. However, the difficulty resides in the low volumes of samples collected in the mouse (maximum of a few hundred microliters), as compared to humans. So the main problem for biochemical analysis of mouse samples consists in miniaturizing the methods used for humans to low sample volumes. Lipids that are directly measurable in plasma include total cholesterol (TC), triglycerides (TG), free cholesterol (FC), phospholipids (PL), and free fatty acids (FFA). These are measured using ready-to-use enzymatic colorimetric kits available for biological analysis in humans. Briefly, the lipid reacts with a specific enzyme leading to the formation of a product quantifiable with a spectrophotometer at the indicated optical density. An internal control will allow validation of the assay. This internal control is either provided by the suppliers or homemade as a pool of plasma from normolipidemic mice. To adapt the ready-to-use kits to the low sample volume, assays are performed either manually in 96-well microplates or in automated analytical instruments by appropriate adaptation of sample-to-reactant volume ratio. The term “total cholesterol” refers to cholesterol associated with all fractions of lipoproteins. Due to the clinical significance of cholesterol associated with the different lipoprotein fractions, it is important to specifically quantify cholesterol in the HDL fraction (HDL-C, i.e., non-atherogenic cholesterol) and cholesterol in all other lipoprotein fractions (non-HDL-cholesterol, i.e., atherogenic cholesterol). To measure HDL-C specifically, the sample is incubated with a reactant, such as phosphotungstic acid/magnesium, which precipitates apolipoprotein-B-containing lipoproteins (all lipoprotein fractions except HDL). After centrifugation, cholesterol is measured in the supernatant, which contains only HDL. Non-HDL-C is obtained by subtraction of HDL-C from TC.

It is noteworthy that the plasma glycerol concentration is much higher in the mouse than in humans, and this may have an impact on the values of TG concentrations measured in the mouse. The triglyceride molecule consists of a glycerol esterified to three fatty acids, and the triglyceride assay consists in measuring free glycerol after hydrolysis by a lipase. Thus, in mouse samples, measuring TGs (called in this context “total triglycerides”) consists of measuring free glycerol in addition to “true” TG. To take this particularity into account, it is possible to measure free glycerol, once again using colorimetric ready-to-use kits, and then the true TG value is obtained by subtracting the free glycerol value from the total TGs value. An alternative is the measurement of TG after gel-filtration chromatography (see below). True TGs are not measured systematically in all mouse experiments, but specifically in mice subjected to metabolic conditions that tend to modify glycerol concentrations, e.g., after fasting when intense lipolysis occurs in adipose tissue that releases glycerol. Free cholesterol and TC concentrations are used to calculate two other parameters. Esterified cholesterol (EC) is obtained by subtraction of FC from TC. Cholesteryl esters (CE) are calculated as 1.68× (TC-FC), 1.68 being the mean molecular mass of a free fatty acid. The methods mentioned above allow the discrimination of HDL only from non-HDLcholesterol concentrations. However, it is often necessary to further detail the cholesterol and triglyceride distribution in the different lipoprotein fractions. For that, lipoproteins are separated by gel-filtration chromatography according to their size (VLDL, LDL, and HDL). It is noteworthy that chylomicrons, the largest lipoproteins, are generally absent because samples are collected in the fasting state. Two different systems are available: 1. In the first one, the chromatograph effluent is immediately and continuously mixed with cholesterol or TG assay reactant, and incubated at the appropriate conditions of temperature and duration to allow the enzymatic reaction to occur. The OD measured at the appropriate wavelength is automatically converted into a graphic signal representing the lipid distribution profile. The area under each peak is proportional to the lipid concentration in the respective lipoprotein fraction. For example, the cholesterol concentration in a lipoprotein fraction is calculated as (area of

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A 0.035 0.03

HDL

A.U.

0.025 0.02 0.015 VLDL

0.01

LDL

0.005 0 20

30

40

50

60 70 Elution time (min)

80

90

100

B Lipoprotein fraction

VLDL

LDL

HDL

Total

Area under peak (A.U.)

2,132,304

5,897,279

17,300,096

25,329,679

Area under peak (% of total)

8.4

22.3

68.3

100

Cholesterol concentration (mg/dl)

8.0

22.3

65.9

95.2

Figure 1 Example of a cholesterol distribution profile in the different lipoprotein fractions separated by gel-filtration chromatography according to their size using plasma from a C57BL6 mouse (A. Tailleux and B. Staels, unpub. observ.). Cholesterol is measured in the total plasma and continuously in the eluate. (A) chromatograph; (B) calculation of VLDL-C, LDL-C, and HDL-C plasma concentrations.

Overview of the Measurement of Lipids and Lipoproteins in Mice

the peak corresponding to the lipoprotein fraction/sum of peak areas) × plasma cholesterol concentration (see Fig. 1). This method can be applied to each individual sample with a small volume (less than 50 μl for one parameter, depending on the apparatus) and has the potential of being highly automatizable. For example, a number of samples could be loaded in the evening; samples are then automatically analyzed during the night, and results are available the next day. 2. A variant is to use a gel-filtration chromatography system connected to a fraction collector. The column is loaded with the sample, the eluted fractions are then collected into tubes, and lipid concentrations are measured in each tube. Lipid profiles are determined by plotting lipid concentration in function of the eluted fraction. Although this is poorly applicable to each individual plasma sample, as

the volume required is at least 200 μl per sample, it can be recommended for analysis of pooled plasma from different animals. Moreover, this method allows one to isolate and collect each lipoprotein fraction, and to measure lipids other than cholesterol and TG, in addition to protein concentrations. Thus, this method allows one to determine the mass composition of each lipoprotein fraction or to perform additional qualitative analysis on the lipoprotein fractions. For example, a lipoprotein fraction can be loaded on an SDS-PAGE to analyze its apolipoprotein composition. It is noteworthy that gel-filtration chromatography separates glycerol, a very low-mass molecule compared to lipoproteins, from the other fractions. Thus, TG profiles obtained in these conditions reflect “true TG” concentrations. In addition to gel-filtration chromatography, flotation ultracentrifugation is another

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technique to isolate lipoprotein fractions. Briefly, appropriate KBr quantities are added to the plasma to adjust its density followed by centrifugation. Either a simple centrifugation at density <1.21 g/ml to obtain all lipoproteins, or sequential ultracentrifugations to obtain each lipoprotein fraction are performed. Generally, the following densities are used: VLDL (density <1.006 g/ml), LDL (density =1.019 − 1.063 g/ml), and HDL (density =1.063 − 1.21 g/ml). Centrifugation is performed either in large tubes (for 24 hr at 107,328 × g, 15◦ C for VLDL, or 4◦ C for others) or in small tubes more suitable for small plasma samples (for 2 hr at 5,590 × g, 15◦ C for VLDL, or 4◦ C for others). Each isolated fraction is washed with density solution. Finally, an extensive dialysis is performed to eliminate the salt. Lipoproteins obtained by preparative ultracentrifugation or gel chromatography are analyzed in several ways. The lipoprotein fraction is assayed for its protein and lipid (cholesterol, triglyceride, and phospholipid) content and results are expressed as a percentage of mass (Singaraja et al., 2002, 2006; Ribas et al., 2005). A semi-quantitative analysis of the apolipoprotein composition is performed by polyacrylamide gel electrophoresis (PAGE) under denaturing (0.1% sodium dodecyl sulfate, SDS) and nonreducing conditions on ready-to-use gels (4% to 20%; Singaraja et al., 2002; MacDonald et al., 2008). After electrophoresis, gels are stained with Coomassie brilliant blue and bands corresponding to apolipoproteins are identified using molecular weight markers. Images are then scanned using a densitometer to obtain a semi-quantitative apolipoprotein composition.

Apolipoprotein measurements Mice express the same types of apolipoproteins as humans with some particularities. For example, murine apoAII is a homodimer consisting of two 82 amino acid chains, whereas its human counterpart is a single polypeptide chain. Moreover, human apoB exists in two different forms: apoB100, which is produced by the liver, and apoB48, a truncated form produced by the intestine. Although mouse apoB also exists in these two forms, apoB48 is not exclusively produced by the intestine, but also by the liver. Immunoreactants developed to measure human apolipoproteins cannot be used to measure murine apolipoproteins because either there is no sequence homology between the two species or sequence homology is only partial and thus insufficient for

correct assaying. For example, murine apoAII should be measured using an immunoreactant specifically developed against the murine protein. Two different strategies are available to obtain antigens for immunization. Native apolipoproteins can be obtained by preparative electrophoresis. For that, appropriate lipoproteins are obtained from mouse plasma by preparative ultracentrifugation (e.g., HDL and LDL to prepare apoAI and apoB, respectively) and separated by SDS-PAGE. The separated apolipoproteins are identified by comparing their electrophoretic migration with molecular weight markers; the gel band is recovered and used for immunization. An alternative strategy is to use synthetic peptides reproducing the mouse apolipoprotein sequence in its complete or partial form. Native apolipoprotein or synthetic peptides can then be used to immunize animals, such as rabbits or goats, with the exception of the mouse, and immunoserum is used to develop an immunoassay. The advantage of nephelometry or turbidimetry versus ELISA lies in the fact that immunoserum or derivatived immunoglobulins can be used instead of purified antibodies. Particular attention must be paid to apolipoprotein assays in transgenic mice expressing human apolipoproteins. In such animals, it is important to measure specifically both the human and the endogenous mouse proteins. The human protein should be measured using an immunoreactant obtained by immunization of mice with the appropriate antigen, whereas the mouse protein should be measured with an immunoreactant obtained against synthetic peptides encompassing specifically mouse, but not human, sequences. For example, in human apoAI transgenic mice, human and mouse apoAI can be measured using monoclonal antibodies obtained in immunized mice and immunoreactants against synthetic peptides, respectively. In human apoAII transgenic mice, human and mouse apoAII can be measured with immunoreactants obtained against each protein, as no cross-reactivity exists between apoAII of the two species. In addition to plasma lipid measurements, development of appropriate immunosera has been extensively used to measure apoAI, apoAII, apoB, as well as apoCIII in different experimental models (Castro et al., 1997; Singaraja et al., 2001, 2002, 2006; Coutinho et al., 2005; Ribas et al., 2005; Brunham et al., 2006, 2009; Calpe-Berdiel et al., 2008; MacDonald et al., 2008, 2009; Tancevski et al., 2008; Julve et al., 2010).

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Pre-β and α-HDL Pre-β-HDLs are referred to as nascent HDLs because these discoidal particles must undergo maturation processes that ultimately result in their conversion into mature spherical plasma HDLs. Although pre-β-HDLs are minor constituents of plasma HDL, there is an intense interest in these particles, as they are the initial acceptors of peripheral tissue cholesterol in the reverse cholesterol transport pathway (Castro and Fielding, 1988; Huang et al., 1993). The quantification of mouse pre-β and αHDL fractions is adapted from the method first described by Castro and Fielding (1988) using a nondenaturing two-dimensional electrophoresis-based quantification of human HDL fractions. The first separation is achieved by running plasma samples on an agarose gel, which separates lipoproteins according to their charge, namely in α-lipoproteins (containing α-HDL), pre-β-lipoproteins (containing VLDL and preβ-HDL), and β-lipoproteins (containing LDL). The second separation is carried out by migration of the separated proteins on a nondenaturing PAGE. Fractionated lipoproteins are then transferred onto a

nitrocellulose membrane and incubated with an anti-mouse apoAI antibody, revealing all apoAI-containing subfractions (Zanotti et al., 2008). Additionally, the nitrocellulose membrane is incubated with protein A (a protein from Staphylococcus aureus with a high affinity for immunoglobulins) labeled with 125 I; this radiolabeled protein A binds to anti-mouse apoAI-antibodies. Determination of radioactivity of each pattern allows quantification of pre-β and α-HDL fractions (Castro et al., 1997; Fig. 2). Alternatively, a quantitative method without radioactivity may be used. Pre-β and αHDL can be separated by a double immunoelectrophoretic technique according to their mobility. The first separation is achieved by running the plasma sample on an agarose gel as described above. The second separation is obtained by migration of the separated proteins on an agarose gel containing mouse anti-apoAI immunoserum. Finally, the gel is washed, dried, and stained with Coomassie blue revealing the two peaks of apoAI-containing lipoproteins corresponding to pre-β and αHDL. The peak area measured by scanning is proportional to the apoAI contained in each

A IgG

B pre␤

␣

MW

IgG

pre␤

␣

256

116 92

Overview of the Measurement of Lipids and Lipoproteins in Mice

Figure 2 Pre-β and α-HDL particles in plasma from a wild-type control (A) and human apoAI transgenic mouse (B). Twenty microliters of mouse serum were used to separate lipoprotein particles by bidimensional electrophoresis (first dimension, agarose gel electrophoresis; second dimension, PAGE). Fractionated lipoproteins were transferred onto a nitrocellulose membrane. Fractions were visualized by immunoblotting with rabbit specific anti-murine apoAI followed by radiolabeled protein A. The nontransgenic control mouse plasma contains predominantly particles with α mobility (2A), while in contrast, plasma from the apoAI transgenic mice showed pre-β and α-HDL particles. The pre-β profile consisted of three mice apoAI-containing particle types with apparent molecular weights (MW) estimated to be 92, 116, and 256 kDa (Reproduced with permission from Castro et al., 1997).

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fraction. Pre-β and α-HDL fractions are calculated by converting the percentage in each subfraction using apoAI concentrations previously measured in plasma (Singaraja et al., 2002; Coutinho et al., 2005).

MEASUREMENT OF ENZYME ACTIVITIES Lipoproteins are not static entities, but molecular complexes in permanent remodeling through numerous exchanges of lipids and apolipoproteins between the different fractions of lipoproteins and/or between lipoproteins and cells. Parts of these processes are spontaneous, whereas others are mediated by enzymes. In addition to plasma lipid, apolipoprotein, and lipoprotein measurements, it may be of interest to measure these enzymatic activities to better delineate the lipoprotein phenotype of a mouse model. It is noteworthy that CETP, one of the main enzymes responsible for lipoprotein remodeling in humans, is not expressed in the mouse.

Lecithin-cholesterol acyltransferase (LCAT) LCAT catalyses the esterification of free cholesterol. It thus plays a crucial role in the early steps of the cholesterol efflux process through conversion of unesterified cholesterol coming from peripheral cells and taken up by lipoprotein particles to cholesteryl esters (CEs). Thus, formed CEs are then internalized within the lipoprotein particle core. LCAT is activated by apoAI on the surface of HDL particles and it increases the size of particles from small pre-β HDL to α-migrating particles. Serum LCAT activity is determined using the so-called exogenous substrate method as described by Chen and Albers (Chen and Albers, 1982). Mouse serum, the source of LCAT, is incubated with a modified proteoliposome substrate containing apoAI, the obligate co-activator of LCAT, 14 C-free cholesterol, and POPC. Serum LCAT converts radioactive-free cholesterol to radioactive esterified cholesterol. After incubation, lipids are extracted and separated by thinlayer chromatography. Esterified cholesterol and free cholesterol are identified by reference to standards, and the corresponding radioactivity is measured. LCAT activity is expressed as number of moles of cholesterol esterified per volume of serum per time unit (Castro et al., 1997).

Phospholipid transfer protein (PLTP) PLTP promotes the transfer of phospholipids from triglyceride-rich lipoproteins (VLDL and chylomicrons) into HDL (for review, see Quint˜ao and Cazita, 2010). Free cholesterol is also a substrate of PLTP, but with a much lower activity. PLTP contributes to the remodeling of HDL particles, leading to the generation of large α-HDL, as well as preβ-HDL particles (Settasatian et al., 2001). In addition to lipids, PLTP is also able to transfer other amphipatic compounds, such as vitamin E and LPS (Jiang et al., 2002; Tzotzas et al., 2009). PLTP is present in all animal species and is expressed at high levels in the mouse (for review, see Masson et al., 2009). PLTP mass can be measured by immunoenzymologic methods using appropriate antibodies. In the mouse, PLTP mass has been essentially measured in mice overexpressing human PLTP using anti-HuPLTP polyclonal antibodies against specific sequences of human PLTP, which show no cross-reactivity with mouse PLTP (van Haperen et al., 2000). Plasma PLTP activity is determined with labeled liposomes as the PL donors and plasma HDL as the PL acceptor. PLTP activity is measured in total plasma by assessing the transfer of radiolabeled phosphatidylcholine (14 C or 3 H phosphatidylcholine liposomes) to an excess of exogenous human HDL3. Liposomes are precipitated by centrifugation and the radioactivity transferred to HDL is counted in the supernatant. PLTP activity is expressed as micromoles of phosphatidylcholine transferred from liposomes to exogenous human HDL3 per milliliter of plasma. Variations on this protocol have been described (Damen et al., 1982; van Haperen et al., 2000; Bouly et al., 2001; Kaser et al., 2003).

Lipoprotein lipase (LPL) and hepatic lipase (HL) LPL is an enzyme located at the surface of endothelium cells, which promotes hydrolysis of triacylglycerols in TG-rich lipoproteins to liberate FFAs. LPL is activated by apoCII and apoAV, and inhibited by apoCIII and angiopoietin, like protein 3 and 4. In mice, the action of LPL on TG is achieved by hepatic lipase action in the vascular compartment. After an intravenous injection of heparin to release cell-surface LPL, blood is collected and plasma isolated. Total HL and LPL activity are analyzed by incubation of post-heparin plasma, containing the released lipases, with a TG substrate emulsion containing

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radiolabeled triolein glyceride, in the presence or absence of 1M NaCl, which inhibits LPL activity and thus allows assessment of HL activity. The radioactivity of released FFA is measured. LPL activity is calculated as the fraction of total lipase activity inhibited by the presence of 1 M NaCl and it is expressed as the amount of free FA released per hour per ml of plasma. This method, first described by Olivecrona (Peterson et al., 1985), has been extensively used since then, with limited adaptations (Julve et al., 2000; Escol`a-Gil et al., 2001). Alternatively, a ready-to-use commercial kit is available, including a nonfluorescent substrate emulsion that becomes intensely fluorescent upon enzymatic digestion by LPL (Altomonte et al., 2004; Qu et al., 2007; Perdomo et al., 2010).

FUNCTIONAL TESTS IN VIVO Numerous functional tests have been developed to investigate the in vivo biological properties of lipoproteins and/or features of lipoprotein metabolism. Here, we will focus on the tests that are currently the most frequently used.

Functional tests to explore TG metabolism in vivo

Overview of the Measurement of Lipids and Lipoproteins in Mice

To determine if an abnormal plasma TG concentration is due to an altered hepatic production of TG-rich lipoproteins or to an alteration of their catabolism through LPL, the enzyme responsible for TG hydrolysis, the LPL inhibitor Triton WR-1339 (Tyloxapol), is often used. In vivo hepatic TG production is determined by assessing the plasma TG appearance rate after intravenous injection of Triton WR-1339 (Moir et al., 1995). Briefly, mice are injected in the tail vein with a solution containing Triton WR-1339, blood samples are collected during 2 hr, and plasma TG is measured in each sample as described in the Lipid and lipoprotein analysis section. The triglyceride production (PR) is calculated as follows: PR= a × PV (expressed in mg per hour), where a is the slope of the regression line of time (in hours) and TG concentrations after Triton injection, and PV is plasma volume expressed in ml [body weight (g) × 0.033]. The change in TG kinetics upon the Triton WR-1339 injection is directly proportional to the hepatic production of TG-rich lipoproteins, whereas absence of change suggests an altered catabolism of TG via LPL, indirectly measured this way. The ability of Triton WR-1339 to inhibit TGrich lipoprotein clearance may also be used to measure the hepatic apoB production rate

in vivo. After an injection of both Triton WR1339 and 35 S-methionine, de novo synthesized hepatic apoB is quantified by assessing radioactivity of apoB100 and apoB48 separated by SDS-PAGE (Voyiaziakis et al., 1999; Noga and Vance, 2003; Goldberg et al., 2008). It may be of interest to specifically measure turnover of the lipid (mainly TG) and the protein (mainly apoB) component of VLDL. For that, 3 H-TG-VLDL or 125 I-apoB-VLDL are intravenously injected and the kinetic changes of plasma radioactivity is measured to determine TG and apoB clearance rates (Shimizugawa et al., 2002). The oral fat gavage (OFG) test is used to measure the metabolism of TG comprising intestinal absorption, production of TG-rich lipoproteins, and subsequent vascular clearance. A single bolus of olive oil (a TG predominant solution) is administered and the kinetics of plasma TG concentrations are measured for ∼6 hr (Altomonte et al., 2004; Perdomo et al., 2010). To quantify the disappearance of TG in plasma and its parallel capture by tissues, a variant of OFG can be performed with a single bolus of radiolabeled TG and radioactivity counted in plasma and tissues (mainly liver, muscle, heart, and adipose tissues).

In vivo assay of macrophage reverse cholesterol transport The reverse cholesterol transport (RCT) pathway involves cholesterol efflux from peripheral extrahepatic tissues to HDL, plasma transport, and cholesterol uptake by the liver and excretion into bile and feces, or direct secretion in the intestine and excretion into feces. Rader and co-workers developed the first in vivo method to trace RCT. Macrophages loaded in vitro with radiolabeled cholesterol are intraperitoneally injected into mice. Feces are collected during the days after the injection, and blood and liver are collected after euthanasia. Radioactivity is measured in plasma, liver lipids, total sterols, and specifically in the cholesterol and bile acid fractions of feces. The amount of the tracer is expressed as a fraction of the injected dose (Zhang et al., 2003). Macrophages used can be primary cells, such as bone marrow–derived macrophages (BMDM), or cell lines (J774 or the macrophage-like cell line P388D1 derived from DBA/2 mice), allowing one to test the influence of the BMDM donor phenotype or the cell receiver phenotype, respectively, on RCT modulation. Since the first publication of the method, this protocol has been extensively used (Calpe-Berdiel et al., 2005; Rotllan

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et al., 2005; Naik et al., 2006; Wang et al., 2007; Calpe-Berdiel et al., 2008; Zanotti et al., 2008; Tancevski et al., 2010).

In vivo analysis of cholesterol absorption and cholesterol excretion via hepatobiliary and intestinal pathways Regulation of cholesterol homeostasis is a complex interplay of numerous metabolic pathways acting in different organs. The liver plays a central role in cholesterol excretion from the body, through the hepatobiliary cholesterol pathway, which was considered for years to be the most substantial route. Recent progress also identified a role for the intestine in cholesterol excretion, and evidence emerges that the proximal part of the small intestine is able to secrete cholesterol actively, a pathway called trans-intestinal cholesterol excretion (TICE). In mice, the cholesterol secreted in the intestine primarily originates from blood rather than from the intestine itself and intestinal cholesterol removal accounts for up to 70% of fecal neutral sterol excretion (for a review, see van der Velde et al., 2010). Numerous methods have been developed to measure intestinal cholesterol absorption, cholesterol excretion via the hepatobiliary and via the TICE pathways, as well as bile acid metabolism through liver and gallbladder, which are briefly described below. Cholesterol absorption is measured using the fecal dual isotope method (Salisbury et al., 1995; Schwarz et al., 1998). Mice receive a gavage with a single dose of 14 C -cholesterol and 3 H -sitosterol (a non-absorbable reference sterol) in oil. Each mouse is housed individually and feces are collected for 3 days. Fecal 14 C and 3 H are determined after lipid extraction. Cholesterol absorption is calculated by the fecal ratio method as follows: % cholesterol absorbed = [(gavaged 14 C/3 H – fecal 14 C/3 H)/(gavaged 14 C/3 H) ] × 100 (Carter et al., 1997; Wang and Carey, 2003; Lalloyer et al., 2006). An alternative method to measure cholesterol absorption is the plasma dual isotope ratio method (Turley et al., 1994). Briefly, mice receive an intravenous injection of radiolabeled 3 H-cholesterol dissolved in Intralipid (Fresenius Kabi, France) and an oral dose of 14 C-cholesterol dissolved in medium-chain FA-containing triglyceride oil. Plasma and feces are collected daily for 3 to 4 days, and bile is collected from anesthetized mice. 3 H and 14 C activity in plasma, bile, and feces is measured and cholesterol absorption is calculated (van der Veen et al., 2005).

Recently, a variant to measure the fractional cholesterol absorption rate using the plasma dual isotope method has been developed using nonradioactive isotopes. Briefly, mice receive an intravenous dose of cholesterol-D7 dissolved in Intralipid, together with an oral dose of cholesterol-D5 dissolved in medium-chain fatty-acid containing TG oil. Plasma enrichment of cholesterol-D7 and -D5 is measured by means of GC-MS after cholesterol extraction from blood samples. Fractional cholesterol absorption is calculated by division of the plasma ratio of oral cholesterol-D5 and intravenous cholesterol-D7 at the time of analysis by the administered D5/D7 ratio (van der Veen et al., 2009; Vrins et al., 2009; Jakulj et al., 2010; Sokolovi´c et al., 2010). In addition, the trans-intestinal cholesterol excretion can also be measured using intestine perfusion procedures in anesthetized mice, the bile duct being cannulated via the gallbladder, and bile acid and cholesterol measured in bile (van der Velde et al., 2008). This technique requires a very large expertise in mouse organ micromanipulation.

CONCLUSION: TECHNOLOGIES ON THE HORIZON All the qualitative and quantitative analysis methods of mouse lipoproteins described here concern known lipids and proteins and could be further developed for more global analysis. Although lipidomic and proteomic technologies have only been developed very recently, lipidomic and proteomic studies of plasma lipoproteins have already provided numerous examples of detailed characterization of distinct metabolic pathways, which may modulate lipoprotein metabolism under different conditions (e.g., health and disease states, lifestyle and dietary modification, lipid-modifying treatments, etc.). Recent reviews (Ekroos et al., 2010; Kontush and Chapman, 2010) present the results of lipidomic studies of both isolated plasma lipoproteins and total plasma lipids, which offer important clues about lipoprotein composition, metabolism, and functionality. A particular focus has been the proteomic analysis of HDL subclasses. At present, most of these studies are performed on human lipoproteins, and translation of lipidomic/proteomic methodologies from humans to mouse could facilitate the detailed characterization of lipid classes and molecular species present in plasma, as well as in lipoprotein fractions, and provide molecular

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details on composition, metabolism, and functions of lipoproteins in different genetically modified mouse models. To our knowledge, proteomic analysis of isolated lipoproteins has not yet been performed in mouse models. However, a few proteomic analyses have been performed in whole serum. For example, proteomic profiling has been done on whole serum of apoE-deficient mice during atherosclerosis progression (Dursun et al., 2010b), as well as upon pharmacological treatment with compounds such as diarbepoetin, a synthetic erythropoietin analogue that stimulates erythropoiesis (Dursun et al., 2010a), and nebivolol (a third-generation vasodilatory beta-blocker; Ozben et al., 2009). Moreover, the impact of the APOE3-Leiden isoform on the protein profile has been investigated by comparing whole plasma of transgenic apoE*3-Leiden mice versus isogenic wild-type controls (Davidov et al., 2004). Finally, progress is needed to better investigate the functionality of HDLs in the mouse, and more precisely the interplay between size, morphology, stability, and biological properties of HDLs (Kontush and Chapman, 2006; Cavigiolio et al., 2008) in the wide varieties of genetically modified mouse models.

LITERATURE CITED Altomonte, J., Cong, L., Harbaran, S., Richter, A., Xu, J., Meseck, M., and Dong, H.H. 2004. Foxo1 mediates insulin action on apoC-III and triglyceride metabolism. J. Clin. Invest. 114:14931503. Bouly, M., Masson, D., Gross, B., Jiang, X.C., Fievet, C., Castro, G., Tall, A.R., Fruchart, J.C., Staels, B., Lagrost, L., and Luc, G. 2001. Induction of the phospholipid transfer protein gene accounts for the high density lipoprotein enlargement in mice treated with fenofibrate. J. Biol. Chem. 276:25841-25847. Brunham, L.R., Kruit, J.K., Iqbal, J., Fievet, C., Timmins, J.M., Pape, T.D., Coburn, B.A., Bissada, N., Staels, B., Groen, A.K., Hussain, M.M., Parks, J.S., Kuipers, F., and Hayden, M.R. 2006. Intestinal ABCA1 directly contributes to HDL biogenesis in vivo. J. Clin. Invest. 116:1052-1062. Brunham, L.R., Singaraja, R.R., Duong, M., Timmins, J.M., Fievet, C., Bissada, N., Kang, M.H., Samra, A., Fruchart, J.C., McManus, B., Staels, B., Parks, J.S., and Hayden, M.R. 2009. Tissue-specific roles of ABCA1 influence susceptibility to atherosclerosis. Arterioscler. Thromb. Vasc. Biol. 29:548-554. Overview of the Measurement of Lipids and Lipoproteins in Mice

Calpe-Berdiel, L., Rotllan, N., Palomer, X., Ribas, V., Blanco-Vaca, F., and Escol`a-Gil, J.C. 2005. Direct evidence in vivo of impaired macrophage-specific reverse cholesterol transport in ATP-binding cassette transporter

A1-deficient mice. Biochim. Biophys. Acta 1738:6-9. Calpe-Berdiel, L., Rotllan, N., Fi´evet, C., Roig, R., Blanco-Vaca, F., and Escol`a-Gil, J.C. 2008. Liver X receptor-mediated activation of reverse cholesterol transport from macrophages to feces in vivo requires ABCG5/G8. J. Lipid Res. 49:1904-1911. Carter, C.P., Howles, P.N., and Hui, D.Y. 1997. Genetic variation in cholesterol absorption efficiency among inbred strains of mice. J. Nutr. 127:1344-1348. Castro, G.R. and Fielding, C.J. 1988. Early incorporation of cell-derived cholesterol into pre-betamigrating high-density lipoprotein. Biochemistry 27:25-29. Castro, G.R., Nihoul, L.P., Dengremont, C., de Geit`ere, C., Delfly, B., Tailleux, A., Fievet, C., Duverger, N., Den`efle, P., Fruchart, J.C., and Rubin, E.M. 1997. Cholesterol efflux, lecithincholesterol acyltransferase activity, and prebeta particle formation by serum from human apolipoprotein A-I and apolipoprotein AI/apolipoprotein A-II transgenic mice consistent with the latter being less effective for reverse cholesterol transport. Biochemistry 36:22432249. Cavigiolio, G., Shao, B., Geier, E.G., Ren, G., Heinecke, J.W., and Oda, M.N. 2008. The interplay between size, morphology, stability, and functionality of high-density lipoprotein subclasses. Biochemistry 47:4770-4779. Chen, C.H. and Albers, J.J. 1982. Characterization of proteoliposomes containing apoprotein A-I: A new substrate for the measurement of lecithin: Cholesterol acyltransferase activity. J. Lipid Res. 23:680-691. Coutinho, J.M., Singaraja, R.R., Kang, M., Arenillas, D.J., Bertram, L.N., Bissada, N., Staels, B., Fruchart, J.C., Fievet, C., JosephGeorge, A.M., Wasserman, W.W., and Hayden, M.R. 2005. Complete functional rescue of the ABCA1-/- mouse by human BAC transgenesis. J. Lipid Res. 46:1113-1123. Damen, J., Regts, J., and Scherphof, G. 1982. Transfer of [14C]phosphatidylcholine between liposomes and human plasma high density lipoprotein. Partial purification of a transfer-stimulating plasma factor using a rapid transfer assay. Biochim. Biophys. Acta 712:444-452. Davidov, E., Clish, C.B., Oresic, M., Meys, M., Stochaj, W., Snell, P., Lavine, G., Londo, T.R., Adourian, A., Zhang, X., Johnston, M., Morel, N., Marple, E.W., Plasterer, T.N., Neumann, E., Verheij, E., Vogels, J.T., Havekes, L.M., van der Greef, J., and Naylor, S. 2004. Methods for the differential integrative omic analysis of plasma from a transgenic disease animal model. OMICS 8:267-288. Dursun, E., Monari, E., Cuoghi, A., Bergamini, S., Ozben, B., Suleymanlar, G., Tomasi, A., and Ozben, T. 2010a. Proteomic profiling during atherosclerosis progression using SELDI-TOFMS: Effect of darbepoetin treatment. Acta Histochem. 112:432-443.

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Dursun, E., Ozben, B., Monari, E., Cuoghi, A., Tomasi, A., and Ozben, T. 2010b. Proteomic profiling in apolipoprotein E-deficient mice during atherosclerosis progression. Acta Histochem. 112:178-188. Ekroos, K., J¨anis, M., Tarasov, K., Hurme, R., and Laaksonen, R. 2010. Lipidomics: A tool for studies of atherosclerosis. Curr. Atheroscler. Rep. 12:273-281. Escol`a-Gil, JC., Julve, J., Marzal-Casacuberta, A., Ord´on˜ ez-Llanos, J., Gonz´alez-Sastre, F., and Blanco-Vaca, F. 2001. ApoA-II expression in CETP transgenic mice increases VLDL production and impairs VLDL clearance. J. Lipid Res. 42:241-248. Fievet, C., Fruchart, J., and Staels, B. 2007. Genetically-engineered animals as research models for atherosclerosis: Their use for the characterization of PPAR agonists in the treatment of cardiometabolic disorders. Front. Biosci. 12:4132-4156. Goldberg, I.J., Hu, Y., Noh, H., Wei, J., Huggins, L.A., Rackmill, M.G., Hamai, H., Reid, B.N., Blaner, W.S., and Huang, L.S. 2008. Decreased lipoprotein clearance is responsible for increased cholesterol in LDL receptor knockout mice with streptozotocin-induced diabetes. Diabetes 57:1674-1682. Huang, Y., von Eckardstein, A., and Assmann, G. 1993. Cell-derived unesterified cholesterol cycles between different HDLs and LDL for its effective esterification in plasma. Arterioscler. Thromb. 13:445-458. Jakulj, L., Vissers, M.N., van Roomen, C.P., van der Veen, J.N., Vrins, C.L., Kunne, C., Stellaard, F., Kastelein, J.J., and Groen, A.K. 2010. Ezetimibe stimulates faecal neutral sterol excretion depending on abcg8 function in mice. FEBS Lett. 584:3625-3628. Jiang, X., Tall, A.R., Qin, S., Lin, M., Schneider, M., Lalanne, F., Deckert, V., Desrumaux, C., Athias, A., Witztum, J.L., and Lagrost, L. 2002. Phospholipid transfer protein deficiency protects circulating lipoproteins from oxidation due to the enhanced accumulation of vitamin E. J. Biol. Chem. 277:31850-31856. Julve, J., Escol`a-Gil, J.C., Marzal-Casacuberta, A., Ord´on˜ ez-Llanos, J., Gonz´alez-Sastre, F., and Blanco-Vaca, F. 2000. Increased production of very-low-density lipoproteins in transgenic mice overexpressing human apolipoprotein A-II and fed with a high-fat diet. Biochim. Biophys. Acta 1488:233-244. Julve, J., Escol`a-Gil, J.C., Rotllan, N., Fi´evet C., Vallez, E., de la Torre, C., Ribas, V., Sloan, J.H., and Blanco-Vaca, F. 2010. Human apolipoprotein A-II determines plasma triglycerides by regulating lipoprotein lipase activity and high-density lipoprotein proteome. Arterioscler. Thromb. Vasc. Biol. 30:232-238. Kaser, S., Sandhofer, A., H¨olzl, B., Gander, R., Ebenbichler, C.F., Paulweber, B., and Patsch, J.R. 2003. Phospholipid and cholesteryl ester transfer are increased in lipoprotein lipase deficiency. J. Intern. Med. 253:208-216.

Kontush, A. and Chapman, M.J. 2006. Antiatherogenic small, dense HDL–guardian angel of the arterial wall? Nat. Clin. Pract. Cardiovasc. Med. 3:144-153. Kontush, A. and Chapman, MJ. 2010. Lipidomics as a tool for the study of lipoprotein metabolism. Curr. Atheroscler. Rep. 12:194-201. Lalloyer, F., Fi´evet, C., Lestavel, S., Torpier, G., van der Veen, J., Touche, V., Bultel, S., Yous, S., Kuipers, F., Paumelle, R., Fruchart, J.C., Staels, B., and Tailleux, A. 2006. The RXR agonist bexarotene improves cholesterol homeostasis and inhibits atherosclerosis progression in a mouse model of mixed dyslipidemia. Arterioscler. Thromb. Vasc. Biol. 26:2731-2737. MacDonald, M.L.E., Singaraja, R.R., Bissada, N., Ruddle, P., Watts, R., Karasinska, J.M., Gibson, W.T., Fievet, C., Vance, J.E., Staels, B., and Hayden, M.R. 2008. Absence of stearoylCoA desaturase-1 ameliorates features of the metabolic syndrome in LDLR-deficient mice. J. Lipid Res. 49:217-229. MacDonald, M.L.E., van Eck, M., Hildebrand, R.B., Wong, B.W., Bissada, N., Ruddle, P., Kontush, A., Hussein, H., Pouladi, M.A., Chapman, M.J., Fievet, C., van Berkel, T.J., Staels, B., McManus, B.M., and Hayden, M.R. 2009. Despite antiatherogenic metabolic characteristics, SCD1-deficient mice have increased inflammation and atherosclerosis. Arterioscler. Thromb. Vasc. Biol. 29:341-347. Masson, D., Jiang, X., Lagrost, L., and Tall, A.R. 2009. The role of plasma lipid transfer proteins in lipoprotein metabolism and atherogenesis. J. Lipid Res. 50:S201-S206. Moir, A.M., Park, B.S., and Zammit, V.A. 1995. Quantification in vivo of the effects of different types of dietary fat on the loci of control involved in hepatic triacylglycerol secretion. Biochem. J. 308:537-542. Naik, S.U., Wang, X., Da Silva, J.S., Jaye, M., Macphee, C.H., Reilly, M.P., Billheimer, J.T., Rothblat, G.H., and Rader, D.J. 2006. Pharmacological activation of liver X receptors promotes reverse cholesterol transport in vivo. Circulation 113:90-97. Noga, A.A. and Vance, D.E. 2003. A genderspecific role for phosphatidylethanolamine Nmethyltransferase-derived phosphatidylcholine in the regulation of plasma high density and very low density lipoproteins in mice. J. Biol. Chem. 278:21851-21859. Ozben, B., Dursun, E., Monari, E., Cuoghi, A., Bergamini, S., Tomasi, A., and Ozben, T. 2009. Proteomic profiling during atherosclerosis progression: Effect of nebivolol treatment. Mol. Cell. Biochem. 331:9-17. Perdomo, G., Kim, D.H., Zhang, T., Qu, S., Thomas, E.A., Toledo, F.G., Slusher, S., Fan, Y., Kelley, D.E., and Dong, H.H. 2010. A role of apolipoprotein D in triglyceride metabolism. J. Lipid Res. 51:1298-1311. Peterson, J., Olivecrona, T., and BengtssonOlivecrona, G. 1985. Distribution of lipoprotein lipase and hepatic lipase between plasma and

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tissues: Effect of hypertriglyceridemia. Biochim. Biophys. Acta 837:262-270. Qu, S., Perdomo, G., Su, D., D’Souza, F.M., Shachter, N.S., and Dong, H.H. 2007. Effects of apoA-V on HDL and VLDL metabolism in APOC3 transgenic mice. J. Lipid Res. 48:14761487. Quint˜ao, E.C.R. and Cazita, P.M. 2010. Lipid transfer proteins: Past, present and perspectives. Atherosclerosis 209:1-9. Ribas, V., Palomer, X., Roglans, N., Rotllan, N., Fievet, C., Tailleux, A., Julve, J., Laguna, J.C., Blanco-Vaca, F., and Escol`a-Gil, J.C. 2005. Paradoxical exacerbation of combined hyperlipidemia in human apolipoprotein A-II transgenic mice treated with fenofibrate. Biochim. Biophys. Acta 1737:130-137. Rotllan, N., Ribas, V., Calpe-Berdiel, L., Mart´ınCampos, J.M., Blanco-Vaca, F., and Escol`aGil, J.C. 2005. Overexpression of human apolipoprotein A-II in transgenic mice does not impair macrophage-specific reverse cholesterol transport in vivo. Arterioscler. Thromb. Vasc. Biol. 25:e128-e132. Salisbury, B.G., Davis, H.R., Burrier, R.E., Burnett, D.A., Bowkow, G., Caplen, M.A., Clemmons, A.L., Compton, D.S., Hoos, L.M., McGregor, D.G., Schnitzer-Polokoff, R., Smith, A.A., Weig, B.C., Zilli, D.L., Clader, J.W., and Sybertz, E.J. 1995. Hypocholesterolemic activity of a novel inhibitor of cholesterol absorption, SCH 48461. Atherosclerosis 115:45-63. Schwarz, M., Russell, D.W., Dietschy, J.M., and Turley, S.D. 1998. Marked reduction in bile acid synthesis in cholesterol 7alpha-hydroxylasedeficient mice does not lead to diminished tissue cholesterol turnover or to hypercholesterolemia. J. Lipid Res. 39:1833-1843. Settasatian, N., Duong, M., Curtiss, L.K., Ehnholm, C., Jauhiainen, M., Huuskonen, J., and Rye, K.A. 2001. The mechanism of the remodeling of high density lipoproteins by phospholipid transfer protein. J. Biol. Chem. 276:2689826905. Shimizugawa, T., Ono, M., Shimamura, M., Yoshida, K., Ando, Y., Koishi, R., Ueda, K., Inaba, T., Minekura, H., Kohama, T., and Furukawa, H. 2002. ANGPTL3 decreases very low density lipoprotein triglyceride clearance by inhibition of lipoprotein lipase. J. Biol. Chem. 277:33742-33748.

Overview of the Measurement of Lipids and Lipoproteins in Mice

Singaraja, R.R., Bocher, V., James, E.R., Clee, S.M., Zhang, L.H., Leavitt, B.R., Tan, B., Brooks-Wilson, A., Kwok, A., Bissada, N., Yang, Y.Z., Liu, G., Tafuri, S.R., Fievet, C., Wellington, C.L., Staels, B., and Hayden, M.R. 2001. Human ABCA1 BAC transgenic mice show increased high-density lipoprotein cholesterol and ApoAI-dependent efflux stimulated by an internal promoter containing liver X receptor response elements in intron 1. J. Biol. Chem. 276:33969-33979. Singaraja, R.R., Fievet, C., Castro, G., James, E.R., Hennuyer, N., Clee, S.M., Bissada, N., Choy, J.C., Fruchart, J.C., McManus, B.M., Staels, B.,

and Hayden, M.R. 2002. Increased ABCA1 activity protects against atherosclerosis. J. Clin. Invest. 110:35-42. Singaraja, R.R., Van Eck, M., Bissada, N., Zimetti, F., Collins, H.L., Hildebrand, R.B., Hayden, A., Brunham, L.R., Kang, M.H., Fruchart, J.C., Van Berkel, T.J., Parks, J.S., Staels, B., Rothblat, G.H., Fi´evet, C., and Hayden, M.R. 2006. Both hepatic and extrahepatic ABCA1 have discrete and essential functions in the maintenance of plasma high-density lipoprotein cholesterol levels in vivo. Circulation 114:13011309. Sokolovi´c, M., Sokolovi´c, A., van Roomen, C.P.A.A., Gruber, A., Ottenhoff, R., Scheij, S., Hakvoort, T.B., Lamers, W.H., and Groen, A.K. 2010. Unexpected effects of fasting on murine lipid homeostasis–transcriptomic and lipid profiling. J. Hepatol. 52:737-744. Tailleux, A., Torpier, G., Mezdour, H., Fruchart, J.C., Staels, B., and Fi´evet, C. 2003. Murine models to investigate pharmacological compounds acting as ligands of PPARs in dyslipidemia and atherosclerosis. Trends Pharmacol. Sci. 24:530-534. Tancevski, I., Wehinger, A., Demetz, E., Eller, P., Duwensee, K., Huber, J., Hochegger, K., Schgoer, W., Fievet, C., Stellaard, F., Rudling, M., Patsch, J.R., and Ritsch, A. 2008. Reduced plasma high-density lipoprotein cholesterol in hyperthyroid mice coincides with decreased hepatic adenosine 5 -triphosphate-binding cassette transporter 1 expression. Endocrinology 149:3708-3712. Tancevski, I., Demetz, E., Eller, P., Duwensee, K., Hoefer, J., Heim, C., Stanzl, U., Wehinger, A., Auer, K., Karer, R., Huber, J., Schgoer, W., Van Eck, M., Vanhoutte, J., Fievet, C., Stellaard, F., Rudling, M., Patsch, J.R., and Ritsch, A. 2010. The liver-selective thyromimetic T-0681 influences reverse cholesterol transport and atherosclerosis development in mice. PLoS ONE 5:e8722. Turley, S.D., Herndon, M.W., and Dietschy, J.M. 1994. Reevaluation and application of the dualisotope plasma ratio method for the measurement of intestinal cholesterol absorption in the hamster. J. Lipid Res. 35:328-339. Tzotzas, T., Desrumaux, C., and Lagrost, L. 2009. Plasma phospholipid transfer protein (PLTP): Review of an emerging cardiometabolic risk factor. Obes. Rev. 10:403-411. van der Veen, J.N., Kruit, J.K., Havinga, R., Baller, J.F., Chimini, G., Lestavel, S., Staels, B., Groot, P.H., Groen, A.K., and Kuipers, F. 2005. Reduced cholesterol absorption upon PPARdelta activation coincides with decreased intestinal expression of NPC1L1. J. Lipid Res. 46:526534. van der Veen, J.N., van Dijk, T.H., Vrins, C.L.J., van Meer, H., Havinga, R., Bijsterveld, K., Tietge, U.J., Groen, A.K., and Kuipers, F. 2009. Activation of the liver X receptor stimulates trans-intestinal excretion of plasma cholesterol. J. Biol. Chem. 284:19211-19219.

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van der Velde, A.E., Vrins, C.L.J., van den Oever, K., Seemann, I., Oude Elferink, R.P., van Eck, M., Kuipers, F., and Groen, A.K. 2008. Regulation of direct transintestinal cholesterol excretion in mice. Am. J. Physiol. Gastrointest. Liver Physiol. 295:G203-G208. van der Velde, A.E., Brufau, G., and Groen, A.K. 2010. Transintestinal cholesterol efflux. Curr. Opin. Lipidol. 21:167-171. van Haperen, R., van Tol, A., Vermeulen, P., Jauhiainen, M., van Gentxys, T., van den Berg, P., Ehnholm, S., Grosveld, F., van der Kamp, A., and de Crom, R. 2000. Human plasma phospholipid transfer protein increases the antiatherogenic potential of high density lipoproteins in transgenic mice. Arterioscler. Thromb. Vasc. Biol. 20:1082-1088. Voyiaziakis, E., Ko, C., O’Rourke, S.M., and Huang, L.S. 1999. Genetic control of hepatic apoB-100 secretion in human apoB transgenic mouse strains. J. Lipid Res. 40:2004-2012. Vrins, C.L.J., van der Velde, A.E., van den Oever, K., Levels, J.H., Huet, S., Oude Elferink, R.P., Kuipers, F., and Groen, A.K. 2009. Peroxisome proliferator-activated receptor delta activation

leads to increased transintestinal cholesterol efflux. J. Lipid Res. 50:2046-2054. Wang, D.Q. and Carey, M.C. 2003. Measurement of intestinal cholesterol absorption by plasma and fecal dual-isotope ratio, mass balance, and lymph fistula methods in the mouse: an analysis of direct versus indirect methodologies. J. Lipid Res. 44:1042-1059. Wang, M., Franklin, V., and Marcel, Y.L. 2007. In vivo reverse cholesterol transport from macrophages lacking ABCA1 expression is impaired. Arterioscler. Thromb. Vasc. Biol. 27:1837-1842. Zanotti, I., Pot`ı, F., Pedrelli, M., Favari, E., Moleri, E., Franceschini, G., Calabresi, L., and Bernini, F. 2008. The LXR agonist T0901317 promotes the reverse cholesterol transport from macrophages by increasing plasma efflux potential. J. Lipid Res. 49:954-960. Zhang, Y., Zanotti, I., Reilly, M.P., Glick, J.M., Rothblat, G.H., and Rader, D.J. 2003. Overexpression of apolipoprotein A-I promotes reverse transport of cholesterol from macrophages to feces in vivo. Circulation 108:661663.

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Using the Auditory Brainstem Response (ABR) to Determine Sensitivity of Hearing in Mutant Mice Neil J. Ingham,1 Selina Pearson,1 and Karen P. Steel1 1

Wellcome Trust Sanger Institute, Wellcome Trust Genome Campus, Hinxton, Cambridge, United Kingdom

ABSTRACT Measurements of auditory evoked potentials can be used to determine reliably an audiometric representation of hearing sensitivity in mice. In a high-throughput phenotyping screen of mice carrying targeted mutations of single genes, the auditory brainstem response (ABR) is used to gain an estimate of hearing threshold for broadband click stimuli and pure tone frequencies ranging from 6 to 30 kHz. Comparison of thresholds obtained in mutant and wild-type mice give a means to determine mild, moderate, and severe hearing impairment. This gives a clear advantage over using a “clickbox” test to assess hearing by observations of the Preyer reflex. The ABR screen has identified several mutant lines with mild and moderate hearing loss, which appear to demonstrate normal Preyer responses. The ABR technique also allows frequency-selective hearing loss to be C 2011 by John Wiley & Sons, Inc. identified. Curr. Protoc. Mouse Biol. 1:279-287  Keywords: hearing r evoked potential r electrophysiology r high-throughput screening

The auditory brainstem response (ABR) is an established method for determination of hearing sensitivity in patients in the clinic and in animal studies of hearing loss in the laboratory. Evoked electrical potentials recorded from the scalp are averaged to produce a physiological waveform representing auditory neural activity in the brainstem and can be achieved using a wide variety of equipment. In human testing, it is common for a very high number of potentials to be averaged to generate a recognizable ABR waveform. In animal studies, the use of anesthesia to sedate and immobilize the animal reduces the magnitude of biological/myogenic noise in the recording system and facilitates fewer record sweeps to be required to produce a clear averaged waveform.

BASIC PROTOCOL

Here, a custom hardware and software system for high-throughput ABR studies is described. It is typical to record click-evoked and tone-evoked ABRs across a range of sound pressure levels to determine the ABR threshold for each stimulus. Using tone pips of 6, 12, 18, 24, and 30 kHz allows an audiometric profile for each mouse tested to be constructed and an average audiogram for each mutant mouse line to be generated and compared with a baseline dataset of wild-type mice on the same genetic background. This approach has been used to screen over 330 lines of mutant mouse. Phenotyping results can be viewed online at http://www.sanger.ac.uk/mouseportal. NOTE: All protocols using live animals must first be reviewed and approved by an Institutional Animal Care and Use Committee (IACUC) or must conform to governmental regulations regarding the care and use of laboratory animals [e.g., the guidelines of the UK Home Office Animals (Scientific Procedures) Act 1986]. Auditory Brainstem Response (ABR) in Mice Current Protocols in Mouse Biology 1: 279-287, June 2011 Published online June 2011 in Wiley Online Library (wileyonlinelibrary.com). DOI: 10.1002/9780470942390.mo110059 C 2011 John Wiley & Sons, Inc. Copyright 

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Materials Male or female mice: control and mutant, tested at an appropriate age (no ABRs are likely to be recordable before the onset of hearing at around postnatal day 12) Ketamine/xylazine anesthetic (see recipe) Viscotears (liquid eye gel containing 2 mg/g Carbomer, with cetrimide; Dr Mann-Pharma) Atipamezole mix (recovery agent; see recipe) Sound-attenuating chamber (e.g., Industrial Acoustics) Heating blanket (to prevent hypothermia in the mouse) Stimulus-generation and calibration equipment (Tucker-Davis Technologies, TDT) including: RP2.1 Enhanced Real-time processor PA5 Programmable Attenuator SA1 Stereo Amplifier ACO Pacific Microphone (7017) and Preamplifier (PS9200) for calibration* MA3 Stereo Microphone Amplifier Sound Transducer (CTS Type 341; RS Components part no. 172-7712)* BNC and other connector cables 1-ml syringes (for injection; BD Plastipak) 27-G, 13-mm length hypodermic needles (for injection; BD Microlance) Response-processing equipment (Tucker-Davis Technologies) including: Needle electrodes (Chalgren Enterprises, cat. no. 112-812-48-TP; disposable low-profile EEG needle electrodes; Fig. 1A) Low-Impedance Recording Headstage/Preamplifier (RA4LI + RA4PA)* RA16 Medusa Base Station RP2.1 Enhanced Realtime processor HB7 Headphone Buffer (Optional) MS2 Monitor Speaker (Optional) BNC and other connector cables Personal computer, housing TDT gigabit interface, TDT driver software and bespoke averager software (available on request) Digital Oscilloscope (to view stimulus and electrode signals; optional) NOTE: Items with an asterisk (*) next to them are housed within the sound attenuating chamber.

Calibrate the sound system 1. Amplify and digitize (RA4LI and RA4PA) the electrode signals before returning them via an optical link to the RA16 basestation for filtering and then to the second RP2.1 for sampling at 100 kHz. An optional connection from the RA16 basestation, via the HB7 headphone buffer to the MS2 monitor speaker gives an audible output of the electrode signal to provide a very useful means of monitoring the condition of the animal. The signal allows the ECG of the mouse along with baseline myogenic activity and breathing rhythm to be heard. Synchronized triggering of stimulus presentation and recording allows short samples (20 msec) of electrode activity to be averaged under software control to generate the ABR-evoked potential waveform trace. It is not necessary to use artifact rejection (e.g., to reject sweep records containing an ECG component).

Auditory Brainstem Response (ABR) in Mice

Anesthetize the mouse and prepare for recording 2. Anesthetize the mouse with a 10 ml/kg intraperitoneal injection of ketamine/xylazine (0.1 ml/10 g body weight containing 10 mg ketamine/10 g body weight and 0.1 mg xylazine/10 g body weight). Once the mouse is immobilized, place it on the heating blanket in a sound-attenuating chamber and add a drop of Viscotears to each eye to prevent drying of the cornea.

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Figure 1 Positioning of sub-dermal electrodes for ABR recording. (A) Electrode configuration, to demonstrate the hook introduced onto commercially available EEG electrodes. The three views, from left to right, indicate an oblique view, an end-on view and a side view of the hook in the needle electrode. (B) The positioning of the active electrode on the vertex of the mouse. (C) The positioning of the reference and ground electrodes in the patch of bare skin behind the pinna overlying the bulla.

3. Once sufficient depth of anesthesia is achieved, as determined by abolition of the righting reflex and pedal withdrawal reflex, and following calibration of the sound system (step 2, above), insert needle electrodes (active electrode on the vertex, reference electrode overlying the left bulla, ground electrode overlying the right bulla; Fig. 1). Lay the mouse in a natural position, facing the loudspeaker, at a distance of 20 cm from the loudspeaker leading edge to the mouse interaural axis (Fig. 2).

Record ABRs 4. Set up the sound system equipment (Fig. 2) and calibrate the sound delivery system each day. Position the loudspeaker oriented along the mouse’s interaural axis 20 cm in front of where the mouse will subsequently be placed. Position the microphone (ACO 7017) where the mouse’s ears will be. Present 500-msec noise bursts 10 times. Auditory Brainstem Response (ABR) in Mice

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Figure 2 A schematic to illustrate the ABR setup used for high-throughput screening with free-field sound stimulation. The mouse is shown, with electrodes attached, on a heating blanket, with 20 cm between the leading edge of the loudspeaker and the mouse’s interaural axis. For sound system calibration, the microphone is positioned where the center of the animal’s head will be later positioned, oriented along the line of the interaural axis. Each TDT System3 module is housed within a zBus caddy. A gigabit (or optibit) interface provides communication between programmable modules and the computer system.

Using the MA3 amplifier at 40 dB gain, amplify microphone signals (from the ACO 7017) and digitize at 100-kHz sample rate (RP2.1). Subject each response to a Fast Fourier Transformation (FFT) and analyze the average FFT to produce an equalization curve, which is then used to ensure that all ABR stimuli are presented at a known sound pressure level (accounting for the frequency response of the sound transducer). Digitally generate stimuli under software control and convert to an analog waveform in the RP2.1 processor at 100-kHz sampling rate at 5 V pk-pk amplitude. Use the PA5 Programmable Attenuator to attenuate the waveform to produce the desired sound pressure level according to the calibration/equalization curve. By generating a large amplitude waveform in the RP2.1 digital-analog converters (DACs) followed by attenuation, an improved signal-to-noise ratio for the stimulus is produced; any noise inherent to the DAC is attenuated along with the signal. This is of particular importance for the production of the very low sound pressure level stimuli, critical for assessment of auditory sensitivity. If a lower amplitude signal waveform is generated on the DAC, removing the need to use the analog attenuators, then DAC noise is higher relative to the signal amplitude and a poorer signal-to-noise ratio results, which can result in noise masking of very low amplitude signals and a poorer estimate of pure tone ABR thresholds. The attenuated signal is then amplified and sent to the loudspeaker, producing the required sound pressure level at 20-cm distance from the loudspeaker, where the mouse’s head is now positioned.

5. Record a test ABR trace to ensure the system is functioning correctly, using clicks at 70 dB SPL (sound pressure level). 6. Record a series of ABRs using an array of click stimuli (a 10-μsec duration positive transient, 42.6/sec, 256 sweeps at a fixed phase), from 0 to 85 dB SPL in 5-dB steps. These responses are used to determine the ABR threshold to the click stimuli.

7. Record a series of ABRs using an array of stimuli of various frequencies and SPLs. Auditory Brainstem Response (ABR) in Mice

The run presents tones pips (5 msec duration, 1 msec rise/fall time, 42.6/sec, 256 sweeps at a fixed phase) across five frequencies (30, 24, 18, 12, and 6 kHz) at intensity levels

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increasing from 0 to 85 dB SPL in 5-dB steps. For routine screening, different SPL ranges are recorded for different test frequencies to improve time efficiency (6 kHz, 20 to 85 dB; 12 kHz, 0 to 70 dB; 18 kHz, 0 to 70 dB; 24 kHz, 10 to 70 dB; 30 kHz, 20 to 85 dB). Responses are recorded at each stimulus level, in decreasing frequency order before stepping up to the next highest stimulus level. These responses are used to determine the ABR thresholds to the tone stimuli. If mice appear to have hearing impairment, the upper limit of SPLs is extended to 95 dB for each test frequency and for clicks (this represents the upper limit of the linear range of our sound system at these frequencies).

8. Record a final test trace, again using clicks at 70 dB and compare with the initial test trace to ensure there has been no deterioration in the response during the measurements.

Promote recovery 9. Promote recovery of the mice from anesthesia with a 10 ml/kg intraperitoneal injection of atipamezole mix (0.1 ml/10 g body weight containing 0.01 mg atipamezole/10 g body weight). Return the mouse to its cage, place on a thermal mat or in a temperature-controlled ventilated rack. Monitor the recovering mice over the next 1 to 3 hr and return them to the holding racks once they are able to move well and respond to external stimuli. Analyze the data 10. Analyze data files to visualize the ABR waveforms and allocate thresholds. Organize ABR traces such that increasing dB SPL responses for a particular test stimulus are stacked together, aligned on the abscissa (time, msec; see Fig. 3)

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Figure 4 ABR audiograms for wild-type and Atp2b2Obv /+ mice. ABR thresholds are plotted for individual wild-type and mutant mice in the right and left panels, respectively. Mean ABR thresholds (± standard deviation, SD) are plotted for wild-type (green) and Atp2b2Obv /+ mice (blue) for the range of frequencies tested and for clicks. In the center panel, mean thresholds (± SD) are re-plotted for wild-type (green) and Atp2b2Obv /+ mice (blue) over a reference mean threshold (±1 SD and 2 SDs and shown as lighter green and darker green areas, respectively) calculated from a large population of wild-type mice on the same genetic background.

11. For each stimulus, determine the threshold by visual inspection of the trace stack as the lowest stimulus level (dB SPL) where any recognizable feature of the waveform can be identified (Fig. 3). As the stimulus level decreases, waveform amplitude reduces and wave peak latency extends. These trends are useful to help to identify true waveform features for determination of threshold.

12. For different cohorts of mice, calculate mean thresholds for each test stimulus and plot as a function of the test frequency, in addition to plotting the individual mouse response thresholds. Click thresholds are plotted at an arbitrary labeled point on the frequency abscissa to separate them from pure tone thresholds, which are plotted linked by a line (see Fig. 4). Plot control and mutant thresholds together to facilitate comparison of mean thresholds and add a baseline, reflecting the mean threshold (± 1 and 2 standard deviations) of a large population of control mice on the same genetic background. If the mutant mean threshold for any test stimulus exceeds ±2 SDs from the baseline average, the mutant line is considered to exhibit a hearing impairment phenotype (see Fig. 4).

REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps.

Atipamezole mix Mix 0.2 ml antisedan (Atipamezole hydrochloride 5 mg/ml; Pfizer) and 9.8 ml H2 O (Atipamezole 0.1 mg/ml) Ketamine/xylazine anesthetic mix Auditory Brainstem Response (ABR) in Mice

Mix 1 ml ketamine (Ketaset 100 mg/ml; Fort Dodge Animal Health), 0.5 ml xylazine (20 mg/ml; Rompun 2%, Bayer), and 8.5 ml H2 O. continued

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Store the anesthetic mix and recovery agent dilutions for a maximum of 7 days before it is disposed of via incineration (after binding with Clan Uni-Safe chemical spillage reagent; Fisher Scientific) and keep refrigerated at 4◦ C when not in use. Solutions should be warmed to room temperature before use. Stock bottles are stored at room temperature and any unused contents disposed of after 28 days using authorized local protocols.

COMMENTARY Background Information The ABR method has been widely used in clinical and research settings for many years. It provides an accurate means to estimate hearing sensitivity using electroencephalographic measures rather than behavioral responses. Because the recordings are derived from brainstem activity, with little or no contribution from higher centers, the measurements are largely unaffected by the use of anesthetic agents. Other methodologies can be used as screening methods for hearing with recovery of the mouse after the testing procedure. Testing for the Preyer reflex gives a crude indication of the presence or absence of hearing at high stimulus intensity but it will not detect mild or moderate hearing impairments. Otoacoustic emissions (distortion product, DPOAEs) can also be employed as a rapid screening method (Martin et al., 2006), but these measurements will only detect outer hair cell or middle ear pathology and will not identify as wide a range of hearing deficits as ABRs. The auditory steady-state response (ASSR) can also potentially be used as a screening technique based on electroencephalographic recordings similar to ABR (Pauli-Magnus et al., 2007). The ABR technique presented here is an optimized, fast recording protocol (approximately 15 to 20 min per mouse). It yields high-quality recordings and reliable estimates of thresholds in mice anesthetized with an easily reversible agent. This optimized protocol can be easily integrated into high-throughput phenotyping pipelines, which can cover a wide range of biological screening tests. Using a combination of click-evoked and tone-evoked ABRs, it is possible to identify more potential deafness-associated mutations than with the Preyer reflex testing or click ABR screening alone. Observations of Preyer reflex alone will not detect mutants showing mild or moderate hearing impairment. By using tone-evoked ABRs, it is possible to identify mutants with hearing deficits in particular frequency domains (e.g., mice showing severe high frequency deficits can have normal low frequency and click-evoked ABR thresholds).

Critical Parameters and Troubleshooting Anesthesia 1. Use artificial tears (Viscotears) during the procedure to prevent corneal drying, thus improving the welfare of the mice in recovery after the recording procedure. 2. Use a heating pad/blanket to maintain body temperature in the mice. Anesthetized mice will become hypothermic very quickly and the accompanying drop in metabolic rate will seriously affect the efficiency of inner ear function and result in artifactually high thresholds. 3. Ensure that the mouse achieves a sufficient depth of anesthesia. With the indicated dose of ketamine/xylazine, mice will quickly lose their righting and corneal reflexes. Abolition of pedal withdrawal reflexes may take a few minutes longer. It is important to achieve this depth so that the animal does not feel the insertion of the needle electrodes. Furthermore, if the mouse remains only lightly anesthetized, it will maintain a higher degree of muscle tone, which will introduce excessive myogenic electrical activity on the recorded traces. It is important to minimize sources of electrical noise arising from insufficient anesthesia to prevent the low-amplitude ABR waveform from being masked. Such masking can produce an artificial elevation of the estimated ABR threshold as the peak and trough features of the waveform used to identify an ABR will become buried in the noisefloor of the recording and only become visible at higher dB SPLs. Equipment 1. Use a sound-attenuating booth to reduce background (masking) acoustic noise. The booth should also be grounded (and thus act as a Faraday cage) to ensure low levels of ambient electrical noise, which can interfere with low-impedance recording systems. 2. Ensure that the needle electrodes are inserted sub-dermally. This provides better electrical contact with the animal and reduces electrical noise on the recorded traces.

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3. The microphone used for daily calibration of the recording system should itself be regularly calibrated (e.g., quarterly or biannually) to ensure that the sound levels presented to mice are consistent over long periods of time. Recording 1. The stimulus and recording parameters described in this protocol provide a reliable and time-efficient method to estimate ABR thresholds in a high-throughput phenotyping screen. The relatively low number of sweep records (256) contributing to each averaged waveform is made possible by minimizing sources of acoustic and electrical noise in the recording environment. In addition, the stimuli are presented relatively rapidly (42.6/sec), which helps to desynchronize any potential noise from the 50-Hz mains electricity supply. Many other studies use a slower rate of presentation (<30/sec). However, the higher stimulus presentation rate has a minimal effect on the recordings and facilitates a shorter time period before the mouse can be allowed to recover from the anesthesia, thus helping to improve the welfare of the mice. 2. As no artifact rejection is used in the recording system it is possible that the averaged trace can be significantly influenced by ECG activity, which happens to synchronize to the stimulus repetition rate. This is a rare occurrence and, if necessary, the affected stimulus frequency/level combinations can be repeated manually once the main automated data collection is complete.

Anticipated Results

Auditory Brainstem Response (ABR) in Mice

Examples of ABR audiometric profiles recorded in control (wild-type) and a hearingimpaired mutant mouse are shown in Figure 4. Using the recording protocol detailed here, mice tend to have the highest sensitivity of hearing (lowest thresholds) at 12 to 18 kHz, with average thresholds being around 10 to 20 dB higher at 6 kHz and 30 kHz. Click threshold often equates well with highest sensitivity tone threshold. Examples of ABR thresholds in hearingimpaired mutant mice are shown in Figure 4. Average thresholds for the Atp2b2Obv heterozygotes are significantly elevated compared to those of control mice on the same genetic background (Spiden et al., 2008). The use of ABR recordings to screen for hearing impairment is more informative and they will

identify more lines of interest than screens using only a test for the presence or absence of a Preyer reflex. Using the ABR method, the sensitivity of hearing across a range of frequencies can be estimated. This allows for even mild-tomoderate degrees of hearing impairment to be detected. The use of a range of test frequencies also gives a more comprehensive description of any hearing impairment detected. The use of click-evoked ABRs alone will give a good estimate of overall hearing sensitivity, but will not detect frequency-specific deficits in hearing; for example, loss of high frequency sensitivity would be missed as such lines can show normal sensitivity to low frequency tones and broadband clicks.

Time Considerations Using this protocol, it is possible to overlap the experiment on subsequent mice; for example, while one mouse is in the recording booth, the next mouse can be given its anesthesia injection. Induction of anesthesia requires ∼6 to 8 min following injection and prior to the start of electrophysiological recordings. Once in place in the booth, all ABRs can be recorded in 10 min using the semi-automated software routines. A minimum of four mutant mice per line should be subjected to ABR recording. More mice are subsequently tested if initial results show features of interest. Additionally, four to eight wild-type mice per week (from the same genetic background as the mutants) should be subjected to ABR recording. At full pipeline capacity, 44 to 48 mice per week can be tested and fully analyzed (plus any additional mice exhibiting ABR behavior warranting further examination). Allowing ∼15 min per mouse (overlapping anesthetic inductions and recordings), four mice can be tested per hour. This equates to ∼12 to 13 hr of experimental work per week, plus ∼1 hr set up/clean-up time per experimental day. Allowing for ∼5 min per mouse to upload data and allocate thresholds totals 3 to 4 hr of analysis per week. General organization and scheduling requires ∼3 to 4 hr per week. An additional 3 to 4 hr per week is required for quality control of analyses and project management. Thus, approximately 30 working hours per week are required to screen the hearing of 44 to 48 mice. This allows time for testing of additional mice from lines already identified as being of interest.

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Acknowledgments This work was supported by the Wellcome Trust (grant no. 077189).

Literature Cited Martin, G.K., Stagner, B.B., and Lonsbury-Martin, B.L. 2006. Assessment of cochlear function in mice: Distortion product otoacoustic emissions. Curr. Protoc. Neurosci. 8.21C.1-8.21C.18. Pauli-Magnus, D., Hoch, G., Strenzke, N., Anderson, S., Jentsch, T.J., and Moser, T. 2007. Detection and differentiation of sensorineural hearing loss in mice using auditory-steady state responses and transient auditory brainstem responses. Neuroscience 149:673–684. Spiden, S.L., Bortolozzi, M., Di Leva, F., Hrabe de Angelis, M., Fuchs, H., Lim, D., Ortolano, S., Ingham, N.J., Brini M., Carafoli, M., Mammano, F., and Steel, K.P. 2008. The novel mouse mutation Oblivion inactivates the PMCA2 pump and causes progressive hearing loss. PLoS Genetics 4:e1000238.

Internet Resources http://www.sanger.ac.uk/mouseportal Web site for phenotyping and mutant mouse resources at the Wellcome Trust Sanger Institute.

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Characterizing Bile Acid and Lipid Metabolism in the Liver and Gastrointestinal Tract of Mice Salvatore Modica,1 Stefania Murzilli,2 and Antonio Moschetta2,3 1

Institute of Food, Nutrition, and Health, ETH Zurich, Schwerzenbach, Switzerland Laboratory of Lipid Metabolism and Cancer, Department of Translational Pharmacology, Consorzio Mario Negri Sud, Santa Maria Imbaro, Italy 3 Clinica Medica “A. Murri,” Department of Internal and Public Medicine, University Aldo Moro of Bari, Bari, Italy 2

ABSTRACT Mouse models that mimic human diseases are invaluable tools to study and discover genetic and pharmacological therapies for human diseases. The protocols described in this article are intended to assess general clinical parameters in the context of the enterohepatic system under both normal and pathological conditions. Methods are presented for characterizing liver and intestinal function with a focus on bile acid and lipid metabolism C 2011 by John Wiley & Sons, in the gut-liver axis. Curr. Protoc. Mouse Biol. 1:289-321  Inc. Keywords: nuclear receptors r bile acids r gut-liver axis

INTRODUCTION Bile acids (BAs) are end products of cholesterol catabolism secreted from the liver into the intestine to allow the absorption of dietary lipids such as cholesterol and liposoluble vitamins. The presence of BAs in the intestine is also important to inhibit bacterial overgrowth and translocation across the mucosa barrier by maintaining the integrity of the intestinal architecture (Inagaki et al., 2006). BAs also represent the main factor determining bile flow and are important to solubilize biliary cholesterol and avoid gallstone formation (Moschetta et al., 2004). As signaling molecules, BAs can also regulate hepatic lipogenesis and decrease serum triglyceride levels (Watanabe et al., 2004). Despite their beneficial effects, BAs are also detergents and can be extremely toxic if they accumulate to high levels. Thus, during cholestasis, a medical condition characterized by interruption of bile flow, subsequent hepatic BA overload results in liver dysfunction and potentially liver failure. In this article, a series of protocols are presented to characterize the liver and gastrointestinal tract of mice when challenged with conditions that mimic metabolic human diseases related to BA and lipid metabolism. Thus, methods are provided for measuring BA levels and composition in serum, bile, liver, intestine, and feces (Basic Protocols 1 to 4), along with protocols for assessing serum and tissue lipid levels (Basic Protocols 8 to 11) and intestinal BA and cholesterol absorption (Basic Protocols 6 and 12). Protocols are also provided for the measurement of bile flow (Basic Protocol 5) and quantification of intestinal bacterial overgrowth and translocation across the mucosa barrier (Basic Protocol 7). Basic Protocol 13 describes a method for studying in vivo fatty acid synthesis. Finally, Basic Protocols 14 to 17 describe how to assess clinical parameters indicative of liver damage, such as serum transaminases (AST, ALT, ALP) and bilirubin. Characterizing Bile Acid and Lipid Metabolism Current Protocols in Mouse Biology 1: 289-321, June 2011 Published online June 2011 in Wiley Online Library (wileyonlinelibrary.com). DOI: 10.1002/9780470942390.mo100226 C 2011 John Wiley & Sons, Inc. Copyright 

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Abbreviations Used in This Article Bile acids (BAs), bile salt export pump (BSEP), multidrug-resistance related protein 2 (MRP2), ileal bile acid transporter (IBAT), cholic acid (CA), chenodeoxycholic acid (CDCA), lithocholic acid (LCA), deoxycholic acid (DCA), muricholic acid (MCA), taurocholic acid (TCA), tauro-β-muricholic acid (TβMCA), hyodeoxycholic acid (HDCA), colony-forming units (cfu), cholesterol gallstone disease (CGD), familial intra-hepatic cholestasis type 3 (PFIC3), N-ethyl-N-(2-hydroxy-3-sulfopropyl)-3,5-dimethoxyaniline (DAOS), peroxidase (POD), sterol regulatory element binding protein 1c (SREPB1c), serum aspartate aminotransferase (AST), glutamic oxalacetic transaminase (SGOT), serum alanine aminotransferase (ALT), glutamate pyruvate transaminase (SGPT), reticuloendothelial system (RES), gas-liquid chromatography (GLC), high performance liquid chromatography (HPLC), 3-α-hydroxysteroid dehydrogenase (3α-HSD), nitrotetrazolium blue (NTB). NOTE: All protocols using live animals must first be reviewed and approved by an Institutional Animal Care and Use Committee (IACUC) and must follow officially approved procedures for the care and use of laboratory animals. BASIC PROTOCOL 1

ENZYMATIC MEASUREMENT OF BILE ACIDS BAs are secreted into the bile duct via an active transport system against a concentration gradient. This active transport system involves ABC transporters such as the bile salt export pump (BSEP) and the multidrug-resistance related protein 2 (MRP2). Once in the bile duct, a small portion of BAs is reabsorbed by biliary ductal cells (cholangiocytes) and returns to hepatocytes through the periductal capillary plexus to be resecreted into bile. The majority of BAs reach the gallbladder, where they are concentrated up to 1000-fold. Upon a postprandial stimulus, BAs are released from the gallbladder into the duodenum to allow the absorption of dietary lipids and lipid-soluble vitamins. Since BA synthesis is an energetically expensive process that requires many enzymes, 95% of BAs are actively absorbed from the distal ileum via the ileal bile acid transporter (IBAT) and returned to the liver through the portal vein to be resecreted into bile (Love and Dawson, 1998). This cycle of secretion, absorption, and resecretion is termed “enterohepatic circulation” and ensures that only 0.5 g of BAs is lost per day via fecal excretion. This loss is compensated by de novo hepatic synthesis of BAs from cholesterol, which contributes less than 3% to hepatic BA secretion (Russell, 2003). Newly synthesized BAs are termed primary BAs, and in humans consist of cholic acid (CA) and chenodeoxycholic acid (CDCA). In mice, CDCA is converted to muricholic acid (MCA). Before secretion into the canalicular lumen, primary BAs are conjugated with the amino acids taurine (in human) or glycine (in mice) (Falany et al., 1994). While BAs are recycling in the gut-liver axis, bacterial enzymes present in the distal intestine modify their structure. Indeed, BAs undergo a deconjugation process, with formation of unconjugated BAs plus taurine or glycine. Some of these BAs are passively absorbed from enterocytes and shuttled back to the liver, where they are reconjugated. The rest is lost in feces. Thus, almost all BAs recovered in feces are deconjugated. In addition to the deconjugation process, bacterial enzymes present in the colon also convert the primary BAs CA and CDCA to the secondary BAs deoxycholic acid (DCA) and lithocholic acid (LCA), respectively.

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Under normal conditions, BAs are secreted into the gallbladder where they are stored to be delivered after a postprandial stimulus. As BAs recirculate between intestine and liver, analysis of biliary BA amount and composition is an indirect measure of the total BA pool size. Under hepatic pathological conditions such as cholestasis, BAs are released from the liver into the systemic circulation for clearance via the kidneys. Indeed, quantification of serum BAs represents an indicative marker of liver function. Current Protocols in Mouse Biology

The content of BAs in bile, serum, tissue, and feces can easily be measured by an enzymatic colorimetric assay. All BAs include a 3α-hydroxyl group, which can be oxidized to a 3-keto group by the enzyme 3-α-hydroxysteroid dehydrogenase. This enzyme simultaneously converts NAD+ to NADH, which can be measured spectrophotometrically using different approaches. Many commercial kits are available for this purpose. However, most of these kits suffer from interference from lipemic and hemolytic samples. The assay from Diazyme used here shows excellent precision (%CV < 5%), with no significant interference from lipemic and hemolytic samples. Moreover, being an enzymatic cycling method, it amplifies the signal for accurate and fast measurement using a small sample, up to 50% less than conventional nitrotetrazolium blue (NTB) methods.

Materials Mice Colorimetric Total Bile Acid Assay kit (containing Reagents R1 and R2, and a calibrator; http://www. diazyme.com; also see recipe for total bile acid reagents in Reagents and Solutions) Ethanol Refrigerated centrifuge 1-ml syringe 30-G needle 1.5-ml cuvettes or 96-well plates Spectrophotometer capable of measuring absorbance at 405 nm Additional reagents and equipment for hepatic bile acid extraction (Basic Protocol 2) and obtaining fecal bile acid pool (Basic Protocol 4) 1a. When measuring serum BAs: Centrifuge blood drawn from mouse 5 min at 6000 × g, 4◦ C, to separate serum. In the case of liver injury, as under cholestatic conditions, it is recommended to dilute serum samples with distilled water at least 1:4.

1b. To determine biliary BA levels: Recover the gallbladder contents using a 1-ml syringe with a 30-G needle. Since BAs are highly concentrated in bile, samples should be diluted at least 1:104 with distilled water.

1c. For measuring hepatic and fecal BAs: Refer to Basic Protocols 2 and 4, respectively. 1d. For measuring total BA pool: See Basic Protocol 3. 2. Pipet 270 μl of Reagent R1 into a 1.5-ml cuvette (pipet 210 μl if using a 96-well plate with 300-μl well depth), to which 4 μl of sample, calibrator, or blank [distilled water for serum and biliary BAs, 75% (v/v) ethanol for hepatic BAs (also see Basic Protocol 2), 100% ethanol for total BA pool (also see Basic Protocol 3)] have been added. 3. Incubate 5 min at 37◦ C. 4. Pipet 90 μl of Reagent R2 into the cuvette and mix immediately (pipet 70 μl if using a 96-well plate). 5. Incubate 2 to 3 min at 37◦ C. 6. Read the absorbance at 405 nm. 7. Calculate the total BA concentration using the following formula: [(sample A405 – blank A405 ) / (calibrator A405 – blank A405 )] × 50 μmol/liter

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where 50 μmol/liter is the calibrator concentration. This procedure is linear from 1 to 180 μmol/liter. A sample with a BA level exceeding the linearity limit should be diluted with blank and reassayed, incorporating the dilution factor in the calculation of the volume. BASIC PROTOCOL 2

HEPATIC BILE ACID EXTRACTION BAs are produced from hepatocytes through a multistep process that involves many enzymes, and are then actively secreted into the gallbladder. When this process is impaired, BAs accumulate in the liver and lead to liver inflammation and necrosis (e.g., cholestasis). Thus, the measurement of hepatic BA levels is important to identify and characterize hepatic dysfunctions.

Materials Mice 75% (v/v) ethanol Surgical instruments including scissors and clamps Dounce homogenizer with tight-fitting glass pestle 50◦ C heating block 5-ml glass tubes (5 ml) Refrigerated centrifuge Additional reagents and equipment for euthanasia of the mouse (Donovan and Brown, 2006) 1. Euthanize mouse by carbon dioxide inhalation (Donovan and Brown, 2006). Test for absence of vital signs before proceeding. 2. Perform laparotomy, then identify and remove the liver. 3. Weigh 50 to 100 mg of liver. 4. Homogenize at room temperature liver fragments in 1 ml of 75% ethanol using a Dounce homogenizer with a tight-fitting glass pestle, at setting 3. 5. Incubate the homogenate 2 hr at 50◦ C in glass tubes. 6. Centrifuge 10 min at 6000 × g, 4◦ C. 7. Remove and retain the supernatant fraction, which contains BAs. The sample is now ready for assay of BA concentration as described in Basic Protocol 1. BASIC PROTOCOL 3

Characterizing Bile Acid and Lipid Metabolism

DETERMINATION OF TOTAL BILE ACID POOL SIZE AND COMPOSITION Under normal conditions, only a small amount of BAs circulates in serum, and by definition the BA pool is composed of all BAs that are present in the liver, gallbladder, and small intestine. Thus, in this protocol, liver, gallbladder, and small intestine are harvested, collected, and minced with scissors into small pieces. BAs are extracted by boiling samples in ethanol and purified by filtration. After extraction, the BA pool size can be determined using an enzymatic assay (Basic Protocol 1), while HPLC-MS/MS is used to measure the exact composition of BAs present in the pool. The most abundant species of BAs recovered into the total BA pool are expected to be tauro-β-muricholic acid (TβMCA) and the taurocholic acid (TCA). This protocol is extremely important to define the total amount of circulating BAs in the mouse, which reflects both hepatic bile acid neosynthesis (5% of the pool) and

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enterohepatic circulating bile acids (95% of the pool). Comparing the total pool size with the biliary and fecal BA levels allows the investigator to dissect the partitioning of the pool with relevance to a series of pathogenic mechanisms that regulate quantity and quality of circulating BAs. The data obtained are needed for the identification of novel therapeutic strategies in diseases such as cholestasis, cancer, and, recently, diabetes.

Materials Mice Ethanol (HPLC grade) 1 mg/ml deuterated glycol-cholic acid (glycol-CA; Point-Clare) Ultrapure H2 O Methanol (HPLC-grade) Formic acid (mass spectrometry grade; Fluka) Animal balance Surgical instruments including scissors and clamps 250-ml beakers with watch glasses Hemostats Multi-position hot plate/stirrer Glass funnels No. 2 Whatman filter paper 100-ml volumetric flasks 5-ml glass tubes Additional reagents and equipment for euthanasia of the mouse (Donovan and Brown, 2006), enzymatic BA assay (Basic Protocol 1), and HPLC-MS/MS (Support Protocol) 1. Record animal body weight to normalize results. 2. Euthanize mouse by CO2 inhalation (Donovan and Brown, 2006). Test for absence of vital signs before proceeding. 3. Perform laparotomy. Remove the liver with the gallbladder and the small intestine from the animal and place all in a 250-ml glass beaker containing approximately 100 ml of 100% ethanol. When removing the small intestine, clamp both the proximal and distal ends with hemostats and carefully remove all attached mesentery without tearing the intestinal wall. Cut the common bile duct just above where it enters the duodenum to avoid tearing open the intestine when it is removed. It is important that the entire small intestine with all of its contents be placed into the ethanol. This also applies to the gallbladder and its contents.

4. Cut the liver and small intestine into small pieces with scissors. Rinse the scissors with ethanol between mice and add a stir bar to each beaker containing the collected organs from an individual mouse. 5. As internal standard, add 100 μl of 1 mg/ml deuterated glycol-CA to the beaker with the organs. This is important to evaluate the recovery of BAs by HPLC-MS/MS (e.g., if the recovery is 100%, 100 mg of deuterated glycol-CA will be found in a final volume of 100 ml).

6. Place the beakers on a multi-position hot plate/stirrer for extraction and cover each beaker with a watch glass. Set the heat so that the ethanol comes to a gentle boil and set the stir rate so that there is uniform stirring. 7. When the ethanol is reduced to around 30 ml, remove the beakers and allow them to cool to room temperature.

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8. Filter each sample using a glass funnel and no. 2 Whatman filter paper into a 100-ml volumetric flask. Be sure to rinse each beaker three times, each time with 15 to 20 ml ethanol. When filtering is complete, bring each volumetric flask to a volume of 100 ml with ethanol. 9. Pour samples into 5-ml glass tubes and store indefinitely at −20◦ C (until analysis). 10. Determine the total amount of BA in the pool (in μmol) by the enzymatic assay described in Basic Protocol 1 and correct for dilution (if one was made) and the amount of deuterated glycol-CA recovered (reported by HPLC-MS/MS analysis; see Support Protocol). Divide the results by the body weight and express the amount of BAs in the total pool as μmol/100 g body weight. 11. For the analysis of BA composition, dilute samples from 1:2 to 1:20 (v/v) with water/ethanol/methanol/formic acid (25%, 50%, 25%, 1% v/v/v/v). Sample dilution depends on the abundance of the different species of BAs.

12. Use 20 μl for HPLC-MS/MS analysis of BA composition (see Support Protocol). 13. Quantify BA fractions in the pool using the calibration curve generated with BA standards in the range of 0.01 to 10 μg/ml (see Support Protocol). SUPPORT PROTOCOL

HPLC MEASUREMENT OF BILE ACIDS The knowledge of the BA composition is of extreme importance for several pathophysiological conditions such as cholestasis, cholesterol gallstone disease, and intestinal malabsorption. HPLC is the method of choice when we measure BA composition in bile and total pool.

Materials Extracted BAs (Basic Protocol 3) Ultrapure water via Milli-Q system (Millipore) Methanol (HPLC grade) Formic acid (mass spectrometry grade; Fluka) BA standard working solutions (see recipe for BA standards and internal standards) in the range of 0.01 to 10 μg/ml for preparing calibration curves Internal standard working solutions (1S1 and IS2; see recipe for BA standards and internal standards) Gradient solution A: methanol (HPLC grade) containing 10 mM ammonium acetate (analytical grade) and 0.25% (v/v) formic acid (mass spectrometry grade; Fluka) Gradient solution B: Ultrapure H2 O containing 10 mM ammonium acetate (analytical grade) and 0.25% (v/v) formic acid (mass spectrometry grade; Fluka) Perkin Elmer 200 autosampler with 20-μl sample loop Perkin Elmer 200 quaternary HPLC pump 250 × 3.0 mm i.d., 5-μm Luna C18(2) column provided with a 4.0 × 2.0–mm i.d., 5-μm Luna C18(2) SecurityGuard System (Phenomenex) API 365 triple quadrupole mass spectrometer (PE Sciex, http://www.absciex.com/) Turbo ion spray source (PE Sciex, http://www.absciex.com/) TurboQuan 1.0 software (PE Sciex, http://www.absciex.com/) 1. Dilute extracted BAs 1:20 for the quantification of taurocholic and tauromuricholic acids, and 1:2 for the quantification of all the other bile acids, with water/methanol/formic acid (49.5:49.5:1, v/v/v, final solvent composition). Characterizing Bile Acid and Lipid Metabolism

2. Add 100 μg/ml IS1 working solution to obtain a 0.5 μg/ml final concentration. 3. Mix by vortex for 5 min.

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4. Centrifuge 15 min at 20,000 × g, room temperature. 5. For quantitative analyses, prepare calibration curves in the range of 0.01 to 10 μg/ml in methanol/water/formic acid (49.5:49.5:1, v/v/v), with IS1 (for conjugated BAs) or IS2 (for unconjugated BAs) at the concentrations of 0.5 and 1 μg/ml, respectively. 6. Place samples in an autosampler (e.g., Perkin Elmer 200 autosampler) thermostatted at 4◦ C and equipped with a 20-μl sample loop. 7. Achieve separations with a Perkin Elmer 200 quaternary HPLC pump equipped with a 250 × 3.0–mm i.d., 5 μm, Luna C18(2) column provided with a 4.0 × 2.0 mm i.d., 5 μm, Luna C18(2) SecurityGuard System, both from Phenomenex. Use a flow rate of 0.5 ml/min and a linear gradient of methanol (A) in water (B), both containing 10 mM ammonium acetate and 0.25% formic acid (75% A for 5 min, then to 95% A in 10 min and to 100% A in 3 min with 5-min hold). Connect the HPLC system to an API 365 triple quadrupole mass spectrometer through a turbo ion spray source. Perform analyses in negative ion mode by Selected Reaction Monitoring (SRM) using the most sensitive precursor to product ion transitions (see Table 1). 8. Set dwell time at 150 msec for each compound and optimize all the other mass spectrometric parameters for maximum sensitivity after infusion of the single compound standards. The system and the analytical conditions used allow the chromatographic separation of the isomeric compounds having the same precursor to product ion transitions.

9. Perform peak integration of extracted ion chromatograms and all calculations of concentrations and regression parameters using PE Sciex TurboQuan 1.0 software. Table 1 LC-MS/MS Method Performance

Compound

Precursor ion (m/z)

Production (m/z)

Linearity range (μg/ml)

LOD (μg/ml)

TCA

514

80

0.50-10.0

0.010

TDCA

498

80

0.025-1.0

0.025

TCDCA

498

80

0.025-1.0

0.010

THDCA

498

80

0.025-1.0

0.025

TUDCA

498

80

0.025-1.0

0.010

TLCA

482

80

0.025-1.0

0.010

TMCA

514

80

0.50-10.0

0.010

GCA

464

74

0.025-1.0

0.010

GDCA

448

74

0.025-1.0

0.010

GCDCA

448

74

0.025-1.0

0.010

CA

407

407

0.050-5.0

0.010

DCA

391

391

0.050-1.0

0.025

CDCA

391

391

0.25-2.5

0.100

UDCA

391

391

0.25-2.5

0.100

HDCA

391

391

0.25-2.5

0.100

LCA

375

375

0.25-2.5

0.100

GCA-D4

468

74

—

—

CA-D4

411

411

—

—

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10. Calculate standard curve equations using weighted (1/y) linear regression of internal ratios (analyte/IS peak areas) versus analyte concentrations. The linearity of the method is assessed as the range of concentrations in which the correlation coefficient (r2 ) is ≥ 0.985 with precision and accuracy ≤ 15%, except for the lower limit of quantification, for which precision and accuracy ≤ 20% are accepted. The limit of detection is assessed as the lowest concentration with a signal intensity at least three times greater than the background level. Method performance is summarized in Table 1. BASIC PROTOCOL 4

DETERMINATION OF FECAL BILE ACID POOL SIZE AND COMPOSITION After being delivered into the small intestine for the intestinal solubilization and uptake of dietary lipids, the majority of BAs are reabsorbed at the level of the distal ileum, and only a small amount is lost in feces. However, the assessment of the size and composition of BAs in feces can provide information about diseases related to the enterohepatic system. For instance, low levels of BAs in feces may indicate a problem related to their hepatic secretion (e.g., cholestasis), while high levels of BAs may be responsible for chronic diarrhea occurring in the context of Crohn’s disease, post-gastrectomy syndrome, and short bowel syndrome (Jung et al., 2007). Moreover, high levels of secondary BAs in feces have been linked to colorectal cancer development (Nagengast et al., 1995). In this protocol, BAs are extracted by boiling feces under reflux in the presence of sodium borohydride, HCl, and NaOH, and purified by filtration. The majority of BAs recovered in feces are deconjugated, and the most abundant species are TβMCA, DCA, and hyodeoxycholic acid (HDCA).

Materials Mice 1 μg/100 μl deuterated cholic acid (CA) (https://www.cdnisotopes.com/) 2 mg/ml sodium borohydride in ethanol (prepare fresh) 2 N HCl 10 N NaOH Nitrogen source 20% (v/v) methanol and 100% methanol (HPLC grade) Ultrapure H2 O Ethanol (HPLC grade) Formic acid (mass spectrometry grade) BA standard working solutions (see recipe for BA standards and internal standards) in the range of 0.01 to 10 μg/ml for preparing calibration curves 100-ml glass beakers 50-ml plastic and glass tubes with caps 60◦ C, 70◦ C, and 120◦ C heat blocks No. 2 Whatman filter paper C18 Bond Elute column (500 mg/6 ml; Varian) Suction manifold (Varian) 15-ml conical tubes (e.g., BD Falcon) Additional reagents and equipment for HPLC-MS/MS analysis of BA composition (see Support Protocol)

Characterizing Bile Acid and Lipid Metabolism

1. Collect feces from either group-housed or individually housed animals over three consecutive 24-hr periods or over a single 72 hr period toward the end of the study. Weigh all animals before and after the 3-day collection.

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2. Place feces in 100-ml glass beaker and dry in air until the weight no longer changes (∼1 to 2 days). Record weights of dried feces for each collection period. 3. Grind each lot of feces to a fine powder using a mortar and pestle and transfer to 50-ml plastic tubes for storage at room temperature. 4. For each ground fecal sample, take single or duplicate 1-g aliquots and add to 50-ml glass tubes. 5. As internal standard, add 300 μl of 1 μg/100 μl deuterated CA. This is important to evaluate the recovery of BAs by HPLC-MS/MS (e.g., if the recovery is 100%, 3 μg of deuterated CA will be found in a final volume of 3 ml).

6. To reduce 3-keto-BAs to 3-α-BAs, add 10 ml of freshly prepared 2 mg/ml sodium borohydride solution in ethanol to each tube, vortex vigorously, and let stand at room temperature for 1 hr. 7. For saponification of BAs, add 500 μl 2 N HCl, vortex vigorously, then add 2 ml 10 N NaOH and vortex again. 8. Cap the tubes and digest under reflux (by placing the sealed tube in a heat block at about 120◦ C) for 12 to 14 hr to extract BAs. 9. Filter each sample through no. 2 Whatman filter paper into a 50-ml glass tube, and rinse the original tube and filter paper with ethanol (5 ml, three times). 10. Dry extract under nitrogen at about 70◦ C and then add 9 ml of Ultrapure water to each tube (the sample should be essentially dry before water is added). Vortex vigorously and then let stand for at least 1 hr before chromatography. Gently vortex samples immediately before applying to the column. 11. Apply 3 ml of each aqueous suspension to individual C18 Bond Elute columns (500 mg/6 ml) that have been prewashed with 3 ml of 100% methanol followed by 3 ml of Ultrapure water. Run columns on a suction manifold. 12. After a sample has been loaded into its column, wash the column twice, each time with 6 ml of 20% methanol. 13. Elute BAs from column with 6 ml methanol into 15-ml conical tubes. Dry down extracts under nitrogen gas at about 60◦ C. 14. Redissolve BAs in 3 ml of methanol. Vortex tubes vigorously. Sample will usually contain some insoluble material. Store samples indefinitely at −20◦ C in glass tubes, until analysis. 15. Determine the amount of BAs in feces (μmol) by the enzymatic assay described in Basic Protocol 1 and correcting for dilutions (if they were made) and the amount of deuterated CA recovered (reported by HPLC MS/MS analysis). Divide the results for the number of days in which the feces were collected and express the results per 100 g of body weight. For example, dilute 20 μl of the 3 ml of methanol (step 14) with 80 μl of water for a dilution factor of 5 (the blank has to be made by methanol and water in the same proportion). After performing the enzymatic assay, assume, for this example, a concentration of 30 μM. The final concentration is 30 μM × 5 = 150 μM. Indeed, the μmol of BAs present in 3 ml of methanol are (150 μM/1000 μl) × 3 = 0,45 μmol. If the percentage of recovery is 30%, 70% of BAs have been lost. Thus, it will be necessary to correct for this (70% of 0.45 μmol is 0.315 μmol). The final μmol in 3 ml of methanol are 0.45 μmol + 0.315 μmol = 0.765 μmol. However, these are the μmol present in 3 ml of the starting 9 ml (see step 10). So, the total amount of BAs is given by 0.765 μmol × 3 = 2.295 μmol.

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From this amount, it is necessary to subtract the amount of the internal standard that was added (3 μg of deuterated CA = 0.0073 μmol). Thus, 2.295 μmol –0.0073 μmol = 2.2877 μmol. These are the μmol present in 1 g of feces (see step 4). So, if 4 g of feces were collected over 3 days, there will be 2.2877 μmol × 4 = 9.1508 μmol. Dividing now by 3 will provide the μmol of BAs per day, which is 9.1508 μmol/3 =3.05 μmol. Assuming that the body weight (BW) of a mouse is 30 g, the final result is given by (3.05 μmol/ 30 grams) × 100 g = 10 μmol/day per 100 g of BW.

16. For the analysis of BA composition, dilute samples from 1:2 (v/v) with water/ethanol/methanol/formic acid (25%, 50%, 25%, 1% v/v/v/v final composition). 17. Use ∼20 μl for HPLC-MS/MS analysis of BA composition. See Support Protocol for procedural details. 18. Quantify BA fractions in feces by comparing with a BA standard curve in the range of 0.01 to 10 μg/ml. BASIC PROTOCOL 5

BILE FLOW MEASUREMENT Bile is a complex fluid containing water, electrolytes, and organic molecules such as BAs, cholesterol, phospholipids, and bilirubin. Bile flows out of the liver through the left and right hepatic ducts, which come together to form the common hepatic duct. This duct then joins with a duct connected to the gallbladder, called the cystic duct, to form the common bile duct. The common bile duct enters the small intestine at the sphincter of Oddi (a ring-shaped muscle), located several cm below the stomach. About half of the bile secreted between meals flows directly through the common bile duct into the small intestine. The rest of the bile is diverted through the cystic duct into the gallbladder to be stored. In the gallbladder, up to 90% of the water in bile is absorbed into the bloodstream, making the remaining bile highly concentrated. When food enters the small intestine, a series of neuronal hormonal signals triggers the gallbladder to contract and the sphincter of Oddi to relax and open. Bile then flows from the gallbladder into the small intestine to mix with food contents and perform its digestive functions. Correct bile flow is important since many waste products such as bilirubin and excess cholesterol are cleared from the body by secretion into bile and subsequent elimination in feces. The levels of conjugated BAs provide the primary determinant for the rate of bile flow. Genetic inactivating mutations in the bile salt export pump (BSEP) gene result in progressive familial intra-hepatic cholestasis type 2 with greatly reduced bile flow. Thus, measuring bile flow may provide useful information in the study of hepatic diseases.

Materials Mice, fasted for 4 hr Ketamine (Sigma-Aldrich) Xylazine (Bayer) Animal balance Syringe and 26-G needle Surgical instruments PE-10 catheter Additional reagents and equipment for euthanasia of the mouse (Donovan and Brown, 2006) 1. Weigh mice. Characterizing Bile Acid and Lipid Metabolism

2. Induce anesthesia by administration via intraperitoneal injection (using a 26-G needle) of ketamine (Sigma-Aldrich, 100 mg/kg body weight) and xylazine, 2% (Bayer, 500 μl), up to 10 ml total volume.

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Figure 1

Common bile duct ligation.

3. Open the chest area with a midline incision. 4. Ligate cystic duct and common bile duct (Fig. 1). 5. Cannulate the common bile duct just above the ligated area with a PE-10 catheter. 6. Collect bile for 10 to 15 min. 7. Determine bile flow as μl per min per 100 g of body weight, assuming a density of 1 g/ml. 8. At the end of the study, euthanize mouse by CO2 inhalation (Donovan and Brown, 2006).

INTESTINAL BILE ACID ABSORPTION BA reabsorption by ileal enterocytes is essential for cholesterol homeostasis, intestinal absorption of dietary fats and vitamins, and proper regulation of bile flow and biliary lipid secretion. The initial step in BA reabsorption by these epithelial cells is largely mediated by IBAT (Oelkers et al., 1997). 95% of BAs are reabsorbed at the level of the distal ileum by this transporter and returned to the liver as part of the cycle referred to as enterohepatic circulation. A reduction in intestinal BA reabsorption and subsequent increase in fecal bile acid excretion contributes to chronic diarrhea and steatorrhea that occur in different clinical contexts. Moreover, high levels of BAs in the colon can promote colorectal cancer development. Assessing intestinal BA reabsorption can therefore provide important insights in mouse models of human diseases. Indeed, the knowledge of the exact amount of BAs that are absorbed in the ileum, together with measurement of total BA pool and hepatic BA neosynthesis, biliary, and fecal BA levels, allow a detailed description of the partitioning of BAs in the body with relevance to a series of pathogenic mechanisms for metabolic diseases.

BASIC PROTOCOL 6

Materials Mice Ketamine (Sigma-Aldrich) Xylazine (Bayer) 100 μM [3 H(G)] taurocholic acid (sp. act. 1 μCi/100 μl; American Radiolabeled Chemicals)

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Solvable sample solubilizer (Perkin-Elmer) 10% (v/v) H2 O2 Opti-Fluo (Perkin-Elmer) Surgical instruments including scissors and clamps Syringes with 26-G needles Tared 20-ml glass vials Scintillation counter Additional reagents and equipment for euthanasia of the mouse (Donovan and Brown, 2006) 1. Induce anesthesia with intraperitoneal administration (using a 26-G needle) of ketamine and xylazine (see Basic Protocol 5, step 2). Perform a median laparotomy and inject 100 μl of 100 μM [3 H(G)]taurocholic acid (1 μCi/100 μl) using a 26-G needle directly into the ileum (∼3 cm above the junction between small intestine and cecum). 2. Euthanize mouse by CO2 inhalation (Donovan and Brown, 2006) 15 min after [3 H] taurocholic acid administration. 3. Remove and segment the small intestine into duodenum, jejunum and ileum, and also the colon. 4. Place tissue samples in separate, tared 20-ml glass vials. 5. Add 1 ml/0.1 g tissue of Solvable to each vial. 6. Incubate 2 to 3 hr at 60◦ C. 7. Allow the vials to cool at room temperature. 8. Add 0.1 ml of 10% H2 O2 per 1-ml aliquot of Solvable or tissues. 9. Let stand at room temperature for 15 to 30 min. 10. Cap tightly the vials and incubate 1 hr at 60◦ C. 11. Allow the vials to cool to room temperature. 12. Add the scintillation fluid (4 ml of Opti-Fluo per 200 μl of each sample) and allow the samples to stand for at least 2 hr at room temperature before counting. 13. Quantify radioactivity in each sample using a scintillation counter. 14. Express the results as taurocholate (% of dose). BASIC PROTOCOL 7

INTESTINAL BACTERIAL COUNTS BAs have long been known to exhibit surfactant properties and to keep intestinal bacterial overgrowth under control. Moreover, it has been shown that part of their antibacterial effect is due to their activity as signaling molecules (Inagaki et al., 2006). The absence of BAs in the intestinal lumen (e.g., obstructive cholestasis) compromises the intestinal architecture and can allow bacterial translocation across the mucosa barrier and possible subsequent systemic infection. A well established animal model to reduce intestinal BA content and mimic human obstructive cholestasis consists of performing bile duct ligation (BDL) in rodents. At a time point 4 days after BDL, ileum, cecum, and mesenteric lymph node complex are harvested and processed as described below. After BDL, the content of bacteria is expected to dramatically increase due to absence of intraluminal BA content.

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ileum

jejunum

duodenum

cecum

small intestine

Figure 2

colon

Removal of various small intestinal segments from the mouse.

Materials Mice Trypticase soy broth (BD Difco; also see recipe in Reagents and Solutions) Trypticase soy agar plates with 5% sheep blood (BD Difco) Surgical instruments including scissors and clamps Balance Additional reagents and equipment for euthanasia of the mouse (Donovan and Brown, 2006) and anaerobic culture of bacteria (Speers et al., 2009) 1. Euthanize mouse by CO2 inhalation (Donovan and Brown, 2006) and remove ileum, cecum, and mesenteric lymph node complex (Fig. 2). 2. Using a syringe, inject, in one end of the lumen of the ileum, 1.5 ml of trypticase soy broth, and collect the contents from the opposite end. 3. Weigh the cecum and mesenteric lymph node complex. 4. Homogenize cecum and mesenteric lymph node complex with scissors in 1.5 ml of trypticase soy broth. 5. Make serial dilutions in trypticase soy broth of ileal contents and cecal and mesenteric lymph node complex extracts (for ileum try from 1:10 to 1:103 , for cecum at least 1:105 , for lymph node complex try from 1:10 to 1:102 ). 6. Plate dilutions on trypticase soy agar plates with 5% sheep blood and culture at 37◦ C under aerobic and anaerobic conditions. Anaerobic culture is described in Speers et al. (2009).

7. After 20 hr incubation, count the number of colony-forming units (cfu). 8. Express the results as cfu/g of ileum, cecum, or mesenteric lymph node complex One calculation is made for the aerobic condition, and one calculation is made for the anaerobic condition. Characterizing Bile Acid and Lipid Metabolism

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BASIC PROTOCOL 8

MEASUREMENT OF SERUM AND HEPATIC TRIGLYCERIDES Triglycerides are esters of fatty acids and are hydrolyzed to glycerol and free fatty acids. Triglyceride determinations, when performed in conjugation with other lipid assays, are useful in the diagnosis of primary and secondary hyperlipoproteinemia. They are also of interest in following the course of diabetes mellitus, nephrosis, biliary obstruction, and various metabolic abnormalities arising from endocrine disturbances. Standard methods for the determination of triglyceride concentrations have involved either enzymatic or alkaline hydrolysis to release glycerol. This protocol makes use of the enzymatic hydrolysis and quantification since it is specific and not subject to interference by phospholipids (Searcy, 1969). The enzymatic reaction sequence employed in the assay of triglycerides is as follows: Triglycerides → Glycerol + Fatty Acids (catalyzed by lipase) Glycerol + ATP → Glycerolphosphate +ADP (catalyzed by glycerol kinase) Glycerol-1-Phosphate + O2 → DAP + H2 O2 (catalyzed by glycerol-3-phosphate oxidase; GPO) H2 O2 + 4-Aminoantipyrine + DHBS → Quinoneimine Dye + 2H2 O The procedure described here involves triglyceride hydrolysis by lipase. The glycerol concentration is then determined by enzymatic assay coupled with Trinder reaction that terminates in the formation of a quinoneimine dye. The amount of the dye formed, determined by its absorption at 520 nm, is directly proportional to the concentration of triglycerides in the samples (McGowan et al., 1983). Among the current commercial available kits, the authors have good experience with the Colorimetric Triglycerides Liquid Kit from Sentinel due to absence of interference from hemolytic (hemoglobin < 0.25 g/dl) and jaundiced (total bilirubin < 40 mg/dl) samples, in addition to its high linear range (2 to 1000 mg/dl).

Materials Triglyceride-containing samples Colorimetric Triglycerides Liquid Kit (containing Reagent 1 and Standard; Sentinel, http://www.sentinel.it); see recipe for Triglyceride reagents in Reagents and Solutions for compositions of items Methanol Triton X-100 1.5-ml cuvettes or 96-well plates (300-μl well volume) Spectrophotometer at 546 nm 1. Pipet 10 μl of sample, standard (from kit), or blank (distilled water for serum triglycerides, 10% Triton X-100 in methanol for hepatic triglycerides) per well of a 96-well plate. If cuvettes are used, pipet 10 μl/cuvette.

2. Into each well containing sample, standard, or blank (from step 1), pipet 200 μl/well of Reagent 1. Incubate 5 min at room temperature in the dark. 3. Zero the spectrophotometer with the blank at 546 nm. Read and record absorbance of samples (wavelength range: 540 to 560 nm; scan to find wavelength of maximum absorbance). Characterizing Bile Acid and Lipid Metabolism

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4. Calculate triglyceride concentration using the following formula: (sample A546 /standard A546 ) × conc. of standard where 200 mg/dl is the concentration of the standard. The final color developed is stable up to 30 min in the dark. Linearity is between 2 and 1000 mg/dl. If the values exceed 1000 mg/dl, the test must be repeated by diluting the samples with blank. One must then multiply by the dilution factor.

MEASUREMENT OF SERUM, BILIARY, AND HEPATIC CHOLESTEROL Cholesterol serves as a precursor of BAs, steroids, and vitamin D. The determination of cholesterol is a major aid in the diagnosis and classification of lipemias, cholesterol gallstone disease (CGD), and other conditions such as hepatic thyroid diseases.

BASIC PROTOCOL 9

Enzymatic methods have replaced older methodologies involving cholesterol esterase, cholesterol oxidase, and Trinders color methods. Allain et al. (1974) developed a total enzymatic technique in which hydrogen peroxide produced during the oxidation of cholesterol is used in conjugation with peroxidase, 4-aminoantipyrine, and phenol to form a quinoneimine dye. The enzymatic reaction sequence employed in the assay of cholesterol is as follows: Cholesterol Esters → Cholesterol + Fatty Acids (catalyzed by cholesterol esterase) Cholesterol + O2 → Cholesten-3-one + H2 O2 (catalyzed by cholesterol oxidase) 2H2 O2 + 4-Aminoantipyrine + hydroxybenzoic acid → Quinoneimine + 2H2 O (catalyzed by peroxidase) Cholesterol esters are hydrolyzed to produce cholesterol. Hydrogen peroxide is then produced from the oxidation of cholesterol by cholesterol oxidase. In a coupled reaction catalyzed by peroxidase, red-colored quinoneimine dye is formed from 4-aminoantipyrine, hydroxybenzoic acid, and hydrogen peroxide. The absorption of dye this solution at 510 nm is proportional to the cholesterol concentration in the sample. Among the current commercial available kits, the authors have good experience with the Colorimentric Cholesterol Liquid Kit from Sentinel, due to absence of interference from lipemic (triglycerides < 1000 mg/dl), hemolytic (hemoglobin < 0.5 g/dl), and jaundiced (total bilirubin < 15 mg/dl) samples, in addition to its high linear range (4 to 700 mg/dl).

Materials Cholesterol-containing samples Colorimetric Cholesterol Liquid Kit (containing Reagent 1 and Standard; Sentinel, http://www.sentinel.it); see recipe for Cholesterol reagents in Reagents and Solutions for compositions of items Methanol Triton X-100 1.5-ml cuvettes or 96-well plates (300-μl well volume) Spectrophotometer at 546 nm 1. Into wells of a 96-well plate place 10 μl of sample, standard (from kit), or blank (distilled water for serum and biliary cholesterol, 10% Triton X-100 in methanol for hepatic cholesterol). If cuvettes are used, pipet 10 μl/cuvette.

2. Into each well containing sample, standard, or blank (from step 1), pipet 200 μl of Reagent 1. Incubate 5 min at room temperature in the dark.

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3. Zero the spectrophotometer with the blank at 546 nm. Read and record absorbance of samples (wavelength range: 500 to 546 nm; scan to find wavelength of maximum absorbance). 4. Calculate cholesterol concentration using the following formula: (sample A510 /standard A510 ) × conc. of standard where 200 mg/dl is the concentration of the standard. The final color developed is stable up to 1 hr in the dark. Linearity is between 4 and 700 mg/dl. If the values exceed 700 mg/dl, the test must be repeated by diluting the samples with blank. One must then multiply by the dilution factor. BASIC PROTOCOL 10

MEASUREMENT OF BILIARY, HEPATIC, AND SERUM PHOSPHOLIPIDS Phospholipids are a class of lipids that constitute a major component of cell membranes and play important roles in signal transduction. Moreover, phospholipids play a role in the emulsification and absorption of fat in the body. The determination of phospholipids in bile and serum is an important index of the hepatic function. High levels of serum phospholipids are present in Niemann-Pick disease, while low levels of phospholipids in bile are a consequence of progressive familial intra-hepatic cholestasis type 3 (PFIC3). Most phospholipids contain one diglyceride, a phosphate group, and one choline. In the past, methods used to measure phospholipids required extractions with organic solvents followed by acid digestion to release the phosphorus, which would then be measured by colorimetric methods. Such methods were complicated and not easily adaptable to automation. In the method presented here, when a sample is added to the Color Reagent of the Colorimetric Phospholipids Kit (Sentinel), phospholipids are hydrolyzed by phospholipase D to produce choline, which in turn is oxidized by choline oxidase to betaine and hydrogen peroxide. The hydrogen peroxide produced causes N-ethylN-(2-hydroxy-3-sulfopropyl)-3,5-dimethoxyaniline (DAOS) and 4-aminoantipyrine to undergo a quantitative oxidative condensation catalyzed by peroxidase (POD), producing a color whose absorbance is proportional to the amount of phospholipids in the sample. The enzymatic reaction sequence employed in the assay of phospholipids is as follows: Phospholipids + H2 O → Choline + Phosphatitic acid (catalyzed by phospholipase D) Choline → Betaine + H2 O2 (catalyzed by choline oxidase) 2H2 O2 + 4-Aminoantipyrine + HB → Quinoneimine + 2H2 O (catalyzed by peroxidase) Among the current commercial available kits, the authors have good experience with the Colorimentric Phospholipids Kit from Sentinel, due to absence of interference from lipemic (triglycerides < 1000 mg/dl), hemolytic (hemoglobin < 0.5 g/dl), and jaundiced (total bilirubin < 25 mg/dl) samples, in addition to its high linear range (5 to 1000 mg/dl).

Material Phospholipid-containing samples Colorimetric Phospholipids Kit (containing Reagent 1a, Reagent 1b, and Standard; Sentinel, http://www.sentinel.it); see recipe for Phospholipid reagents in Reagents and Solutions for descriptions of items Methanol Triton X-100 Characterizing Bile Acid and Lipid Metabolism

96-well plates Spectrophotometer at 520 nm

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1. Add 10 ml of Reagent 1a to Reagent 1b (items from kit) and wait for 10 min. 2. Pipet 10 μl of sample, standard, or blank (distilled water for biliary and serum phospholipids, 10% Triton X-100 in methanol for hepatic phospholipids) into a 96-well plate. 3. Into each well containing sample, standards, or blank (from step 2), pipet 200 μl of the reconstituted Reagent 1b from step 1. 4. Incubate 10 min at room temperature. 5. Zero the spectrophotometer with the blank at 520 nm. Read and record absorbance of samples (wavelength range: 510-530 nm). 6. Calculate phospholipid concentration using the following formula: (sample A520 / standard A520 ) × conc. of standard where 300 mg/dl is the concentration of the standard. The final color developed is stable up to 30 min. Linearity is between 5 and 1000 mg/dl. If the values exceed 1000 mg/dl, the test must be repeated by diluting the samples with blank. Then multiply to account for the dilution factor.

HEPATIC LIPID EXTRACTION The Folch method (Folch et al., 1957), based on a chloroform/methanol extraction, is still used today for the extraction of lipids from fresh or frozen tissue. Liver samples are homogenized in chloroform/methanol to extract lipids into the organic phase. Then, after evaporation of the organic phase, lipids are resuspended in 10% Triton X-100 in methanol. Cholesterol, triglycerides, and phospholipids can then be determined with the same assays reported above used to measure these lipids in the serum.

BASIC PROTOCOL 11

NOTE: Avoid plastic tubes because of the possibility of solvents leaching out and contaminating lipids. Use caps to avoid evaporation of solvents.

Materials Mouse liver tissue 2:1 (v/v) chloroform/methanol 50 mM NaCl 0.36 M CaCl2 in methanol Chloroform Nitrogen source 10% (v/v) Triton X-100 in methanol 5-ml volumetric glass tubes Polytron homogenizer Centrifuge 1. To 100 to 200 mg of liver tissue add 4 ml of 2:1 (v/v) chloroform/methanol. Homogenize with a Polytron for 30 sec at room temperature 2. Wash by adding 1 ml of 50 mM NaCl, vortex, centrifuge 10 min at 1500 × g, room temperature, and carefully remove organic phase and transfer into a new glass tube. 3. Wash organic phase with 1 ml of 0.36 M CaCl2 in methanol, vortex, and centrifuge 10 min at 1500 × g, room temperature. Carefully remove organic phase and transfer into a 5-ml volumetric glass tube. Bring sample volume to 5 ml with chloroform. If you wish to stop here, store samples at −20o C.

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4. Evaporate 500 μl of the sample from step 3 to dryness under nitrogen gas at room temperature. Dissolve the pellet in 200 μl of 10% Triton X-100 in methanol. 5. Measure cholesterol, triglycerides, and phospholipids as described in Basic Protocols 8, 9, and 10. BASIC PROTOCOL 12

INTESTINAL CHOLESTEROL ABSORPTION After a meal, BAs and phospholipids are secreted from the gallbladder into the duodenum in the form of mixed micelles, which are important to allow intestinal cholesterol absorption. When BA or phospholipid concentrations in the intestine decrease due to defects in their secretion, problems with absorption of cholesterol and other fat dietary lipids may arise (Turley et al., 1994). This protocol allows the investigator to obtain an estimate of the intestinal cholesterol absorption fraction over the total circulating cholesterol pool.

Materials Mice Dosing mixture of labeled cholesterol and stigmastanol (see recipe) Soluene 350 (Perkin Elmer) 2-propanol 30% (v/v) hydrogen peroxide UltimaGold scintillation fluid (Perkin Elmer) Animal balance Gavage needle for mice Mortar and pestle 20-ml scintillation vial Heat block β-scintillation counter 1. Weigh mice before the assay. 2. Administer via gavage 100 μl of a dosing mixture containing 0.02 μCi [14 C] cholesterol/g body weight and 0.04 μCi [3 H] stigmastanol/g body weight (see Reagents and Solutions). Store 60 μl final dosing mixture for measuring radioactivity (standards). Therefore, always prepare ∼30% excess dosing mixture than is necessary for the number of mice in the experiment.

3. Collect fecal samples for 48 hr after radiolabeled cholesterol administration IMPORTANT NOTE: Collect cold (nonradioactive) feces from a control mouse and process simultaneously.

4. Place feces in 100-ml glass beaker and dry in air until weight no longer changes (∼1 to 2 days). Grind feces to a fine powder using a mortar and pestle. 5. In a 20-ml scintillation vial, add 0.1 ml water to 20 mg of feces and rehydrate for 30 min. 6. Add 1 ml Soluene 350, vortex, and incubate 2 hr at 50◦ C (vortex every 30 min). 7. Add 1 ml of 2-propanol and incubate for 2 hr at 50◦ C (vortex every 30 min). 8. Add 0.2 ml of 30% hydrogen peroxide dropwise and gently agitate. Allow to stand for 30 min at room temperature. Characterizing Bile Acid and Lipid Metabolism

9. Add 20 μl of the original mixture (step 2) only to the cold stool control.

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10. Add 20 ml UltimaGold and count vials on 3 H/14 C channel after overnight room temperature and light adaptation. 11. Calculate the % of cholesterol absorption using this equation:

( ⎡⎣

14

)

(

)

H ⎤⎦ ⎡⎣ 3 H ⎤⎦ dosing mixture − ⎡⎣ 14 H ⎤⎦ ⎡⎣ 3 H ⎤⎦ feces × 100 ⎡⎣ 14 H ⎤⎦ ⎡⎣ 3 H ⎤⎦ dosing mixture

(

)

MEASUREMENT OF IN VIVO FATTY ACID SYNTHESIS It has been shown that BAs, as signaling molecules, can also regulate lipid metabolism. In particular, by modulating the expression of the lipogenic transcription factor sterol regulatory element binding protein 1c (SREPB1c), BAs can impact fatty acid synthesis (Watanabe et al., 2004). This protocol describes the in vivo measurement of fatty acid synthesis.

BASIC PROTOCOL 13

Material Mice [3 H]2 O (tritiated water; American Radiolabeled Chemicals) 1 M KOH in 66% (v/v) ethanol Petroleum ether Concentrated HCl Hexane Methanol Scintillation cocktail Syringe and 26-G needles Animal balance Surgical instruments including scissors and clamps 50-ml glass tubes Heat block 50- or 100-ml volumetric flask Scintillation vials 80◦ C vacuum oven β-scintillation counter Additional reagents and equipment for euthanasia of the mouse (Donovan and Brown, 2006) 1. Inject mice intraperitoneally with 20 to 50 mCi of [3 H]2 O. 2. Place injected mice in individual plastic cages and euthanize them by CO2 inhalation (Donovan and Brown, 2006) exactly 1 hr later. 3. Determine weights of extracted tissues for all samples. For the liver, weigh the entire tissue and then cut two pieces of 250 to 300 mg for each sample. Tissues such as kidney or brain can also be harvested for controls.

4. Put each sample in a 50-ml glass tube and add 5 ml of 1 M KOH in 66% ethanol. 5. Incubate the tubes at 65◦ C in a heat block until samples are dissolved (this can take several hours). Vortex samples every 30 min.

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6. Once tissues are completely dissolved, add exactly 5 ml of distilled water and 15 ml of petroleum ether. Shake tubes for 1 min to extract sterols. Let stand at least 30 min to allow the complete separation of the organic and water phase. The organic phase, on the top, is cholesterol enriched, while the water phase, is enriched with fatty acids.

7. Remove and discard top phase, which contains labeled cholesterol. 8. Add 1 ml concentrated HCl to the water phase. 9. Add 15 ml hexane and vortex vigorously for 1 min. Allow sample to separate into two phases. Small intestine and carcass (rest of mouse body once organs mentioned above have been removed) may need to be centrifuged 5 min at 800 × g. If the bottom phase is cloudy, it must be allowed to sit longer or be centrifuged.

10. Transfer hexane layer from each tube into a 50- or 100-ml volumetric flask. 11. Add another 15 ml of hexane to each tube and repeat step 9. 12. Bring volumetric flask up to volume with hexane and invert several times to mix. 13. Remove 10 ml hexane from 100-ml volumetric flask (or 5 ml from 50-ml flask) and place it in a numbered scintillation vial. 14. Allow the hexane to evaporate overnight in a chemical fume hood. 15. Heat uncapped vials in a vacuum oven 30 min at 80◦ C. 16. Add 1 ml methanol and 15 ml scintillation counting cocktail. Count in scintillation counter. 17. Use the radioactivity counts and the tissue weight to calculate pmol/mg. 18. Express the rate of fatty acid synthesis as pmol/hr/g tissue.

SERUM TRANSAMINASES (AST, ALT, ALP) AST Serum aspartate aminotransferase (AST), also known as serum glutamic oxalacetic transaminase (SGOT), is an enzyme that catalyzes the exchange of amino and keto groups of alpha amino acids and alpha keto acids. AST is widely distributed in tissues such as the heart and liver. Injury to these tissues causes release of the AST enzyme into the general circulation. Following a myocardial infarction, serum levels of AST are elevated and reach a peak 48 to 60 hr after onset. Hepatobiliary diseases such as cirrhosis, metastatic carcinoma, and viral hepatitis also increase serum AST levels. Earlier colorimetric methods for the determination of serum AST were based on the reaction of oxaloacetate with dinitrophenylhydrazine (Teitz, 1976). However, this reaction is time consuming and nonspecific. The method described here is based on a modification of the colorimetric method of Doumas and Briggs (1969). AST catalyzes the following reaction: L-Aspartate

+ 2-Oxoglutarate → Oxalacetate + L-Glutamate

In the present method (Basic Protocol 15), a diazonium salt is used. It selectively reacts with the oxalacetate to produce a color complex that is measured photometrically. Characterizing Bile Acid and Lipid Metabolism

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ALT Serum alanine aminotransferase (ALT), also known as glutamate pyruvate transaminase (SGPT), is present in a variety of tissues. The major source of elevated serum levels of ALT is the liver, which has led to the introduction of ALT determination in the diagnosis of hepatic diseases. Elevated serum ALT levels are found in hepatitis, cirrhosis, and obstructive jaundice. Levels of ALT are only slightly elevated in patients following a myocardial infarction. Since 1995, many methods and modifications of methods have been proposed for the determination of ALT. Herein we describe a colorimetric method (Basic Protocol 14) based on dinitrophenylhydrazine formation, where ALT catalyzes the following reaction: L-Alanine

+ α-Ketoglutarate → Pyruvate + Glutamate

Pyruvate + 2,4-DNPH → Pyruvate + 2,4-DNPH-one (in the presence of H+ ) This is a modification of the classical Reitman-Frankel colorimetric endpoint reaction (Reitman and Frankel, 1957). In this procedure, ALT catalyzes L-alanine and α-ketoglutarate to form pyruvate and glutamate. Pyruvate then reacts with 2,4dinitrophenylhydrazine (2,4-DNPH) to form 2,4-DNPH-one. The addition of sodium hydroxide dissolves this complex, allowing 2,4-DNPH-one to be measured at 505 nm.

ALP The alkaline phosphatase (ALP) enzyme is distributed in almost every tissue of the body. Serum ALP levels are of interest in the diagnosis of hepatobiliary disorders and bone diseases. Most of the ALP in normal adult serum is from the liver or biliary tract. Normal ALP levels are age dependent, and levels are elevated during periods of active bone growth. Moderate elevations of ALP (not involving the liver or the bone) may be attributed to Hodgkin’s disease, congestive heart failure, and abdominal bacterial infections. Elevations also occur in the third trimester of pregnancy. Previous ALP assays were based on the measurement of phosphate liberated by the action of the enzyme on a β-glycerolphosphate substrate or on the measurement of phenol liberated from disodium phenyl phosphate substrate. Many of these substrates are instable in solution and need to be prepared fresh daily. The substrate prepared by Roy, which uses sodium thymolphthalein monophosphate, is stable up to 1 year when properly stored (Roy, 1970). In the method reported here (Basic Protocol 16), ALP acts upon the AMP-buffered sodium thymolphthalein monophosphate. The addition of an alkaline reagent stops enzyme activity and simultaneously develops a blue chromogen, which is measured photometrically. Today, different commercial kits are available for measuring transaminases. Here, for convenience, kits from BioQuant are used for manual measurement of transaminase activities.

Determination of Serum Alanine Aminotransferase (ALT) Activity Materials Serum sample Colorimetric ALT (SGPT) kit (containing ALT substrate, ALT color reagent, ALT color developer, ALT calibrator; BioQuant, http://www.bio-quant.com); see recipe for ALT reagents in Reagents and Solutions for compositions of items

BASIC PROTOCOL 14

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2-ml test tubes with rack Heat block Spectrophotometer with detection at 505 nm 1. Label 2-ml test tubes as blank (distilled water), calibrator, or sample. 2. Transfer 250 μl of ALT substrate to each tube and incubate 3 to 5 min at 37◦ C. 3. At time intervals (∼15 to 30 sec), add 100 μl of sample (dilute unknown samples if required) to the corresponding labeled tubes. Mix and immediately return to 37◦ C heating bath for exactly 30 min. 4. After exactly 30 min, add 250 μl of ALT color regent to each tube, maintaining the time interval sequence. Mix and return to 37◦ C heating bath for exactly 10 min. 5. After exactly 10 min, add 1 ml ALT color developer, maintaining the same time interval. Mix and return to 37◦ C heating bath for 5 min. 6. Zero the spectrophotometer with the reagent “blank” at 505 nm. Read and record absorbance of all tubes (wavelength range: 500 to 520 nm). The final colored product is stable for 60 min at room temperature.

7. Calculate the ALT activity using the following formula: (sample A505 / calibrator A505 ) × conc. of calibrator (IU/liter) where 70 IU/liter is the activity of the calibrator. The final color developed in the reaction must be read within 60 min. Linearity is up to 120 IU/liter. If the values exceed 120 IU/liter, the test must be repeated by diluting the samples with blank. One must then multiply to account for the dilution factor. BASIC PROTOCOL 15

Determination of Serum Aspartate Aminotransferase (AST) Activity Materials Serum sample Colorimetric AST (SGOT) kit (containing AST substrate, AST color reagent, AST calibrator; BioQuant, http//www.bio-quant.com); see recipe for AST reagents in Reagents and Solutions for compositions of items 0.1 N HCl 2 ml test tubes with rack Spectrophotometer with detection at 530 nm 1. Label 2-ml test tubes as blank (distilled water), calibrator, or sample. 2. Transfer 250 μl of AST substrate to each tube and incubate at least 4 min at 37◦ C 3. At time intervals (∼15 to 30 sec), add 100 μl of sample (dilute unknown samples if required) to the corresponding labeled tubes. Mix and immediately return to 37◦ C for exactly 10 min. 4. After exactly 10 min, add 250 μl of AST color regent to each tube, maintaining the time interval sequence. Mix and return to 37◦ C heating bath for exactly 10 min. 5. After exactly 10 min, add 1 ml of 0.1 N hydrochloric acid and mix by inversion.

Characterizing Bile Acid and Lipid Metabolism

6. Zero the spectrophotometer with the reagent “blank” at 530 nm. Read and record absorbance of all tubes (wavelength range: 500 to 550 nm). The final colored product is stable for 60 min at room temperature.

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7. Calculate the AST activity using the following formula: (sample A530 /calibrator A530 ) × conc. of calibrator (IU/liter) where 88 IU/liter is the activity of the calibrator. The final color developed in the reaction must be read within 60 min. Linearity is up to 500 IU/liter. If the values exceed 500 IU/liter, the test must be repeated by diluting the samples with blank. One must then multiply to account for the dilution factor.

Determination of Serum Alkaline Phosphatase (ALP) Activity Materials

BASIC PROTOCOL 16

Serum sample Colorimetric ALP kit (containing ALP substrate, ALP color developer, ALP standard; BioQuant, http://www.bio-quant.com); see recipe for ALP reagents in Reagents and Solutions for compositions of items 2-ml test tubes with rack Spectrophotometer detecting at 590 nm 1. Label 2-ml test tubes as blank (distilled water), calibrator, or sample. 2. Dispense 250 μl of ALP substrate into each tube and equilibrate 3 min at 37◦ C. 3. At time intervals (∼15 to 30 sec), add 50 μl of blank, standard, and sample to the respective labeled tubes. Mix gently. 4. Incubate for exactly 10 min at 37◦ C. 5. Following the same time interval sequence as in step 3, add 1.25 ml of ALP color developer at time intervals. Mix well. 6. Set the wavelength of the spectrophotometer to 590 nm. Zero with reagent blank (wavelength range: 580 to 630 nm). 7. Read and record the sample absorbance. The final colored product is stable for 60 min at room temperature.

8. Calculate the ALP activity using the following formula: (sample A590 / calibrator A590 ) × conc. of calibrator (IU/liter) where 50 IU/liter is the activity of the calibrator. The final color developed in the reaction must be read within 60 min. Linearity is up to100 IU/liter. If the values exceed 100 IU/liter, repeat the test after diluting the samples with blank. One must then multiply to account for the dilution factor.

MEASUREMENT OF SERUM AND BILIARY TOTAL BILIRUBIN Bilirubin is a metabolite of heme, mainly from breakdown of hemoglobin. Normally, bilirubin is excreted into the intestine from the liver via the bile. The site of the catabolism of hemoglobin is the reticuloendothelial system (RES) in bone marrow, liver, and spleen. Bilirubin is then released into the bloodstream where it binds tightly to albumin (this fraction is referred to as indirect bilirubin) and is transported to the liver. Upon uptake by the liver, bilirubin is conjugated with glucuronic acid to form bilirubin mono- and diglucuronide, which are water-soluble metabolites. The metabolites will react with aqueous diazo reagent and are commonly referred to as “direct bilirubin” (Teitz, 1976). The sum of the indirect and direct bilirubin is referred to as total bilirubin.

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Elevation of total serum bilirubin may occur due to excessive hemolysis or destruction of the red blood cells (e.g., hemolytic disease of the newborn), liver diseases (e.g., hepatitis and cirrhosis), or obstruction of the biliary tract (e.g., gallstones). Thus, determination of total bilirubin levels is useful in the diagnosis and management of hemolytic disorders and liver diseases, including biliary obstruction, cholestasis, hepatitis, jaundice, and cirrhosis. Notably, elevated levels of direct bilirubin in patients with liver or biliary-tract disease have been reported, although total bilirubin levels were normal. Therefore, the greatest diagnostic value of direct bilirubin assays is their ability to detect occult liver disease. The methods most widely employed for the determination of total bilirubin are the diazo coupling method and the bilirubin oxidase enzymatic method (Gambino et al., 1967). However, these methods have disadvantages such as interference from coexistent serum substances and unsatisfactory stability of reagents after preparation. Here, a method is presented for bilirubin quantification based on chemical oxidation, utilizing vanadate as an oxidizing agent. This procedure shows good correlation with conventional methods and no interference by coexistent serum substances, and uses a convenient, ready-to-use liquid reagent. When the sample is mixed with the reagent containing the detergent and the vanadate, at pH 3, total bilirubin in the sample is oxidized to biliverdin. This causes the reduction of yellow absorbance, which is specific to bilirubin. Therefore, the total bilirubin concentration in the sample can be obtained by measuring the absorbances before and after the vanadate oxidation.

Materials Serum or bile sample Colorimetric Bilirubin Total assay kit (containing Reagent R1, Reagent R2, and calibrator; Diazyme, (http://www.diazyme.com); see recipe for total bilirubin reagents in Reagents and Solutions for compositions of items Cuvettes or 96-well plates Spectrophotometer detecting at 450 nm 1. Place 10 μl of sample, calibrator, or water (blank) into a cuvette or into wells of a 96-well plate. 2. Pipet 280 μl of Reagent R1 into the cuvette. If using a 96-well plate, pipet 200 μl/well Reagent R1. 3. Incubate 5 min at 37◦ C. 4. Read and record the absorbance at 450 nm. 5. Pipet 70 μl of Reagent R2 into cuvette (if using a 96-well plate, pipet 50 μl) and incubate 5 min at 37◦ C. 6. Read and record the absorbance at 450 nm. 7. Calculate the total bilirubin concentration using the following formula: [S450 (sample) – S450 (blank)] / [S450 (calibrator) – S450 (blank)] × conc. calibrator where S450 is the difference between the absorbance recorded at 450 nm after 5 and 10 min. The concentration of the calibrator is 12.8 mg/dl. The linearity of this procedure is from 0.1 to 40 mg/dl. When total bilirubin concentration exceeds 40 mg/dl, dilute the sample with saline, repeat the assay, and multiply the result by the dilution factor. Characterizing Bile Acid and Lipid Metabolism

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REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps.

ALP reagents Items in kit from BioQuant (http://www.bio-quant.com): ALP SUBSTRATE (may cause skin irritation!): 3.6 mM sodium thymolphthalein monophosphate in 0.2 M 2-amino-2-methyl-1-propanol buffer. Also contains 1.0 mM MgCl2 , wetting agent, inactive ingredients, preservatives; pH 10.2±0.1 ALP COLOR DEVELOPER (causes burns!): 0.1 M sodium hydroxide, 0.1 M sodium carbonate. ALP STANDARD: 0.5 mM thymolphthalein in n-propanol. Equivalent to 50 U/liter enzyme activity when used according to the ALP procedure All reagents are ready to use and stable until the expiration date. Store ALP substrate, color developer, and standard at 2◦ to 8◦ C. ALT substrate should be a clear amber solution. A precipitation or blue-green color would indicate deterioration. ALP color developer should be a clear colorless solution.

ALT reagents Items in kit from BioQuant (http://www.bio-quant.com): ALT SUBSTRATE: 0.2 M L-alanine, 2.0 mM α-ketoglutarate, 100 mM phosphate buffer, pH 7.4±0.05, 0.2% (v/v) preservatives. ALT COLOR REAGENT (cause burns!): 1.0 mM 2,4 dinitrophenylhydrazine in 1 N hydrochloric acid, preservative. ALT COLOR DEVELOPER (corrosive!): 0.5 N sodium hydroxide. ALT CALIBRATOR: solution of sodium pyruvate in 100 mM phosphate buffer at pH 7.4. All reagents are ready to use and stable until the expiration date. Store ALT substrate, color developer, and standard at 2◦ to 8◦ C. If turbidity and precipitation occur, this may be sign of reagent deterioration.

AST reagent Items in kit from BioQuant (http://www.bio-quant.com): AST SUBSTRATE: 33 mM aspartic acid, 5 mM ketoglutaric acid, phosphate buffer, pH 7.4. AST COLOR REAGENT: 0.25% w/v diazonium salt preserved with formalin AST CALIBRATOR: a lyophilized serum with AST value provided in each lot SUBSTRATE reagent is ready to use. Reconstitute COLOR REAGENT and calibrator with 30 ml and 5 ml of distilled water, respectively. Let stand until dissolved and swirl to mix. Calibrator is stable up to 5 days at 2◦ to 8◦ C after reconstitution. Store AST SUBSTRATE, COLOR REAGENT, and CALIBRATOR at 2◦ to 8◦ C. The AST SUBSTRATE should be a clear colorless solution. Reagents should be discarded if turbidity or discoloration is noted. If AST COLOR REAGENT darkens or if dark brown precipitate is visible, do not use.

Bile acid standards and internal standards Purchase the following standard compounds from Sigma-Aldrich: Sodium taurocholate Sodium taurodeoxycholate Sodium taurochenodeoxycholate Sodium taurolithocholate Sodium glycocholate Sodium glycodeoxycholate continued

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Sodium glycochendeoxycholate Cholic acid Deoxycholic acid Chenodeoxycholic acid Hyodeoxycholic acid Ursodeoxycholic acid Lithocholic acid Sodium tauroursodeoxycholate Sodium taurohyodeoxycholate Purchase tauro-β-muricholic acid from Steraloids (http://www.steraloids.com) Purchase the following deuterated internal standards: D4 -glycocholic acid (IS1) from CDN isotopes (https://www.cdnisotopes.com/) D4 -cholic acid (IS2) from Steraloids (http://www.steraloids.com) Preparation of standard solutions: Prepare standard stock solutions of BAs and internal standards in HPLC-grade ethanol at a concentration of 1 mg/ml. Prepare standard working solutions of BAs (in the range of 0.01 to 10 μg/ml by dilution of their stock solutions with HPLC-grade ethanol. Prepare internal standard working solutions (100 μg/ml for IS1 and 10 μg/ml for IS2) by dilution of their stock solutions with HPLC-grade methanol. Store all standard solutions indefinitely at –20◦ C. Cholesterol reagents Items in kit from Sentinel (http://www.sentinel.it): REAGENT 1: Good’s buffer, 50 mmol/liter, pH 6.7; cholesterol oxidase, ≥ 300 U/liter; cholesterol esterase, ≥ 300 U/liter; hydroxybenzoic acid, 12 mmol/liter; 4-aminoantipyrine, 0.3 mmol/liter; peroxidase, ≥ 10 kU/liter; sodium azide, < 0.1%. STANDARD: cholesterol standard, 200 mg/dl (4.17 mmol/liter). REAGENT 1 is ready to use and stable up to 90 days at 2◦ to 8◦ C after opening. Standard is ready to use and stable up to 120 days at 2◦ to 8◦ C after opening. A slight pink coloration of REAGENT 1 does not affect the reagent performance.

Dosing mixture If 30 mice are being used for the experiment and an average of 25 g per mouse is assumed, then prepare: [14 C] cholesterol (American Radiolabeled Chemicals): 0.02 μCi/g × 25 g/mouse × (30 mice + 10) = 20 μCi total. [3 H] stigmastanol (American Radiolabeled Chemicals): 2 × 20 μCi = 40 μCi total. 20 μCi = 500 μl × 0.04 mCi [14 C] cholesterol (specific activity, 50 Ci/mmol). 40 μCi = 40 μl × 1 mCi/ml [3 H] stigmastanol (specific activity, 50 Ci/mmol) Mix, dry under nitrogen gas. Redissolve in 60 μl ethanol (1/10th volume of cholesterol + stigmastanol). Vortex 30 sec. Add 3.3 to 3.4 ml MCT oil (MEAD Johnson; http://mjn.com) so that: Characterizing Bile Acid and Lipid Metabolism

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final concentration of [14 C]cholesterol = 20 μCi/(540+60+3300) μl = ∼0.005 μCi/μl. final concentration of [3 H]stigmastanol = 40 μCi/(540+60+3300) μl = ∼0.01 μCi/μl. So now, 100 μl mixture corresponds to 0.5 μCi [14 C] cholesterol and 1 μCi [3 H] stigmastanol. Vortex 3 min. Prepare standards: put 10 to 20 μl of mixture into each of two to three beakers containing 50 ml Folch reagent (1:2 chloroform:methanol) to be extracted alongside samples.

Phospholipid reagents Items in kit from Sentinel (http://www.sentinel.it): REAGENT 1a: TES, 50 mmol/liter, pH 7.6; 4-hydroxybenzoic acid, 12 mmol/liter; EDTA, 1.3 mmol/liter; sodium azide, <0.1%; detergents. REAGENT1b (lyophilized): TES, 50 mmol/liter; phospholipase D, >1500 U/liter; choline oxidase, >7500 U/liter; 4-aminoantipyrine, 1.2 mmol/liter; peroxidase, >7000 U/liter. STANDARD: phospholipids standard, 300 mg/dl (3.87 mmol/liter); sodium azide, <0.1%; detergents Add 10 ml of REAGENT 1a to a bottle of lyophilized reagent 1b. Mix gently and wait for 10 min. Reconstituted REAGENT 1b is stable up to 14 days at 2o to 8o C. Standard is stable up to 120 days at 2o to 8o C after opening. A slight coloration or turbidity of the STANDARD will not influence the reagent performance. A slight pink coloration of the solution R1 will not influence the reagent performance.

Total bile acid reagents Items in kit from Diazyme (http://www. diazyme.com): REAGENT R1: Thio-NAD buffer. REAGENT R2: 3-α-HSD, buffer. CALIBRATOR: conjugated cholic acids, buffer. Reagents are ready to use. Unopened reagents are stable up to the expiration date printed on the label. Reagents are light sensitive and should be stored at 2o to 8o C. Reagents from different lots must not be interchanged.

Total bilirubin reagents Items in kit from Diazyme (http://www. diazyme.com): REAGENT R1 (buffer solution): citrate buffer, 0.1 mol/liter, pH 2.9; detergent. REAGENT R2 (vanadate solution): phosphate buffer, 10 mmol/liter, pH 7.0; sodium metavanadate, 4 mmol/liter. CALIBRATOR (lyophilized): bilirubin from bovine bile, ditaurobilirubin, and 0.005% (when reconstituted) sodium azide Reagents are ready to use. Unopened reagents are stable until the expiration date printed on the label. Reagents are light sensitive and should be stored at 2o to 8o C. The lyophilized calibrator is stable until the expiration date marked when stored at 2o to 10o C. The reconstituted calibrator (with 3 ml of distilled water) is stable up to 5 days at 2o to 10o C under protection from light.

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Triglyceride reagents Items in kit from Sentinel (http://www.sentinel.it): REAGENT 1: Good’s buffer, 50 mmol/liter, pH 6.7; ATP, 1.1 mmol; GK, ≥1000 U/liter; GPO, ≥2000 U/liters; POD, ≥2500 U/liter; LPL, ≥1500; 4aminoantipyrine, 0.2 mmol/liter; TOOS, 0.6 mmol/liter; sodium azide <0.1%. STANDARD: triglycerides standard, 200 mg/dl (1.96 mmol/l). REAGENT 1 is ready to use and stable up to 90 days at 2o to 8o C after opening. Standard is ready to use and stable up to 120 days at 2o to 8o C after opening. REAGENT 1 has a typical yellow color. A yellow/orange color of REAGENT 1 does not influence the reagent performance.

Trypticase soy broth Composition of item from BD Difco: 2.5 g dextrose 2.5 g dipotassium phosphate 3.0 g enzymatic digest of soybean meal 17 g pancreatic digest of casein 5.0 g sodium chloride Reconstitution: 1. Suspend the powder in 1 liter of purified water. 2. Mix thoroughly. 3. Warm gently until solution is complete. 4. Autoclave at 121◦ C for 15 min. 5. Store at room temperature. COMMENTARY Background Information

Characterizing Bile Acid and Lipid Metabolism

BAs are amphipathic molecules synthesized in the liver from cholesterol by a multiple enzymatic process (Russell, 2003). Newly synthesized BAs are referred to as primary BAs, and in humans consist of CA and CDCA. In mice, CDCA is converted to βMCA. Before active secretion into the gallbladder through ABC transporters such as BSEP and MRP2, primary BAs are conjugated with glycine (in humans) and taurine (in mice) (Falany et al. 1994). This process is very efficient, so that the majority of BAs in bile are conjugated. Moreover, this amidation event is important to make BAs less hydrophobic and subsequently less toxic. Besides BAs, other lipids, such as cholesterol and phospholipids, are present in bile. Cholesterol is pumped into bile by ABCG 5/8, while phosphatidylcholine, the most abundant phospholipid, is pumped in by MDR3/Mdr2. The correct ratio of biliary BAs, cholesterol, and phospholipids is of crucial importance for maintaining cholesterol in solution in bile and preventing the precipitation of cholesterol, the first step in chronic gallstone disease (CGD) progression. A high cholesterol to BA

and phospholipid ratio predisposes the patient to cholesterol precipitation. Thus, defects in MDR3 transporter are responsible for CGD. Moreover, genetic or pharmacological alterations in the functionality of MDR3 are also responsible for cholestasis, a disease characterized by interruption of the bile flow and consequent hepatic accumulation of BAs that may lead to liver failure. Clinical parameters that are usually assayed to investigate cholestasis consist of serum BA levels, serum AST, ALT, and ALP activities, and serum bilirubin concentrations. During cholestasis, these parameters can be highly elevated. A well established animal model of cholestasis is the bile duct ligation model, which mimics obstructive extrahepatic cholestasis. After meal ingestion, BAs are released from the gallbladder into the small intestine to promote the absorption of dietary lipids. If the levels of BAs in the intestine are low, as in obstructive cholestasis, absorption of cholesterol and lipophilic vitamins will decrease. When reaching the distal ileum, the majority of BAs (95%) are actively absorbed and returned to the liver via the portal vein to be resecreted into the bile (Love and Dawson, 1998). This

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process of BA recycling is referred to as enterohepatic circulation (Love and Dawson, 1998). Only 0.5 g of BAs per day is lost through feces. This loss is compensated by de novo hepatic synthesis. During enterohepatic circulation, conjugated BAs undergo a deconjugation process in the distal intestine, mediated by bacterial enzymes. Deconjugated BAs are then passively absorbed by enterocytes and returned to the liver, where they will be reconjugated. The primary BAs CDCA and CA are transformed by anaerobic bacteria present in the colon to the secondary BAs lithocholic acid (LCA) and deoxycholic acid (DCA), respectively. Also, secondary BAs are passively reabsorbed from enterocytes to travel back to the liver, where they will be conjugated. Thus, the total BA pool is represented by secondary and, for the most part, primary conjugated BAs. In addition to the liver, where accumulation of BAs can induce cholestasis, BA levels must also be tightly regulated in the intestine. Reduced intestinal BA concentration, due to bile flow obstruction, results in bacterial overgrowth and translocation across the mucosa barrier, with possible systemic infection. On the other hand, high levels of BAs in the intestine, which may result from defects in BA reabsorption, lead to in chronic diarrhea, inflammatory bowel disease, and promotion of intestinal tumorigenesis. In the latter case, animal studies have linked high levels of BAs, especially secondary BAs, to colorectal cancer (CRC). Indeed, patients with CRC exhibit higher levels of secondary BAs in their feces compared to normal subjects. In recent years, a fundamental role for BAs as signaling molecules has emerged. BAs can repress their own synthesis, induce their conjugation and secretion into the gallbladder, and regulate their intestinal absorption and hepatic uptake. Moreover, BAs can also impact lipid metabolism by reducing serum and hepatic TG levels. Finally, by repressing CYP7A1, BAs promote hepatic cholesterol accumulation. In conclusion, BAs represent an important link between liver and intestine by promoting bile flow and intestinal cholesterol absorption, and controlling intestinal bacteria overgrowth and translocation across the mucosa barrier. However, beyond these important roles, BAs, as detergent-like molecules, can be toxic when they accumulate to high levels. In particular, in the liver, high levels of BAs can induce cholestasis, while in the intestine they can promote CRC.

Critical Parameters Bile acid measurement Since the majority of BAs are present in the gallbladder and in the small intestine, the hepatic contribution to the total BA pool size and composition is negligible. Thus, it is not necessary to sample all of the liver, but it is important to avoid any loss of bile from the gallbladder and the small intestine. Several assays have been used to determine both total and individual BAs in biological fluids. The methods that have been used to analyze BA concentrations include gas-liquid chromatography (GLC), high-performance liquid chromatography (HPLC), enzymatic assay, and enzyme cycling assay. GLC and HPLC are not commonly used in clinical laboratories, where automated clinical chemistry analyzers are used for most chemical testing including total BA analysis. The enzymatic assay is now mainly used in small laboratories where manual operations are possible, as the reagents of the total BA test are in lyophilized powder form, and manual reconstitution steps are needed before use. At present, the most widely used total BA test in clinical laboratories is the enzymatic cycling method, which is a liquid-stable and ready-to-use assay (Zhang et al., 2005). The enzymatic total BA assay uses the 3α-hydroxysteroid dehydrogenase (3α-HSD) enzyme to catalyze the oxidation reaction converting 3-α-hydroxyl groups of all BAs to 3keto groups, with concomitant formation of coenzyme NADH from NAD+. The NADH formed is further reacted with NTB to form a formazan dye in the presence of diaphorase enzyme. The dye formation is monitored by measuring the absorbance at 540 nm, which is directly proportional to the BA concentration in the biological sample. The enzyme cycling assay allows for signal amplification through cycled regeneration reactions. In the cycling-based total BA assay, BAs are repeatedly oxidized and reduced by the 3α-HSD enzyme, with a concomitant accumulation of reduced coenzyme thioNADH that is detected at a specific wavelength (405 nm). In the forward reaction, the enzyme catalyzes the oxidation reaction in the presence of coenzyme thio-NAD+ to form oxidized BAs and reduced coenzyme thioNADH. On the other hand, in the reverse reaction, the enzyme catalyzes the reduction reaction in the presence of excess coenzyme NADH to convert oxidized bile back to BAs, which are then ready for the next round of forward oxidation reaction. The innovative use

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of this paired coenzyme and coenzyme analog enables significant signal amplification, and therefore leads to a much higher detection sensitivity of the assay. The rate of thio-NADH formation is detected at 405 nm, and is proportional to the amount of total BA in the sample. The enzyme cycling amplification total BA assay offers analytical performance far beyond the capabilities of conventional BA test methods. The major advantages of the enzyme cycling assay over the conventional enzymatic assays consist in liquid-stable, ready to use reagents, high detection sensitivity, less interference from lipidemic and hemolytic samples, less sample volume needed, and no contamination by formazan dye.

tions with no light exposure, since bilirubin degrades to biliverdin in the presence of light. Most chemical methods for the determination of total bilirubin are based on the reaction between diazotized sulfanilic acid and bilirubin to produce azobilirubin, which absorbs maximally at 560 nm. Such tests are often run in the presence or absence of an organic solvent (e.g., methanol) to distinguish free bilirubin from conjugated bilirubin on a differential solubility basis. Notably, serum substances such as hemoglobin and ascorbic acid interfere with this test. Conversely, in the vanadate oxidation method, hemoglobin up to 500 mg/dl and ascorbate up to 50 mg/dl have been shown not to interfere.

Serum AST, ALP, and ALT activity Reports have indicated that ALT and AST in serum remain stable at 4◦ to 8◦ C or 4◦ C, respectively, for a minimum of 7 days. With regard to ALP, samples should be kept cold and assayed as soon as possible after collection. A timed routine for sample collection and analysis should be established in each laboratory, because ALP levels in serum or plasma, or in reconstituted control serum, rise significantly when stored at 2◦ to 8◦ C or at room temperature. When assaying ALT and AST, hemolyzed specimens should not be used, as erythrocytes contain fifteen and ten times the ALT and AST activity, respectively, that serum contains. For optimal ALP measurement, unhemolyzed serum is preferred, but heparinized plasma may also be used. Oxalate, fluoride, and EDTA inhibit ALP, so they are unsuitable as anticoagulants. Pyridoxal phosphate, which can be found in water contaminated with microbial growth, can elevate ALT and AST values by activating the apoenzyme form of the transaminase. High levels of serum pyruvate may also interfere with assay performance. EDTA, citrate, fluoride, and oxalate inhibit ALP activity. Young at al. (1975) give a list of drugs and other substances that interfere with the determination of AST, ALT, and ALP. The final color developed in the AST and ALT reaction must be read within 60 min. Moreover, if the sample is lipidemic, a serum blank must be run. Bilirubin concentrations of 5 mg/dl and upward can cause falsely elevated values for the AST assay. A serum blank can eliminate this false reading.

Triglyceride measurement Triglycerides in serum are stable for 3 days when stored at 2◦ to 8◦ C. Prolonged storage of the samples at room temperature is not recommended, since other glycerol-containing compounds may hydrolyze, releasing free glycerol. Blood collection devices lubricated with glycerin (glycerol) should be not used. Glycerol in rubber stoppers or in contaminated glassware will elevate triglyceride levels. Lipidemic or hyperbilirubinemic samples will cause falsely elevated results. Samples with gross hemolysis or high bilirubin values will also produce falsely elevated triglyceride values. The kit we proposed for the measurement of triglyceride levels is quite robust, with reduced interference from bilirubin (<40 mg/dl), hemoglobin (<25 mg/dl), and ascorbic acid (<6 mg/dl). However, fresh, clear, nonhemolyzed serum samples from fasting animals are recommended.

Total bilirubin measurement Freshly prepared serum and bile should be used in this procedure. When stored, the serum or bile must be frozen (−20◦ C) under condi-

Cholesterol measurement Cholesterol in serum is stable for 7 days at room temperature and 6 months when frozen and protected against evaporation. Anticoagulant such as fluoride and oxalate will result in false low values. The cholesterol assay here proposed is not influenced by hemoglobin up to 500 mg/dl, bilirubin levels up to 15 mg/dl, and triglycerides up to 1000 mg/dl. However, the use of fresh, clear, nonhemolyzed serum samples is recommended. Phospholipid measurement The phospholipid assay described here is not influenced by hemoglobin up to 500 mg/dl, bilirubin up to 25 mg/dl, and triglycerides up to 1000 mg/dl. However, the use of fresh, clear, nonhemolyzed serum samples is recommended.

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Intestinal bacterial counts For the ileum, make a dilution of 1 to 103 , for the mesenteric lymph node complex, 1 to 102 , and for the cecum, 1:105 .

Troubleshooting When measuring BA, bilirubin, and lipid concentrations, or transaminases in both serum and liver, under certain conditions the values measured may be out of range. In this case, the assay must be repeated after diluting the samples with the respective blank and never with just water (unless the blank is water). When plating bacteria, use more than one dilution to avoid using too high or low a dilution resulting in either an absence of colonies or an excess too numerous to count. Perform at least three dilutions in the range indicated in the protocol. Use plastic pipets and Falcon tubes only for a short time when working with hexane, to avoid extracting chemicals from the tubes. Use glass tubes for organic phases.

Anticipated Results Bile acid measurement In mice, the majority of BAs that recirculate in the enterohepatic system are conjugated with taurine, while glycine-conjugated BAs are almost absent. Indeed, TCA, TβMCA, and TωMCA are the major BAs of the total BA pool. After entering the large intestine, primary BAs are metabolized by the anaerobic bacterial flora. The first major reaction consists of BA deconjugation to release free BAs. Then, deconjugated BAs are dehydroxylated to form more hydrophobic secondary BAs. Notably, the intestines of humans and mice are exposed to different spectra of BAs. Thus, while in mouse feces the predominant BAs are DCA, βMCA, and ωMCA, in human feces DCA, LCA, and CA are the predominant BAs. Under normal conditions, BA levels are extremely low or absent in urine, being reabsorbed at the kidney level. Conversely, in the presence of liver damage (e.g., cholestasis), high levels of BAs are found in both serum and urine. Moreover, most of these BAs are also glucuronidated. In the bile, where BAs are present at a higher concentration, BA levels are reported as mM, while in serum they are reported as μM. Total BA pool is indicated as μmol/100 g body weight and fecal BA content as μmol/day per 100 g body weight. Blood and tissue lipid content Serum lipid concentrations are indicated as mM or mg/dl, while tissue lipid levels are reported as mg lipid/mg tissue. Blood lipid Current Protocols in Mouse Biology

levels are influenced by environmental factors such as the type of diet and the length of fasting. Moreover, genetic factors also impact serum lipid concentration. Thus, for instance, C3H mice exhibit higher plasma cholesterol levels than C57BL/6 mice (208 mg/dl versus 128 mg/dl, respectively). Serum transaminases and bilirubin measurement In addition to serum BA levels, serum AST, ALT, and ALP activity and bilirubin concentration are also dramatically increased in the case of liver damage. Transaminase activity is reported as IU/liter while serum bilirubin levels are indicated as mg/dl. Intestinal bacterial counts After performing bile duct ligation (BDL), bacterial count is expected to dramatically increase. Bacterial count is expressed as 104 cfu/g for the ileum, 105 cfu/g for the mesenteric lymph node complex, and 109 cfu/g for the cecum.

Time Considerations The total time required to perform the enzymatic measurement of BAs, cholesterol, triglycerides, phospholipids, bilirubin, and transaminases (ALP, AST, ALT) is the sum of the hands-on time of the investigators (which depends upon the number of samples to be processed and the experience levels of the workers) and the incubation times of the relative reactions. While the first condition is variable, the second one is constant. In particular, for the measurement of BAs, around 8 min of incubation at 37◦ C is required; for cholesterol and triglycerides, 5 min at room temperature, for phospholipids 10 min at room temperature, for bilirubin 10 min at 37◦ C, for ALT around 50 min at 37◦ C, for AST 24 min at 37◦ C, and for ALP 13 min at 37◦ C are required. For the extraction of BAs from the liver, it is better to conduct the experiment over 2 days, especially if the number of livers to be harvested is high. The first day, livers can be harvested and 50 to 100 mg can be obtained from the whole liver, snap-frozen in liquid nitrogen, and stored at −80◦ C until BAs will be extracted. Then, the time required to extract hepatic BAs is 2 hr and 20 min (incubations and centrifugations) plus the hands-on time of the investigator (which depends on the number of samples to be processed and the worker’s level of experience). For the extraction of hepatic lipids (cholesterol, triglycerides, and phospholipids), it is better to proceed over 2 days, especially if the number of livers to be harvested is high. In this

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case, 100 to 200 mg of liver can be snap-frozen in liquid nitrogen and stored at −80◦ C until the extraction of lipids. 30 sec of homogenization are required for each sample of liver, followed by 20 min of centrifugation. After the second centrifugation, one can stop here and store the samples at –20◦ C. If continuing, at least 1 hr is required to evaporate the organic phase under nitrogen gas; then, 5 more min of incubation at 37◦ C is required for the assay. To measure serum BAs and lipids and bilirubin, at least 100 μl of blood is required. It is recommended to draw blood directly from the heart of the mouse. The time required depends on the number of mice to be sacrificed and the investigator’s experience with the surgical procedure. After being harvested, blood can be stored on ice until the end of the surgery. Then, after collection, samples have to be centrifuged for 5 min at 10,000 × g, 4◦ C, to separate serum, which can be stored at –80◦ C. The times required to perform enzymatic analysis are reported above. To measure biliary BAs and lipids, mice should be fasted for at least 4 hr in order to have a filled gallbladder. The time required to harvest the gallbladder depends on the number of samples and the experience of the investigator. Gallbladders can be stored in ice until the end of the surgery. Then, break up the gallbladder with a needle and centrifuge 1 min at 10,000 × g to separate gallbladder from bile. The time required to assay the content of BAs and lipids in bile is reported above. For fecal BA extraction, feces are collected over 3 days and then air-dried for 2 days. Then, 2 additional days are required to complete the extraction process and enzymatic assay. For the determination of the total BA pool size, the time depends on the experience of the investigator in harvesting tissues, and on the number of samples. 30 min are then required to bring ethanol from 100 ml to 30 ml and at least 1 hr to filter approximately 10 samples. For the time required to perform the enzymatic assay, see above. To measure bile flow, 10 to 15 min are required to collect bile. Thus, the total time to perform this assay is mainly affected by the number of mice to cannulate and by the experience of the investigator in cannulating the gallbladder. To perform the intestinal BA absorption assay, 5 to 6 hr and 45 min are required as basic time. The final time required depends on the number of mice and the experience of the investigator in performing surgery, as well as other hands-on time at the bench.

For measuring intestinal cholesterol absorption, feces are collected over 2 days and then air-dried for 2 more days. Then, 5 hr of incubations are required. The final time depends on the number of mice, and the experience of the investigator in performing gavage, as well as other hands-on time at the bench. The in vivo fatty acids synthesis is performed in 2 days. For the first day, a basic time of 2.5 hr of incubations is required. The total time depends on the number of mice, the experience of the investigator in performing injections and surgery, plus hands-on time at the bench. On the next day, after hexane evaporation overnight, 1 hr is sufficient to finish the assay. To perform an intestinal bacteria count, a basic time of 20 hr of incubation at 37◦ C is required. The total time for this assay is then affected by the number of mice, the experience of the investigator in performing surgery, and the hands-on time needed to plate bacteria. For this assay, two people should be involved—one to perform surgery and one to plate bacteria.

Literature Cited Allain, C.C., Poon, L.S., Chan, C.S., Richmond, W., and Fu, P.C. 1974. Enzymatic determination of total cholesterol. Clin. Chem. 20:470-475. Donovan, J. and Brown, P. 2006. Euthanasia. Curr. Protoc. Immunol. 73:1.8.1-1.8.4. Doumas, B. and Biggs, H.G. 1969. A colorimetric method for assaying serum aspartate aminotransferase activities. Clin. Chim. Acta 23:7582. Falany, C.N., Johnson, M.R., Barnes, S., and Diasio, R.B. 1994. Glycine and taurine conjugation of bile acids by a single enzyme: Molecular cloning and expression of human liver bile acid CoA:amino acid N-acyltransferase. J. Biol. Chem. 269:19375-19379. Folch, J., Lees, M., and Sloane Stanley, G.H. 1957. A simple method for the isolation and purification of total lipids from animal tissues. J. Biol. Chem. 226:497-509. Gambino, S.R., Other, A., and Burns, W. 1967. Direct serum bilirubin and the sulfobromophthalein test in occult liver disease. JAMA 201:1047-1049. Inagaki, T., Moschetta, A., Lee, Y.K., Peng, L., Zhao, G., Downes, M., Yu, R.T., Shelton, J.M., Richardson, J.A., Repa, J.J., Mangelsdorf, D.J., and Kliewer, S.A. 2006. Regulation of antibacterial defense in the small intestine by the nuclear bile acid receptor. Proc. Natl. Acad. Sci. U.S.A. 103:3920-3925. Jung, D., Inagaki, T., Gerard, R.D., Dawson, P.A., Kliewer, S.A., Mangelsdorf, D.J., and Moschetta, A. 2007. FXR agonists and FGF15 reduce fecal bile acid excretion in a mouse

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model of bile acid malabsorption. J. Lipid. Res. 48:2693-2700. Love, M.W. and Dawson, P.A. 1998. New insights into bile acid transport. Curr. Opin. Lipidol. 9:225-229. McGowan, M.W., Artiss, J.D., Strandbergh, D.R., and Zak, B. 1983. A peroxidase-coupled method for the colorimetric determination of serum triglycerides. Clin. Chem. 29:538-542. Moschetta, A., Bookout, A.L., and Mangelsdorf, D.J. 2004. Prevention of cholesterol gallstone disease by FXR agonists in a mouse model. Nat. Med. 10:1352-1358. Nagengast, F.M., Grubben, M.J. and van Munster, I.P. 1995. Role of bile acids in colorectal carcinogenesis. Eur. J. Cancer 31A:1067-1070. Oelkers, P., Kirby, L.C., Heubi, J.E., and Dawson, P.A. 1997. Primary bile acid malabsorption caused by mutations in the ileal sodium-dependent bile acid transporter gene (SLC10A2). J. Clin. Invest. 99:18801887. Reitman, S. and Frankel, S. 1957. A colorimetric method for the determination of serum glutamic oxalacetic and glutamic pyruvic transaminases. Am. J. Clin. Pathol. 28:56-63. Roy, A.V. 1970. Rapid method for determining alkaline phosphatase activity in serum with thymolphthalein monophosphate. Clin. Chem. 16:431-436.

Russell, D.W. 2003. The enzymes, regulation, and genetics of bile acid synthesis. Annu. Rev. Biochem. 72:137-174. Searcy, R.L. 1969. Diagnostic Biochemistry. McGraw-Hill, New York. Speers, A. M., Cologgi, D. L. and Reguera, G. 2009. Anaerobic cell culture. Curr. Protoc. Microbiol. 12:A.4F.1-A.4F.16. Teitz, N.W. 1976. Fundamentals of Clinical Chemistry. W.B. Saunders, Philadelphia, Pa. Turley, S.D, Herndon, M.W., and Dietschy, J.M. 1994. Reevaluation and application of the dualisotope plasma ratio method for the measurement of intestinal cholesterol absorption in the hamster. J. Lipid Res. 35:328-339. Watanabe, M., Houten, S.M., Wang, L., Moschetta, A., Mangelsdorf, D.J., Heyman, R.A., Moore, D.D., and Auwerx, J. 2004. Bile acids lower triglyceride levels via a pathway involving FXR, SHP, and SREBP-1c. J. Clin. Invest. 113:14081418. Young, D.S., Pestaner, L.C., and Gibberman, V. 1975. Effects of drugs on clinical laboratory tests. Clin Chem. 21:1D-432D. Zhang, G.H., Cong, A.R., Xu, G.B., Li, C.B., Yang, R.F., and Xia, T.A. 2005. An enzymatic cycling method for the determination of serum total bile acids with recombinant 3alpha-hydroxysteroid dehydrogenase. Biochem. Biophys. Res. Commun. 326:87-92.

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Hematology Testing in Mice Pontus Lundberg1 and Radek Skoda1 1

Department of Biomedicine, Experimental Hematology, University Hospital Basel, Basel, Switzerland

ABSTRACT The mouse is an increasingly important system for the study of both normal and aberrant hematopoiesis. As a model organism, the mouse recapitulates much of human hematopoiesis; however, there are some important differences. Here, the basic approaches for analyzing hematopoiesis in mice are described. In particular, methods are provided for the collection and analysis of peripheral blood, flow cytometry analysis of peripheral blood, bone marrow, and spleen cells, and isolation and transplantation of bone marrow C 2011 by John Wiley & Sons, Inc. stem cells. Curr. Protoc. Mouse Biol. 1:323-346  Keywords: mouse hematology r peripheral blood r bone marrow transplantation r hematopoiesis r hematopoietic r stem cell

INTRODUCTION The first genetically modified mice were generated by Jaenisch and Mintz (1974) by introducing foreign DNA into the mouse genome. In the early 1980s, the technology of generating transgenic mice received wider appreciation among scientists, and in the 1990s the techniques for homologous recombination in embryonic stem (ES) were successfully applied to generate knockout and knockin mice. The introduction of the loxP-Cre recombinase system allowed studying conditional and tissue-specific effects of gene knockouts. A few years after the first report on transgene integration in mice, another methodological milestone was achieved with the successful engraftment of human cells into SCID mice (McCune et al., 1988; Mosier et al., 1988), which was later refined by the introduction of NOD/SCID (Greiner et al., 1995) and NOD/SCID/gamma(c)(null) mice (Ito et al., 2002). These immunodeficient mice allowed the study of human disease by xenotransplantation of human hematopoietic cells, e.g., leukemic cells (reviewed by Shultz et al., 2007). The aforementioned technologies are now routinely used, and these mouse models have had a profound impact on our understanding of normal and aberrant hematopoiesis. To take full advantage of these mouse models, a systematic approach to studying their blood systems must be taken. Described below are methods and approaches for assessing hematopoiesis in mice. In vivo analysis and manipulations When assaying the hematological status of a mouse, analysis of the peripheral blood is often the first parameter analyzed. In contrast to bone marrow analyses, peripheral blood can be collected via tail vein bleeding (see Basic Protocol 1), thus allowing analyses over an extended period of time. Peripheral blood can be analyzed in several ways to cover various aspects of hematopoiesis. A complete blood count (CBC) is, as in human patients, performed routinely, and will provide an exact picture of the relative and absolute abundance of all cell types present in the peripheral blood (see Basic Protocol 1). The CBC is the single most informative analysis that can be performed when analyzing the hematological status of a mouse. As a complement to the CBC, a peripheral blood smear is frequently performed in parallel (see Basic Protocol 2). Similar to a CBC, the blood smear will provide information on the relative abundance of the cells present in the Hematology Testing in Mice Current Protocols in Mouse Biology 1: 323-346, September 2011 Published online September 2011 in Wiley Online Library (wileyonlinelibrary.com). DOI: 10.1002/9780470942390.mo110088 C 2011 John Wiley & Sons, Inc. Copyright 

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peripheral blood. A blood smear will also provide information concerning cell-specific alterations such as neutrophil segmentation or morphological alterations, e.g., erythroid spherocytosis. The blood smear can also indicate whether an apparent thrombocytopenia is caused by EDTA-induced aggregation, thus being a pseudo thrombocytopenia. When a more in-depth analysis of the peripheral blood is needed to assess, e.g., expression levels of certain cell surface markers or genes in a subset of cells, flow cytometry and fluorescence activated cell sorting (FACS) of peripheral blood cells are the preferred technologies (see Basic Protocol 3). Flow cytometry can also be applied in order to assay for alteration in the half-life of peripheral blood cells (see Support Protocol 5). Terminal workup and analysis The terminal workup will provide information that is not possible to ascertain while the animal is still living, such as organ status. A typical terminal workup will start with a cardiac puncture (see Basic Protocol 4), which will yield large quantities of aseptic blood. From a hematological perspective, the most important information gleaned when performing a terminal workup is the status of the blood-producing tissues, namely the bone marrow and spleen. Extracting cells from bone marrow and spleen is described in Basic Protocols 5 and 6. Bone marrow cells can be transplanted into sublethally/lethally irradiated hosts (see Basic Protocol 7) using tail vein injection (see Basic Protocol 8), and the transplanted animal then analyzed for bone marrow cellular composition. In some cases, the effect of a transgene or knockout can lead to embryonic lethality; thus, no pups with the desired genotype will be born. In these cases, an alternative is to perform adoptive transfer by transplanting fetal liver cells into irradiated hosts (described in Support Protocol 6). The stem cell compartment in mice has been well characterized, with studies beginning in the mid 1980s (Muller-Sieburg et al., 1986; Spangrude et al., 1988), and there are now many markers for dissecting the different stages of hematopoiesis (stem cell analysis is described in Support Protocol 1 and erythroid maturation is described in Support Protocol 2) from the earliest stem/progenitor cells until the exit to the periphery. Another assessment of the bone marrow stem/progenitor lineage commitment is to perform colony assays (see Support Protocol 3), which can be further extended into serial replating experiments to assess self-renewal capacity (Support Protocol 4). Plating and replating cells from the bone marrow can provide information on, e.g., increased self-renewal or a skewing of the differentiation of progenitor cells (where more cells of a specific lineage are produced). It also provides a way to assay cytokine dependence or efficacy of pharmacological inhibitors ex vivo.

STRATEGIC PLANNING

Hematology Testing in Mice

The following steps should be considered when approaching a new mouse strain with a suspected phenotype in the hematopoietic system. Depending on the mouse mutant and experimental setup, additional customized tests may be needed. Initially, observe the mouse: shortness of breath and pale mucosa (e.g., nostrils) are signs of anemia. Loss of activity and ruffled fur can indicate an underlying disease, such as leukemia or infection. After the initial observation of the mouse, collect peripheral blood and perform a blood smear, a complete blood count (CBC), and flow cytometric analysis. These analyses will provide information on the basic hematological state of the mouse. Continue to draw blood and perform serial blood counts every 4 to 6 weeks until the termination of the experiment. The timing of the terminal workup will be a function of the phenotype of the mice. If there is a strong phenotype with a decrease in health status, the terminal workup should be performed as soon as a set of predetermined health conditions are observed. If the phenotype does not cause a rapid and severe decline in health, monitoring of the mice over an extended period of time (>1 year) may be necessary to obtain conclusive results. When terminating the experiment, the basic procedures usually

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include a cardiac puncture to obtain a sufficient volume of blood for performing a blood smear, CBC, flow cytometric analysis, and extraction of DNA, RNA, and/or protein from peripheral blood cells. Bone marrow and spleen cells are collected for flow cytometry, cell sorting, and freezing of live cells for later use. Cultures of bone marrow cells can be initiated at this stage. Histopathology of all major organs, including the bones, should be performed. Depending on the context, transplantation of bone marrow cells or purified stem cell populations into lethally or sublethally irradiated hosts can be added to the analysis. Important for all experiments, particularly the terminal workup, is careful planning. Many of the protocols listed should be performed on fresh cells; thus, the analyses will be carried out on the same day that the mice are euthanized. Often, certain experiments must be prioritized, since performing all the procedures listed here for terminal workup can be too much for one day when working with large groups of mice. NOTE: All protocols using live animals must first be reviewed and approved by an Institutional Animal Care and Use Committee (IACUC) or must conform to governmental regulations regarding the care and use of laboratory animals. For ethical considerations, see Baertschi and Gyger (2011).

BLOOD COLLECTION IN THE MOUSE VIA TAIL VEIN BLEEDING There are various ways to take blood from a mouse. The most commonly used methods are retro-orbital bleeding and tail vein bleeding. Mouse tail vein bleeding is considered easier, since anesthetics are required for retro-orbital bleeding but not for tail vein bleeding. Also, a higher level of expertise is required for the retro-orbital bleeding, since injuries can result in blindness of the animals. Table 1 gives guidelines on the amounts of blood and percentages of total circulating blood that can safely be drawn from variously sized mice, as well as the recovery periods needed subsequent to these blood draws. Tail vein bleeding provides large amounts of blood and is relatively nontraumatic for the animals. The risk of contamination is low; however, when “milking” the tail to achieve a larger volume of blood (i.e., pressing with the finger along the vein to push out blood from the vein), the quality of the blood might decrease and the measured blood parameters might be misleading due to increased chance of tissue contamination.

BASIC PROTOCOL 1

It is typical to draw ∼200 μl of blood from a mouse every 4 to 6 weeks for monitoring changes in blood parameters. This amount is sufficient for a CBC as well as flow cytometry analysis. For the CBC analysis, the blood is diluted in 0.9% NaCl at a 1:2 ratio (100 μl blood plus 200 μl 0.9% NaCl) or 1:3 ratio (70 μl blood plus 210 μl 0.9% NaCl). Thus, about 70 μl of blood is sufficient for performing a CBC. Described below is arterial bleeding of a mouse; however, all bleedings of mouse tails are commonly referred to as “tail vein bleeding.” Table 1 Recommended Amounts for Bleeding and Estimated Recovery Timesa

Body weight (g)

Circulating blood volume (ml)

7.5% (1-week recovery time)

15% (4-weeks recovery time)

20

1.10–1.40

∼95 μl

∼190 μl

25

1.37–1.75

∼120 μl

∼240 μl

a From http://oacu.od.nih.gov/ARAC/Bleeding.pdf.

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Figure 1 Tail vein bleeding. (A) Position the scalpel so that it is touching the artery. (B) Directly after making the cut, blood should start dripping. (C) Collect the blood in a tube already containing EDTA until the desired amount is obtained. Do not collect more blood than recommended, as excessive bleeding could harm the mice.

Table 2 List and Explanation of the Most Commonly Used Parameters in a Complete Blood Count (CBC)

Abbreviation

Full name

RBC

Red blood cell count

HGB

Hemoglobin

Comment

Hct

Hematocrit

(RBC × MCV) ÷ 10

MCV

Mean corpuscular volume

Mean of RBC volume histogram

MCH

Mean corpuscular hemoglobin

(HGB ÷ RBC) × 10

MCHC

Mean corpuscular hemoglobin concentration

(HGB ÷ [RBC × MCV]) × 1000

RDW

Red cell volume distribution width

Distribution of RBC volume

WBC

White blood cell count

Plt

Platelet count

MPV

Mean platelet volume

Mean of platelet volume histogram

LUC

Large unstained cells

Can contain leukemic blast cells

Materials Mouse subject 0.9% NaCl BD Microtainer blood collection tube (Becton Dickinson): EDTA for CBC analysis or heparin for blood chemistry Heat lamp Mouse restrainer Scalpel 1. Heat the mouse tail by either using a warm compress, putting the tail in warm water, or placing a heat lamp on top of the mouse’s cage. When the tail is heated the veins become more visible and blood flow is increased. During the heating process, carefully monitor the behavior of the mice, as excessive heating is harmful for the animals.

Hematology Testing in Mice

2. Put the mouse in a restrainer to immobilize the body, but maintain a firm grip on the tail (Fig. 1A).

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3. While stretching the tail, use a scalpel to cut the artery on the distal side of the mouse tail (Fig. 1B). 4. Collect the blood directly into a tube that contains the appropriate anticoagulant (Fig. 1C), e.g., EDTA for CBC analysis or heparin for blood chemistry. Immediately invert, but do not shake the tube, to mix the blood with the anticoagulant. 5. After the blood is collected, press on the wound with a tissue to stop the bleeding and return the mouse to the cage A complete blood count (CBC) should be included in the standard workup for a comprehensive characterization of a mouse’s hematological status (Table 2). A number of automated analyzers that are capable of accurately measuring multiple parameters in mouse blood are commercially available. Most automated analyzers will also determine a differential blood count of the white blood cells that can be divided into lymphocytes, monocytes, neutrophil granulocytes, eosinophil neutrophils, and basopohil granulocytes.

BLOOD SMEARS A blood smear requires a very small amount of blood, and some parameters such as morphological abnormalities can only be properly determined using a microscopic assessment. Thus, a blood smear should be part of every standard hematological workup. The microscopic evaluation of a stained (May-Gr¨unwald-Giemsa or Wright-Giemsa) blood sample will provide information on alterations in erythrocytes, platelets, and white blood cell morphology. Performing a blood smear is fast and relatively simple.

BASIC PROTOCOL 2

Figure 2 The process of performing a blood smear. (A) See Basic Protocol 2, step 1. (B) See Basic Protocol 2, step 2. (C) See Basic Protocol 2, step 3. (D) See Basic Protocol 2, step 4.

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Materials Peripheral blood (Basic Protocol 1; in some cases the source might be cardiac puncture) Microscope slides (26 × 76 mm) Wright-Giemsa stain (e.g., Hemacolor, Merck) 1. Put one drop of blood on the top end of a microscope slide (Fig. 2A). 2. With another microscope slide, held in the hand, oriented at a low angle to the slide lying on the bench with the blood drop, make very light contact with the blood drop so that the capillary forces hold the blood drop between the lying slide and the shorter edge of the slide held in the hand. (Fig. 2B). 3. Within the time span of ∼1 sec, drag the held microscope slide in one smooth motion over the lying microscope slide (Fig. 2C), exerting as little force as possible with the first slide upon the second. 4. Let the smear dry, after which a thin film of blood should be present on the microscope slide (Fig. 2D) A successful blood smear ideally exhibits a comet-shaped appearance, tapering in the direction in which the upper slide is pulled.

5. Stain the slide with Hemacolor according to the manufacturer’s instructions. BASIC PROTOCOL 3

FLOW CYTOMETRY ANALYSIS AND FLUORESCENCE ACTIVATED CELL SORTING (FACS) OF PERIPHERAL BLOOD Important additional information on the composition of peripheral blood cells, bone marrow cells, spleen cells, and cells from lymphoid organs can be obtained by flow cytometry. Furthermore, subpopulations of cells can be purified using FACS sorting. Mouse hematopoietic cells are very well characterized, which allows prospective sorting/analysis of subpopulations using various cell-surface antigens. Described below are methods to prepare peripheral blood and bone marrow/spleen cells for sorting/analysis of peripheral blood and bone marrow stem/progenitor cells. The most commonly used markers and their corresponding antibodies are listed in Table 3.

Materials Peripheral blood (Basic Protocol 1) Antibodies (eBioscience, Biolegend, BD Pharmingen) FACS buffer: phosphate-buffered saline (PBS) containing 0.1% (v/v) fetal bovine serum (FBS) and (optionally) 0.1% (w/v) sodium azide if the cells are not to be cultured/transplanted after sorting Red blood cell lysis buffer (e.g., BD FACS lysis buffer; Becton Dickinson) FACS tubes (Becton Dickinson) Centrifuge Fluorescence-activated cell sorter (FACS; Robinson et al., 2011) NOTE: After drawing blood from the mouse, keep the blood in a tube supplemented with EDTA to avoid clotting.

For analysis/sorting of erythrocytes and platelets 1a. Add 1 μl of blood into a FACS tube containing 100 μl FACS buffer. 2a. Add the antibody at the titrated concentration. Hematology Testing in Mice

3a. Incubate for ∼20 min in the dark at room temperature.

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A

B

104

30K 103 20K SS Lin: SS

SS Log: SS

4 2 102 1

10K

101

5 3 100

0 100

101

102

103

104

0

10K

20K

30K

FS Lin: FS

FS Log: FS

Figure 3 Representative scattergrams for peripheral blood. (A) Scattergram of peripheral blood before RBC lysis in log scale with platelets (1), and red blood cells together with lymphocytes (2). (B) Scattergram of peripheral blood after RBC lysis: lymphocytes (3), granulocytes (4), and monocytes (5).

Table 3 Commonly Used Cell Surface Antigens for Analysis/Sorting of Peripheral Blood

Cell type

Antibody target

Other names

Erythrocytes

Ter119

Glycophorin A–associated protein (ly-76)

Platelets

CD61

Integrin beta chain beta 3

Monocytes

Mac-1

CD11b/CD18

Granulocytes

Gr-1

Ly-6G

B cells

B220

CD45R

T cells

CD3

CD3 epsilon

NK cells

NK1.1

NKR-P1C, Ly-55

4a. Wash the cells by adding 4 ml of FACS buffer to the tube so that the contents are well mixed. 5a. Centrifuge 5 min at 300 × g, room temperature. 6a. Remove the supernatant and resuspend the pellet containing the cells in 1 ml FACS buffer. When acquiring FACS data on platelets and RBCs, use the log scale for forward/side scatter, where platelets will be forward/side scatter–low cells and red blood cells forward/side scatter high–cells (Fig. 3A). Be sure that the exclusion filtering of the FFS/SSC is not set to default, which will, in most cases, exclude the platelets because of their small size. Even though it is possible to do RBC lysis when acquiring platelets, the membrane debris of the erythrocytes tends to create a considerable background, so it is generally not recommended to perform RBC lysis when acquiring platelets. Detailed protocols for FACS and flow cytometry are provided in Robinson et al. (2011).

For analysis/sorting of lymphocytes, monocytes, and granulocytes 1b. Add ∼20 μl blood to 80 μl of FACS buffer.

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2b. Add the antibody at the titrated concentration. 3b. Incubate for ∼20 min in the dark at room temperature. 4b. Add RBC lysis buffer and incubate for 10 min at room temperature. 5b. Fill the tube with FACS buffer and centrifuge the cells 5 min at 300 × g, room temperature. 6b. If there is a red pellet, try washing the cells with FACS buffer a second time. If the pellet is still red, then perform another RBC lysis (step 4b) followed by washing (step 5b). 7b. Resuspend the pellet in 200 μl FACS buffer. Perform FACS analysis (Robinson et al., 2011). Gr-1 and Mac-1 are cross-reactive, but these contaminating cells can often be observed as having weaker fluorescence intensity. However, a relatively pure population of granulocytes can be achieved by forward/side scatter which, together with analyzing/sorting the cells with the highest fluorescence, will give a pure population. A representative scattergram of a peripheral blood sample after RBC lysis is depicted in Figure 3B. BASIC PROTOCOL 4

CARDIAC PUNCTURE When performing terminal bleeding of mice, cardiac puncture will yield large amounts of aseptic blood. Blood withdrawal by cardiac puncture is considered a euthanasia procedure and should be performed only after ensuring that the animal is under deep anesthesia. There are two generally recommended ways to do cardiac puncture, as described below (Fig. 4).

Materials Mouse subject EDTA tubes (BD Microtainer, Becton Dickinson) 1-ml syringes and 25-G, 16-mm needles Additional reagents and equipment for euthanizing mice by CO2 asphyxiation (Donovan and Brown, 2006) 1. Euthanize the mouse by CO2 asphyxiation (Donovan and Brown, 2006). Lay the mouse on its back and push the 25-G needle ∼10 mm deep, either though the

Figure 4 Cardiac puncture. (A) Insert the needle in an angled fashion. (B) Retract the plunger very slowly to collect the blood. If the blood stops flowing, rotating the needle or pushing/pulling a bit might restore the blood flow.

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sternum or below the sternum in an angled fashion. (Fig. 4A and B). Alternatively, hold the mouse by the scruff of the neck, keep the head in an upward direction, and insert the needle in an angled fashion below the sternum. It is recommended to use a 25-G, 16-mm needle with a 1- or 2-ml syringe. A longer needle increases the likelihood of inserting the needle too deeply, thus penetrating through the heart. The needle should be inserted ∼10 mm deep, then blood should be slowly withdrawn.

2. Slowly retract the plunger to collect the blood. When retracting the plunger, do not retract it too rapidly as it might cause the heart to collapse. If no blood is visible, creating a vacuum by pulling back the plunger might start the blood flow. If still no blood is visible, slightly pull back the syringe and try a different angle/depth. When blood appears in the syringe, keep the position of the needle fixed and slowly pull back the plunger to collect the blood. If blood stops flowing, rotating the syringe or slightly pulling it back/forward might restore the flow.

3. When a sufficient amount of blood is collected, or the blood flow has stopped, remove the needle from the syringe (pressing the blood through the needle can cause cell lysis and coagulation problems), and expel the blood into a tube containing EDTA. The amount of blood that can be collected with a cardiac puncture varies depending on the size of the mouse; however, for a medium-sized mouse, ∼1 ml of blood can typically be collected. It is important to perform the heart puncture as rapidly as possible, as the blood might start to coagulate before it is dispensed into the tube containing EDTA.

ISOLATION AND TRANSPLANTATION OF BONE MARROW CELLS Bone marrow must be isolated to evaluate the primary hematopoietic site and to obtain material for serial transplantation and colony assays. Sorting of bone marrow cells for analyzing differences in stem cell populations (Support Protocol 1) and erythroid maturation (Support Protocol 2) is important for assessing aberrant hematopoiesis. Also, plating/replating of cells in methylcellulose can give an indication of lineage commitment and colony-forming ability (Support Protocol 3), as well as self-renewal capacity (Support Protocol 4).

BASIC PROTOCOL 5

Materials Mouse subject Phosphate-buffered saline (PBS), ice cold RPMI medium (e.g., Invitrogen) supplemented with 10% fetal bovine serum (FBS) Red blood cell (RBC) lysis buffer (see recipe) Mouse hematopoietic cell depletion kit (R&D Systems) Dissecting equipment including forceps and scissors Mortar and pestle Cell strainers (e.g., Becton Dickinson) 50-ml conical polypropylene tubes (e.g., BD Falcon) Additional reagents and equipment for euthanizing mice by CO2 asphyxiation (Donovan and Brown, 2006) 1. Euthanize the mouse by CO2 asphyxiation (Donovan and Brown, 2006) and lay it on its back on a dissecting board. Fix the front legs (e.g., with needles) and cut with scissors along the tibia and femur of one leg and open the muscles until the bone is visible. 2. After cleaning the bones of muscle tissue, cut with the scissor at the hip joint to remove the tibia and femur from the pelvic bone. 3. Cut off the hind paw and clean the femur and tibia with scissors and laboratory tissues to remove all flesh from the bone. Current Protocols in Mouse Biology

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4. After the bone is clean, place it in ice-cold PBS while repeating the bone removal/cleaning procedure on the other leg. The two hind legs of a mouse contain ∼50–100 × 106 cells after RBC lysis. If a substantially larger number of cells are needed, proceed with removing the pelvic bones and, if needed, also the spine to increase cell numbers.

5. Put the bones in a mortar with 5 ml ice-cold RPMI supplemented with 10% FBS. Crush the bone marrow with a pestle until all bones are crushed (no more sound of bones being crushed will be heard). 6. Filter the medium containing the cells though a cell strainer into a 50-ml conical polypropylene tube. 7. Optional: Add more medium to the crushed bones and repeat steps 5 to 6 a second time to increase the yield of cells. 8. Add 25 ml of RPMI supplemented with 10% FBS to the tube and centrifuge 8 min at 300 × g, room temperature. Remove the supernatant. The authors usually supplement the medium with 1× penicillin/streptomycin (added from 100× stock, e.g., Invitrogen), but this is not strictly necessary, as the process described here is quick and aseptic.

9. Optional: Lyse RBCs by adding 20 ml 1× RBC lysis buffer to the cells and incubating 10 min at room temperature. After RBC lysis, wash the cells twice with RPMI supplemented with 10% FBS as described in step 8 to remove debris from the lysed cells. 10. Optional: Perform T cell depletion following the protocol of the manufacturer of the mouse hematopoietic cell depletion kit used. For transplantation experiments (Basic Protocol 8), RBC lysis followed by washing and counting is recommended. The bone marrow can be kept for several hours at 4◦ C without harm to the cells. However, it is important to keep the cells in medium supplemented with FBS to maintain high viability. For transplantation of bone marrow cells into lethally irradiated recipient mice, at least 2 × 105 cells (after RBC lysis) are recommended to be certain of reconstitution. However, if there is no shortage of donor cells, quantities of ∼1–2×106 bone marrow cells can be used to assure reconstitution. Often, competitive transplantations are used to assess whether a population of cells with, for example, a mutation, has an altered proliferation capacity. These kinds of studies follow the same protocol as described above; however, in the last step, cells are mixed in a ratio (wild type:mutant). To allow for effective tracking of the two populations, different allelic combination of CD45 are often used (Ly-5.1 and Ly-5.2) whose ratio can be quantified to determine the contribution to the peripheral blood cells as well as other populations. The disadvantage of using CD45 is that it is not expressed on erythrocytes and platelets; thus, if there is a specific interest in tracking these populations, alternatives such as mice expressing GFP under the ubiquitin-C promoter can be used (The Jackson Laboratory). SUPPORT PROTOCOL 1

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FLOW CYTOMETRY ANALYSIS/SORTING OF BONE MARROW STEM/PROGENITOR CELLS Bone marrow contains stem cells, early progenitor cells, late progenitor cells, and mature cells. In the later stages leading to terminal differentiation of hematopoietic cells, the cells exit the bone marrow and enter the bloodstream. In many cases, lineage depletion (removal of cells displaying cell surface antigens for a particular lineage commitment while keeping the more immature cells) is performed to reduce sorting/analysis time.

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Table 4 Commonly Used Antibody Combinations for Stem/Progenitor Cell Analysis

Cell type

Antibody combination

LSK

c-kit+ , sca1+

LT-HSC

c-kit+ , sca-1+ , (CD150+ ), CD34– CD48– , Flk-2–

ST-HSC

c-kit+ , sca-1+ , CD150+ , CD34+

MPP

c-kit+ , sca-1+ , CD150– , CD34+

CMP

c-kit+ , sca-1– ,CD34+ , FcγRlo, IL-7R–

MEP

c-kit+ , sca-1– ,CD34– , FcγRlo, IL-7R–

CLP

c-kitlo , sca-1lo , IL-7Rα + , Flk-2+

GMP

c-kit+ , sca-1– ,CD34+ , FcγRhi , IL-7R–

This is not always necessary for flow cytometry analysis, but when performing FACS it massively decreases the sorting time. Many kits use biotinylated antibodies for lineage depletion. These kits normally remove >90% of the lineage-positive cells but, to ensure that the lineage depletion is successful, addition of a fluorescent streptavidin conjugate in the staining cocktail will provide an indication of the depletion efficacy and allow gating of only lineage-negative cells. As a control to set the gate for the lineage-negative population, cells that are not lineage depleted but stained with the lineage-depletion cocktail should be used (as described in the manufacturer’s protocol). Lineage depletion is not necessary for sorting/analysis of stem/progenitor cells, but it significantly reduces the sorting/analysis time, especially when running multiple samples. Markers commonly used for the analysis of stem/progenitor cells are listed in Table 4.

Materials Bone marrow cells (Basic Protocol 5) Mouse hematopoietic cell depletion kit (R&D Systems) FACS buffer: phosphate-buffered saline (PBS) containing 2% (v/v) fetal bovine serum (FBS) and (optionally) 0.1% (w/v) sodium azide if the cells are not to be cultured/transplanted after sorting Antibody staining cocktail Centrifuge Flow cytometer/cell sorter supporting multi-color analysis (see Robinson et al., 2011) and FACS tubes 1. Take the bone marrow (either before or after RBC lysis) and perform a lineagedepletion procedure following the manufacturer’s protocol for the mouse hematopoietic cell depletion kit used. In this process, bone marrow cells are incubated with antibodies recognizing antigens displayed on differentiated cells. In the R&D mouse hematopoietic cell depletion kit, these antibodies are CD5, CD11b, CD45R (B220), Ly-6G (Gr-1), and TER-119. These linage-positive cells will be separated using magnetic beads, after which a population of immature cells will remain. The amount of cells needed depends on the experiment. The protocol for the R&D Systems mouse hematopoietic cell depletion kit lists 10 × 107 cells, but this is scalable depending on the application.

2. After lineage depletion, wash the cells by centrifuging 8 min at 300 × g, 4◦ C, removing the supernatant, and resuspending the cells in the staining cocktail. Alternatively, if several different stainings are to be made, resuspend the cells in a small volume,

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e.g., 10 μl of FACS buffer per staining, transfer them to individual tubes, and add the staining cocktail. The authors routinely use streptavidin–Pacific Blue in combination with DAPI for the combined removal of dead cells (Violethi cells) and cells with a lineage commitment that were not successfully removed during the lineage depletion assay (Violetmed cells) while analyzing viable primitive cells (Violetlo cells). This gating strategy requires a violet laser (405 nm) or equivalent. It should be noted that even though the mouse markers separate different subpopulations of stem/progenitor origin, there is still quite some heterogeneity even in the populations sorted using several cell-surface antigens.

3. Proceed to flow cytometric analysis/cell sorting. SUPPORT PROTOCOL 2

FLOW CYTOMETRY ANALYSIS OF ERYTHROID MATURATION Bone marrow is the main site of hematopoiesis in mice, as is also true for humans. However, under normal conditions, mice also have an active site of hematopoiesis in the spleen, whereas in humans this is typically restricted to certain disease conditions. A straightforward strategy to assess whether erythroid differentiation in the bone marrow is altered involves tracking the maturation of early erythroid progenitors to the stage when they are mature and ready to exit the bone marrow. To follow the maturation process, a dual-antibody stain with CD71 and ter119 is often used. Do not perform RBC lysis of the bone marrow/spleen, as this will destroy the more differentiated red blood cell progenitors, thus only allowing the analysis of very early CD71hi erythroid progenitors. Based on the staining intensity, erythroid maturation can be analyzed where R1 corresponds to proerythroblasts (CD71hi Ter119lo-med ), R2 corresponds to basophilic erythroblasts (CD71hi Ter119hi ), R3 corresponds to polychromatic erythroblasts (CD71med Ter119hi ), and R4 corresponds to orthochromatic erythroblasts and reticulocytes (CD71lo Ter119hi ) (Fig. 5).

R1

104

0.91

: APC-CD71

103

8.7

R2

1.42

R3

9.36

R4

102

101

100 100

101

102

103

104

: PE-ter119

Figure 5

Analysis of erythroid maturation in bone marrow.

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Materials Bone marrow cells (Basic Protocol 5) or spleen cells (Basic Protocol 6) FACS buffer: phosphate-buffered saline (PBS) containing 0.1% (v/v) fetal bovine serum (FBS) and (optionally) 0.1% (w/v) sodium azide if the cells are not to be cultured/transplanted after sorting Antibody staining cocktail Flow cytometer/cell sorter supporting multi-color analysis (see Robinson et al., 2011) and FACS tubes 1. Take an aliquot of bone marrow cells before RBC lysis. Add 4 ml FACS buffer and centrifuge the cells 6 min at 300 × g, room temperature. 2. Resuspend the cells in the antibody staining cocktail and incubate for 30 min at 4◦ C. 3. Add 4 ml of FACS buffer and centrifuge the cells 6 min at 300 × g, room temperature. Resuspend the cells and proceed to flow cytometric analysis.

PLATING BONE MARROW IN METHYLCELLULOSE FOR COLONY ASSAYS

SUPPORT PROTOCOL 3

Plating of bone marrow to analyze the resulting colonies can show whether there is an alteration in differentiation potential of progenitor cells resulting in a skewing of colony numbers and/or lineage distribution. Plating cells from littermates of the mice of interest is necessary as a comparison, to assess the whether there is a change in colony numbers and/or lineage preference.

Materials MethoCult methylcellulose medium containing the appropriate cytokine combination (StemCell Technologies) Bone marrow cells (Basic Protocol 5) IMDM medium (e.g., Invitrogen) supplemented with 2% fetal bovine serum (FBS) 16-G needle 3-ml syringe 35-mm and 150-mm Petri dishes (e.g., Becton Dickinson) 37◦ C, 5% CO2 humidified incubator Additional reagents and equipment for red blood cell lysis (see Basic Protocol 5) 1. Thaw methylcellulose medium (MethoCult, provided in tubes). Depending on which colonies the researcher aims to study, methylcellulose containing different cytokine combinations is used. There is a plethora of media commercially available, and to find the optimal conditions it is recommended to refer to the Web site of the methylcellulose provider.

2. Extract the bone marrow cells and perform RBC lysis (see Basic Protocol 5). 3. Vortex the thawed methylcellulose medium to mix. 4. Prepare cells by resuspending them in their appropriate medium (IMDM medium containing 2% FBS) at 10× the final concentration required (i.e., to achieve 1 × 105 cells/dish, prepare a suspension of 1 × 106 cells/ml). The number of bone marrow cells plated depends on the genetic background of the mouse. For a wild-type mouse, approximately 1–2×105 cells/plate are used to obtain a few hundred colonies.

5. Add 0.3 ml of cells to triplicate MethoCult tubes each containg 3 ml of MethoCult.

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6. Vortex the tubes to mix. 7. Let the tubes stand 5 min at room temperature. 8. Attach a 16-G needle to a 3-ml syringe. To remove air from the syringe, draw up ∼1 ml of the cells/methylcellulose medium and gently depress the plunger to expel the solution completely. Repeat until no air is visible. 9. Draw up cells/methylcellulose (1 ml for each 35-mm Petri dish) into a syringe and gently dispense it into the dish. 10. Gently tilt and rotate the dish to evenly spread the solution. 11. Place the 35-mm dishes inside a 150-mm Petri dish containing a dish with 3 ml of water to maintain humidity. 12. Place the cultures in 37◦ C, 5% CO2 , humidified (>95%) incubator. Colonies should start to appear after ∼1 week, and analysis is usually performed after 10 to 14 days. There are detailed descriptions of the morphology of the colonies in the manufacturer’s protocol to help distinguish different kind of colonies. SUPPORT PROTOCOL 4

SERIAL REPLATING Replating colonies in fresh plates (serial replating) will provide an indication of whether the cells of interest have an increased or decreased self-renewal capacity. Normally, wildtype cells can be replated three times, whereas cells with, e.g., an oncogenic mutation can be further replated and produce high numbers of colonies due to their increased self renewal capacity.

Materials MethoCult methylcellulose medium containing the appropriate cytokine combination (StemCell Technologies) Petri dishes containing cells from first methylcellulose plating (Support Protocol 3) RPMI medium (e.g., Invitrogen) supplemented with 10% fetal bovine serum (FBS) 15-ml conical tubes Centrifuge Additional reagents and equipment for plating bone marrow cells in methylcellulose medium (Support Protocol 3) and counting viable cells by trypan blue exclusion (e.g., Sandell and Sakai, 2011) 1. Add 3 ml of MethoCult medium to duplicate 35-mm dishes and gently pipet up and down to suspend the methylcellulose in the medium. 2. Pool the three plates from the first plating in a 15-ml tube, fill the tube with medium (RPMI supplemented with 10% FBS) and centrifuge 8 min at 300 × g, room temperature. 3. Remove the supernatant until 1 ml is remaining in the tube. Add 11 ml medium and vortex for 20 sec. 4. Centrifuge again 8 min at 300 × g, room temperature. 5. Remove the medium until 200 to 500 μl remains, and count viable cells by trypan blue exclusion (e.g., Sandell and Sakai, 2011).

Hematology Testing in Mice

6. Add 3 × 104 cells to a 3-ml aliquot of methylcellulose and add medium (RPMI supplemented with 10% FBS) so that the total volume (cells plus medium) added to the methylcellulose is 300 μl.

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The amount of cells used for replating depends largely on what is being studied. This number is what is usually sufficient for replating wild-type cells. When studying cells with, e.g., a leukemic transformation, lower numbers (e.g., 1 × 104 ) can be used.

7. Follow Support Protocol 3 from step 6. Count and replate colonies every 7 to 10 days, averaging the score for 2 plates.

ISOLATION OF SPLEEN CELLS The mouse spleen (Fig. 6) plays a role in normal hematopoiesis, which is an important difference compared to humans where extramedullary hematopoiesis is a sign of abnormal hematopoiesis in conditions such as anemia. Analysis of the spleen to assess extramedullary hematopoiesis is an important tool for understanding many hematopoietic disorders.

BASIC PROTOCOL 6

Materials Mouse subject Phosphate-buffered saline (PBS) Dissecting instruments 40-μm mesh cell strainers (Becton Dickinson) 6-well tissue culture plates 50-ml conical polypropylene tubes (e.g., BD Falcon) Additional reagents and equipment for euthanasia of the mouse by CO2 asphyxiation (Donovan and Brown, 2006) 1. Euthanize the mouse by CO2 asphyxiation (Donovan and Brown, 2006), keep the mouse fixed on its back, and cut an opening from the pelvic bone to the thorax.

Figure 6

Ventral view of a mouse with the thorax opened. (1) Heart. (2) Spleen.

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2. Separate the left and right side of the skin and, if needed, secure them to the side with needles. 3. Hold the spleen carefully with a forceps and carefully cut away the white tissue to remove the spleen. 4. Weigh the spleen to determine if there is splenomegaly. A normal sized spleen weighs ∼50 to 80mg (in a mouse with a body weight of 20 g). Spleen weight is often denoted as the proportion of spleen weight compared to body weight (% or mg spleen/g mouse).

5. If needed, place the spleen in ice-cold PBS while extracting other organs. 6. Put a 40-μm mesh cell strainer into a well of a 6-well plate with PBS inside. Put the spleen into the strainer, and crush the spleen though the mesh into the well. Wash the filter twice to increase the yield. 7. Add the cell suspension to a 50-ml conical polypropylene tube and wash once to remove debris by centrifuging 8 min at 300 × g, room temperature, adding PBS, centrifuging again as before, and removing the supernatant. One normal sized mouse spleen yields ∼100 × 106 cells. These cells can then be used for analyses and/or transplantation experiments. BASIC PROTOCOL 7

SUBLETHAL/LETHAL IRRADIATION OF MICE In most experiments sublethal or lethal irradiation is performed to prepare a recipient for accepting donor cells. The authors always use lethal irradiation in our transplantation experiments, with “lethal” in this case meaning that if no rescue cells are provided, i.e., donor cells, the mouse will die from the irradiation. This method will provide a 100% chimerism of donor cells in the mouse. If a sublethal approach is used, meaning that the recipient would survive without donor cells, a mouse chimera will be produced with hematopoiesis contribution from both donor and recipient cells, which in almost all cases we are trying to avoid in order get reproducible results.

Materials Mouse subject, 8 to 10 weeks old Medicated water (treated with Napil, available from various drug companies) γ-irradiation source (Cs137 mouse irradiator; Theratronics, http://www.theratronics.ca/) Irradiation-compatible box (Theratronics, http://www.theratronics.ca/) 1. Irradiate the mice with the desired dose. For C57BL/6 mice, 10 to 12 Gy (1000 to 1200 Rad) is commonly used as a lethal dose. To improve tolerance to the irradiation, it is recommended to divide the dose in two separate irradiations separated by 4 hr. Other mouse strains can be more radiosensitive and irradiation dose should be optimized for the strain used. To optimize irradiation dose, one can transplant donor cells with a specific marker (e.g., CD45.1 and CD45.2) into a recipient not carrying this allele and assess at what dose the donor chimerism is 100%. We recommend irradiating the mice on the day before transplantation; however it is also possible to perform irradiation earlier on the same day.

2. Put the mice in cages and maintain them with medicated water. Hematology Testing in Mice

The authors use Nopil, provided by various pharmaceutical companies, which are supplied in 5-ml aliquots to be mixed into the normal drinking water according to the manufacturer’s instructions.

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TAIL VEIN INJECTION Tail vein injection is often used for transplantation of bone marrow cells and also for injections of other fluids into the mouse bloodstream. Performing a successful tail vein injection can be difficult for an inexperienced handler and requires practice and patience. In order to increase the success rate, it is recommended to perform trial injections of saline into mice prior to the experiment. Also, the size of the mouse, the diameter of the tail vein, and pigmentation affects the difficulty of the injection. The lateral veins, which are commonly used for injections, are located on the top sides of the mouse tail.

BASIC PROTOCOL 8

Materials Mouse subject (lethally irradiated; Basic Protocol 7) Cells: e.g., bone marrow cell suspension (Basic Protocol 5) 70% ethanol PBS Heat lamp Mouse restrainer Insulin syringe with 29-G needle (micro-fine, 0.5 ml, Becton Dickinson) 1. Heat the mouse tail, either using a warm compress, by putting the tail in warm water, or by placing a heat lamp in the animal’s cage. When the tail is heated, the veins become more visible. During the heating process carefully monitor the behavior of the mice, as excessive heating is harmful for the animals.

2. Fill a syringe with injectate (e.g., bone marrow cell suspension) and remove visible air bubbles. 3. Place the mouse in a restrainer while holding the tail in a firm grip and swab the tail with 70% ethanol. 4. Identify the lateral vein on either side of the tail (Fig. 7) and inject using the smallest syringe available with a 29-G needle (insulin syringe/needle). Start at the distal part of the tail; thus, if an injection is unsuccessful there is a possibility to try another injection site closer to the body. 5. Insert the needle, while keeping it as flat and as parallel as possible with the tail, and only insert a small part of the needle so that the bevel is inside the vein. Make sure that no air bubbles are inside the syringe.

Figure 7 Mouse tail vein injection. (A) Notice the positioning of the mouse, which is lying on its side in the restrainer to expose the tail vein. (B) After a successful injection a drop of blood should be visible.

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6. Administer the injection. When injecting, it should be possible to see the fluid moving inside the tail vein, and resistance when moving the plunger should be very low. If the resistance is high, it most probably means that the needle is outside the vein. In that case, remove the needle and try again in a different spot more proximal to the animal’s body. The maximum volume used for an injection is determined by the body weight of the animal. As a general rule, the maximum injection volume should not be more than 10% of the animal’s body weight; however, for most injections, a recommended volume for a mouse is ∼0.2 to 0.5 ml, and most users try to limit the injected volume to 0.25 ml. When injecting cells (e.g., bone marrow cells or a sorted population of cells), make sure that the cells are washed with PBS to remove serum before injection.

7. After a successful injection, remove the needle and press on the injection site with a tissue. In most cases, after a successful injection, a small drop of blood can be observed.

8. Return the animal to its cage directly after the injection. SUPPORT PROTOCOL 5

MEASURING HALF LIFE OF PERIPHERAL BLOOD CELLS This method provides an easy way to check for alterations in the half life of peripheral blood cells. The method uses N-hydroxysuccinimide-biotin (NHS-biotin), which is cell-impermeable and thus will reside on the cell surface. This method can be used to assess whether e.g. a mutation or over-expression of a certain protein will affect the half life of hematopoietic cells (lineage or non-linage specific), which might be responsible for an increase in their numbers. A relevant reference may be found at http://www.sciencedirect.com/science/article/pii/S0092867407001961#sec4.3.

Materials Mouse subject 3 μg/μl NHS-biotin (Sigma) in 0.9% NaCl Phosphate-buffered saline (PBS) Insulin syringe with 29-G needle Heat lamp Mouse restrainer FACS tubes (e.g., BD) Additional reagents and equipment for tail vein injection (Basic Protocol 8) and tail vein bleeding (Basic Protocol 1) 1. Using an insulin syringe and 29-G needle, inject 200 μl of 3 μg/μl NHS-biotin (600 μg) into a mouse via tail vein injection (Basic Protocol 8). 2. Obtain blood by tail vein bleeding (Basic Protocol 1). If serial samples are needed within a short time interval, drawing blood from the vein is preferred over cutting the artery. Depending on the lineage of interest, time points for blood collection will vary. Cells with a short half-life, such as platelets and granulocytes, can be examined at 24-hr intervals, and the biotin will be almost entirely gone after 4 to 5 days. When examining cells with a longer half life, such as RBCs, the time point between analyses is longer, e.g., weekly intervals.

3. Label the cells using a streptavidin conjugate for recognizing the biotin-labeled cells together with a lineage marker of interest to allow analysis using flow cytometry. Hematology Testing in Mice

This will allow you to assess the half-life of the lineages of interest, e.g., you can label the sample with CD61 (PE), ter119 (FITC), and streptavidin (APC), which will allow you to

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see the biotin intensity in platelets (CD61) or red blood cells (ter119) by gating on the different fluorochromes. The antibodies for different lineages are described in Table 3, and the staining protocol is described in Basic Protocol 3.

4. Calculate the half-life of cells by measuring the biotin signal repeatedly. The biotin signal will be reduced over time, and the kinetics of decrease are proportional to the half-life of the lineage. Thus, if the half-life of a lineage is altered, the biotin signal can be lost with a more rapid kinetics (decreased half-life) or slower kinetics (increased half-life). This can be quantified by comparing the first time point (denoted baseline) with later time points giving a graph where time (x axis) is plotted against % of baseline signal (y axis).

ISOLATION OF FETAL MOUSE LIVERS The site of hematopoiesis in mice is altered during embryonic development. The fetal liver is the main source of hematopoiesis shortly after E10 and continues at high levels until after birth. Therefore, the fetal liver is an excellent source of hematopoietic cells. This can be especially useful when studying conditions which are lethal during fetal development. The fetal liver cells can be examined and transplanted into an irradiated host in order to look for effects on hematopoiesis (adoptive transfer). When analyzing/sorting cells from fetal liver, be aware of the phenotypic/functional difference between HSCs derived from the fetal liver and bone marrow of adult mice (Morrison et al., 1995; Rebel et al., 1996). An example is that fetal liver HSCs are often positive for Mac-1, which often is used in lineage depletion cocktails for assaying adult HSCs. Also, fetal liver HSCs appear to be more potent in repopulating recipients than adult bone marrow HSCs.

SUPPORT PROTOCOL 6

Materials Mice Phosphate-buffered saline (PBS) RPMI medium (e.g., Invitrogen) supplemented with 10% fetal bovine serum (FBS) Dissecting instruments Petri dishes (e.g., Becton Dickinson) Cell strainers (e.g., BD) Additional reagents and equipment for mating of mice (Ayadi et al., 2011), euthanasia of mice (Donovan and Brown, 2006), and isolation of cells (as for spleen cells; Support Protocol 6) 1. Obtain pregnant mice with desired fetus age by using timed mating, i.e., checking the female mice for copulation plugs to determine when mating occurred (Ayadi et al., 2011). 2. Euthanize the mouse using carbon dioxide asphyxiation (Donovan and Brown, 2006), place it on its back, and fix all four legs by using needles in the paws. 3. Carefully make a fine cut from the lower part of the abdomen up to the throat of the mouse, separate the skin from the fatty tissue, and fix the skin on each side of the mouse using needles. 4. Carefully cut open the fatty tissue to expose the abdomen and secure the tissue at the sides of the mouse using the same needles as for the skin. If the mouse is pregnant, the embryos should be clearly visible, lying in the lower part of the abdomen. They will be connected to each other like beads on a string.

5. Carefully remove the embryos and put them in a Petri dish containing PBS. Hematology Testing in Mice

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6. Extract a fetus from the amniotic sac. The fetal liver should be clearly visible as a bright red spot inside the otherwise white/beige fetus.

7. Carefully open the fetus with small scissors and scrape/cut to remove the fetal liver. 8. Put the fetal liver in RPMI medium containing 10% FBS, on ice. 9. Extract the cells following the same protocol as described for the spleen (Basic Protocol 6).

REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps.

Red blood cell (RBC) lysis buffer 10× stock solution: 1.5 M NH4 Cl 0.1 M KHCO3 1 mM Na2 EDTA Adjust pH to 7.4 Store up to several months at 4◦ C 1× working solution: Dilute 10× stock 1:9 with water just before use COMMENTARY Background Information The mouse is often used to model steadystate and aberrant hematopoiesis. Described here are basic methods that are commonly used in laboratories working with mouse models to study hematopoiesis. There are several mouse strains that can be used for this purpose, and the flow cytometry scattergrams in this protocol were achieved using a Bl6/C57J mouse. There is variation between different mouse strains, which should be taken into consideration when examining data, but Bl6/C57J is the most commonly used mouse strain for examining hematopoiesis—it is the strain depicted in the figures of this unit.

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Tail vein bleeding This method (Basic Protocol 1) will result in a small opening in the mouse tail vein/artery and allows a relatively noninvasive way to draw blood from a mouse without the need for anesthetics. Depending on the volume of blood drawn from the mouse, the time interval between obtaining blood samples may vary, but, in general, obtaining samples every 4 to 6 weeks will allow enough blood to be drawn for a CBC and flow cytometry analysis. The CBC provides the most important parameters for understanding the hematological status of a mouse. Blood smear This method (Basic Protocol 2) yields a thin film of peripheral blood on an objective slide. It

was previously used to allow counting of cells, which currently in most cases is replaced by the automated CBC. However, a blood smear is still a valuable assay for understanding parameters that are difficult to evaluate solely with a CBC. Primarily, morphological abnormalities are routinely analyzed using a blood smear, but other parameters can also be analyzed. The advantage of a blood smear is that small amounts (a small drop of blood) can yield a lot of information and the procedure can be performed at short time intervals for tracking rapid alterations. Flow cytometry analysis and FACS of peripheral blood and bone marrow/spleen These systems allow for analysis and sorting of defined cellular populations in the peripheral blood (Basic Protocol 3) or bone marrow/spleen (Support Protocol 1), as well as cells obtained from other tissues/organs at a single-cell level. Cardiac puncture This method (Basic Protocol 4) will allow an experienced experimenter to attain a large quantity of aseptic blood from a euthanized mouse. Isolation and transplantation of bone marrow cells The mouse bone marrow is the primary site for hematopoiesis, and this method (Basic Current Protocols in Mouse Biology

Protocol 5) describes how to remove the bones from a mouse to allow extraction of bone marrow cells, which can be used for transplantation into recipient mice.

field of view. This is because platelets can aggregate and accumulate at the upper part of the slide, whereas neutrophils can accumulate at the lower part of the smear.

Isolation of spleen cells The spleen is the main source of extramedullary hematopoiesis, so evaluating hematopoiesis by isolating spleen cells (Basic Protocol 6) is of importance for understanding this organ’s contribution to medullary hematopoiesis.

Flow cytometry analysis and FACS of peripheral blood (Basic Protocol 3) When analyzing platelets, it is important that the size cutoff (forward/side scatter) not be set at default levels, where the platelets can be excluded as debris.

Sublethal/lethal irradiation of mice Performing sublethal/lethal irradiation (Basic Protocol 7) of mice is necessary to prepare recipient mice for accepting bone marrow transplantation from a donor mouse. Tail vein injection This method (Basic Protocol 8) allows injection of substances into a mouse’s bloodstream. It is often used for transplantation of bone marrow cells into an irradiated recipient, since it is technically less difficult as compared to intrafemoral injection and does not require anesthetics.

Critical Parameters and Troubleshooting Tail vein bleeding (Basic Protocol 1) It is important that the mouse be warmed to a reasonable level to improve blood flow, while not heating it so much that it is harmful for the animal. One must carefully monitor the behavior of the mice while warming them. If the animals start to become stressed while being warmed, reduce the intensity of the lamp or turn it off for a short while. Depicted in Figure 1 is arterial bleeding of the mouse. The arteries are situated on the lower (ventral) and upper (dorsal) part of the tail. When cutting the artery, it is important to apply pressure for an extended period of time (up to a minute) to prevent excessive bleeding. If smaller amounts of blood are needed, it is possible to cut the veins on either side of the mouse (the location of the veins is shown in Figure 7 for tail vein injection). Cutting the vein does not affect hemostasis in the same way as arterial bleeding. For either bleeding method, it is important to properly mix the sample with EDTA to avoid clotting of the blood. Proper mixing should be achieved by inverting the tube several times, not by shaking. Blood smear (Basic Protocol 2) It is important to locate an area in the middle of the smear to identify a representative

Cardiac puncture (Basic Protocol 4) Performing a cardiac puncture is technically demanding and requires extensive experience for reproducible results. It is recommended to practice the procedure before the experiment is performed. It is important that the cardiac puncture be performed as soon as possible after euthanizing the mouse. If there is delay, the amount of blood collected will be lower and there is an increased risk of blood clotting. Remove the needle from the syringe when transferring the blood to the tube containing EDTA. Pressing the blood through the thin opening of the needle can increase the chances of coagulation and can cause artifacts. A common problem with cardiac puncture is coagulation of the blood within the syringe. Therefore, it is important to transfer the blood from the syringe to the tube containing EDTA as quickly as possible. When performing a cardiac puncture, estimate beforehand how much blood will be needed for the subsequent analyses. Since the speed of the procedure is of importance, once sufficient blood is collected for the specific analyses, retract the needle and dispense the blood into the tube containing EDTA. Isolation and transplantation of bone marrow cells (Basic Protocol 5) Always keep the bones and extracted bone marrow at 4◦ C on ice to maximize the viability of the cells. Also, after the bone marrow is extracted, it is important that these cells be kept in media supplemented with FBS at 4◦ C. Be careful when extracting the bones and cleaning them of tissue since they are fragile, especially when handling old or fibrotic bones. Sublethal/lethal irradiation of mice (Basic Protocol 7) Each mouse strain has a varying degree of radiosensitivity. Therefore, it is recommended to titrate the irradiation dose needed for each individual mouse that will be used in

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experiments. This can be done by evaluating the chimerism of the mouse after reconstitution by differentiating the donor/recipient cells using surface markers such as CD45.1 and CD45.2. Carefully monitor the mice after irradiation. If the mice are experiencing irradiation-related damage, such as severe skin inflammation, they should be euthanized. Tail vein injection (Basic Protocol 8) Similar to the cardiac puncture, “dry runs" using PBS should be performed to perfect the injection technique before undertaking the experiment. If there is a resistance when injecting, do not force the plunger. When the needle is inserted correctly, there will be no resistance. Flow cytometry analysis/sorting of bone marrow stem/progenitor cells (Support Protocol 1) For flow cytometry of bone marrow stem/ progenitor cells, an appropriate control for lineage-positive cells must be included for appropriate setting of the lineage-negative gate. Appropriate single-fluorochrome stains should always be included for compensation, which in the case of analyzing stem/progenitor cells is of utmost importance. This is because several fluorochromes are used, so suboptimal compensation will have severe effects. Secondly, different populations are often slightly shifted with respect to the intensity of a fluorescent dye; thus, not having the correct compensations will result in impure populations of cells.

Anticipated Results The goal of the methods provided here is to obtain a general assessment of hematopoiesis in the animal model studied. Any significant changes in abundance or morphology of blood cells will be recognized when performing a CBC and blood smear. There may be many reasons why a change is observed, and at least some alterations observed in the peripheral blood will be better understood by performing a more in-depth flow cytometry analysis of the bone marrow and spleen stem/progenitor compartments. The goal of bone marrow transplantations is to assess whether the phenotype observed in the primary animal is transplantable into a secondary recipient, i.e., transmitted by a cell with extended self-renewal capacity. Hematology Testing in Mice

Cardiac puncture (Basic Protocol 4) One can anticipate obtaining ∼500 to 1000 μl of blood from a normal-sized mouse

from a successful cardiac puncture. With a larger mouse, more blood can be collected. Isolation and transplantation of bone marrow cells (Basic Protocol 5) One can anticipate obtaining ∼100 to 150 × 106 cells after RBC lysis from a single mouse when extracting blood from the femora, tibiae, and the two pelvic bones. The viability after extraction should be >80%. Flow cytometry analysis/sorting of bone marrow stem/progenitor cells If using a linage depletion kit the majority (>80%) of cells should be lineagenegative. Regarding the individual populations, there is a wide discrepancy between various mouse strains (Papathanasiou et al., 2010). The LSK population (lin– , c-Kit+ , and Sca-1+ ) usually accounts for ∼1% of lineagenegative cells, but genetic background and age (Kim et al., 2003) of the mouse can increase/decrease the population substantially. The specific subpopulations are variable depending on markers used and gating strategy. In the LSK population, the long-term stem cells (LT-HSC) are the least abundant population, and usually constitute a few percent of the total LSK population. Two recent reports describing gating strategies of stem progenitor cells are Aucagne et al. (2011) and Quere et al. (2011). It should be noted that what is termed the LT-HSC population is still a quite heterogenous population, and in Table 4, CD150 is in brackets because it has recently been shown that this marker affects the myeloid/lymphoid priming of the stem cells (Morita et al., 2010).

Time Considerations In some of the protocols described here (such as isolation of bone marrow cells), the time required is highly dependent on the level of expertise of the experimenter, whereas some protocols, such as performing a blood smear, are very simple even for an inexperienced experimenter. The time considerations described here are based on the time it that is expected to take for an inexperienced user to perform the method. In the cases where the time required for flow cytometry analyses is taken into account, the time needed for compensations and setting up the gating is not accounted for. Tail vein bleeding (Basic Protocol 1) Expect that heating up the mice will take about 1 min. Cutting the tail and collecting the blood also takes ∼1 min per mouse.

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Usually, several mice are kept under the heat lamp simultaneously to speed the process. Blood smear (Basic Protocol 2) Allow a few minutes to perform the smear and for the blood to dry. Staining the sample takes < 1 min, and a few minutes for the sample to dry. Flow cytometry analysis and FACS of peripheral blood and bone marrow (Basic Protocol 3 and Support Protocol 1) The time required for acquisition/collection of cells depends on the number of cells needed. For the time consideration described here, acquisition of a few thousand cells is accounted for. For samples where no red blood cell lysis is needed (platelets and erythrocytes), expect the incubation with antibody and washing to take ∼30 min. Since both platelets and erythrocytes are abundant cells, sample acquisition takes a few seconds for erythrocytes and ∼10 to 15 sec for platelets. For the lineages where RBC lysis is performed, expect the incubation with antibody, red blood cell lysis, and washing to take ∼1 hr. Sample acquisition takes ∼1 to 5 min/sample depending on the abundance of the population. Cardiac puncture (Basic Protocol 2) After euthanizing the mouse, the procedure from start until end should take no longer than 10 sec/mouse. Isolation and transplantation of bone marrow cells (Basic Protocol 5) Extraction of the bones takes ∼20 min for one mouse. The crushing, washing, red blood cell lysis, and additional washing take another ∼40 min. Sublethal/lethal irradiation of mice (Basic Protocol 7) The time needed for irradiation is hard to estimate since it is almost exclusively dependent on the irradiation source. However, it is recommended to allow the mice to rest for 4 hr between irradiation doses; thus, performing the first irradiation early in the day is recommended. Tail vein injection (Basic Protocol 8) The time required for tail vein injection is totally dependent on the experience level of the experimenter. For an inexperienced handler, it is not unusual to have to spend several minutes per mouse in order to achieve a successful injection. The tail-heating time is similar to that for tail vein bleeding, ∼1 min.

Flow cytometry analysis/sorting of bone marrow stem/progenitor cells (Support Protocol 1) Flow cytometry analysis/sorting of bone marrow/progenitor cells is predominantly performed on fresh material; thus, the time for the isolation of bone marrow cells must be included, which can be several hours when working with a large number of animals. For flow cytometry, lineage depletion can take ∼2 hr, and antibody incubation and washing ∼40 min (∼70 min if a secondary antibody is used, which is often the case). The analysis/sorting step can take considerable time, especially when analyzing/sorting rare stem cell populations, or when analyzing without using lineage depletion. Thus, often several hours are needed for the analysis/sorting step.

Literature Cited Aucagne, R., Droin, N., Paggetti, J., Lagrange, B., Largeot, A., Hammann, A., Bataille, A., Martin, L., Yan, K.P., Fenaux, P., Losson, R., Solary, E., Bastie, J.N., and Delva, L. 2011. Transcription intermediary factor 1 gamma is a tumor suppressor in mouse and human chronic myelomonocytic leukemia. J. Clin. Invest. 121:2361-2370. Ayadi, A., Ferrand, G., Gonc¸alves la Cruz, I., and Warot, X. 2011. Mouse breeding and colony management. Curr. Protoc. Mouse Biol. 1:239264. Baertschi, B. and Gyger, M. 2011. Ethical considerations in mouse experiments. Curr. Protoc. in Mouse Biol. 1:155-167. Donovan, J. and Brown, P. 2006. Euthanasia. Curr. Protoc. Immunol. 73:1.8.1-1.8.4. Greiner, D.L., Shultz, L.D., Yates, J., Appel, M.C., Perdrizet, G., Hesselton, R.M., Schweitzer, I., Beamer, W.G., Shultz, K.L., Pelsue, S.C., et al. 1995. Improved engraftment of human spleen cells in NOD/LtSz-scid/scid mice as compared with C.B-17-scid/scid mice. Am. J. Pathol. 146:888-902. Ito, M., Hiramatsu, H., Kobayashi, K., Suzue, K., Kawahata, M., Hioki, K., Ueyama, Y., Koyanagi, Y., Sugamura, K., Tsuji, K., Heike, T., and Nakahata, T. 2002. NOD/SCID/gamma(c)(null) mouse: An excellent recipient mouse model for engraftment of human cells. Blood 100:31753182. Jaenisch, R. and Mintz, B. 1974. Simian virus 40 DNA sequences in DNA of healthy adult mice derived from preimplantation blastocysts injected with viral DNA. Proc. Natl. Acad. Sci. U.S.A. 14:1250-1254. Kim, M., Moon, H.B., and Spangrude, G.J. 2003. Major age-related changes of mouse hematopoietic stem/progenitor cells. Ann. N.Y. Acad. Sci. 996:195-208. McCune, J.M., Namikawa, R., Kaneshima, H., Shultz, L.D., Lieberman, M., and Weissman, I.L.

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1988. The SCID-hu mouse: Murine model for the analysis of human hematolymphoid differentiation and function. Science 241:1632-1639. Morita, Y., Ema, H., and Nakauchi, H. 2010. Heterogeneity and hierarchy within the most primitive hematopoietic stem cell compartment. J. Exp. Med. 207:1173-1182. Morrison, S.J., Hemmati, H.D., Wandycz, A.M., and Weissman, I.L. 1995. The purification and characterization of fetal liver hematopoietic stem cells. Proc. Natl. Acad. Sci. U.S.A. 92:10302-10306. Mosier, D.E., Gulizia, R.J., Baird, S.M., and Wilson, D.B. 1988. Transfer of a functional human immune system to mice with severe combined immunodeficiency. Nature 335:256-259. Muller-Sieburg, C.E., Whitlock, C.A., and Weissman, I.L. 1986. Isolation of two early B lymphocyte progenitors from mouse marrow: A committed pre-pre-B cell and a clonogenic Thy1-lo hematopoietic stem cell. Cell 444:653-662. Papathanasiou, P., Tunningley, R., Pattabiraman, D.R., Ye, P., Gonda, T.J., Whittle, B., Hamilton, A.E., Cridland, S.O., Lourie, R., and Perkins, A.C. 2010. A recessive screen for genes regulating hematopoietic stem cells. Blood 116:58495858.

Quere, R., Karlsson, G., Hertwig, F., Rissler, M., Lindqvist, B., Fioretos, T., Vandenberghe, P., Slovak, M.L., Cammenga, J., and Karlsson, S. 2011. Smad4 binds Hoxa9 in the cytoplasm and protects primitive hematopoietic cells against nuclear activation by Hoxa9 and leukemia transformation. Blood 117:59185930. Rebel, V.I., Miller, C.L., Eaves, C.J., and Lansdorp, P.M. 1996. The repopulation potential of fetal liver hematopoietic stem cells in mice exceeds that of their liver adult bone marrow counterparts. Blood 878:3500-3507. Robinson, J.P., Darzynkiewicz, Z., Hoffman, R., Nolan, J.P., Orfao, A., Rabinovitch, P.S., and Watkins, S. (eds.) 2011. Current Protocols in Cytometry. John Wiley & Sons, Hoboken, N.J. Sandell, A. and Sakai, D. 2011. Mammalian cell culture. Curr. Protoc. Essen. Lab. Tech. 5:4.3.14.3.32 Shultz, L.D., Ishikawa, F., and Greiner, D.L. 2007. Humanized mice in translational biomedical research. Nat. Rev. Immunol. 72:118-130. Spangrude, G.J., Heimfeld, S., and Weissman, I.L. 1988. Purification and characterization of mouse hematopoietic stem cells. Science 241:5862.

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A Comprehensive Guide to Sleeping Beauty–Based Somatic Transposon Mutagenesis in the Mouse Branden Moriarity1,2,3 and David A. Largaespada1,2,3 1

Department of Genetics, Cell Biology, and Development, University of Minnesota, Minneapolis, Minnesota 2 Center for Genome Engineering, University of Minnesota, Minneapolis, Minnesota 3 Masonic Cancer Center, University of Minnesota, Minneapolis, Minnesota

ABSTRACT Recent advances in whole genome analyses made possible by next-generation DNA sequencing, high-density array comparative genome hybridization (aCGH), and other technologies have made it apparent that cancers harbor numerous genomic changes. However, without functional correlation or validation, it has proven difficult to determine which genetic changes are necessary or sufficient to produce cancer. Thus, it is still necessary to perform unbiased functional studies using model organisms to help interpret the results of whole genome analyses of human tumors. To this end, a Sleeping Beauty (SB) transposon–based mutagenesis technology was developed to identify genes that, when mutated, can cause cancer. Herein a detailed methodology to initiate and carry out an SB transposon mutagenesis screen is described. Although this system might be used to identify genes involved with many cellular phenotypes, it has been primarily implemented for cancer. Thus, SB transposon somatic cell screens for cancer development are C 2011 by John Wiley & Sons, Inc. highlighted. Curr. Protoc. Mouse Biol. 1:347-368  Keywords: Sleeping Beauty r cancer r mutagenesis screen r transposon r mouse

INTRODUCTION Forward genetic screens have been utilized for many years in model organisms using a variety of different mutagens. An ideal mutagen is one that randomly mutates the genome and induces gene gain-of-function (GOF) or loss-of-function (LOF) mutations that are easily trackable. One mutagen that fulfills these criteria is an engineered DNA transposon system, termed Sleeping Beauty (Collier et al., 2005; Dupuy et al., 2005). Sleeping Beauty (SB) is a member of the Tc1/Mariner family of DNA transposable elements that utilizes a cut-and-paste mechanism of mobilization (Ivics et al., 1997). SB is a two-component system comprising a DNA element termed transposon vector and an enzyme component termed transposase. To date, this system has been successfully used to model and identify candidate genes in many types of cancer, including: colorectal (Starr et al., 2009), liver (Keng et al., 2009), lymphoma/leukemia (Dupuy et al., 2005, 2009), and sarcoma (Collier et al., 2005). Each of these studies identified a list of common insertion sites (CISs) that harbored candidate oncogenes or tumor suppressors that were then further investigated and validated. Thus, it has been clearly demonstrated that the SB system is an invaluable tool for cancer gene identification in the mouse. Herein, three basic protocols with detailed methodology to initiate, validate, and carry out an SB forward genetic screen in somatic cells of mice are described. Basic Protocol 1 describes how to properly breed, genotype, and manage a cohort of experimental and control animals. Basic Protocol 2 covers methods necessary to validate that transposon mutagenesis is occurring in the tissue and cell type of interest using Current Protocols in Mouse Biology 1: 347-368, September 2011 Published online September 2011 in Wiley Online Library (wileyonlinelibrary.com). DOI: 10.1002/9780470942390.mo110087 C 2011 John Wiley & Sons, Inc. Copyright 

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immunohistochemistry (IHC) and polymerase chain reaction (PCR)–based techniques. Basic Protocol 3 describes the method of ligation-mediated PCR (LM-PCR) for identification of transposon insertion sites utilizing the high-throughput Illumina sequencing platform.

STRATEGIC PLANNING Due to the large amount of time and financial investment that must go into an SB forward genetic screen, it is recommended to lay out a plan before beginning the experiment. Choosing the best-suited transgenic mouse strains based on the known information about the cancer of interest is the best way to ensure a successful screen. Additionally, knowing the cell of origin and the types of promoters that are active in that cell type is of utmost importance. Herein, multiple aspects of planning for an SB screen that can greatly increase the likelihood of a fruitful experiment are described. To adequately plan for the screen, the details of the SB system and how it works as an effective mutagen using the T2/Onc transposon must be understood thoroughly. SB is a member of the Tc1/Mariner family of transposable elements and utilizes a cutand-paste mechanism of mobilization. SB had been evolutionarily inactivated and was recently molecularly resurrected by compiling a consensus amino acid sequence from the salmonid subfamily of elements (Ivics et al., 1997). It is a two-component system comprising a DNA element termed transposon vector and an enzyme component termed transposase. The transposon vector is identified by the SB transposase via flanking unique inverted repeat/direct repeat (IR/DR) DNA sequences and is excised, followed by reintegration into any TA dinucleotide. The TA dinucleotide is the only strict requirement for transposon integration. To equip the SB transposon for somatic mutagenesis in the mouse, its original cargo, the transposase open reading frame, was replaced with mutagenic cargo. Splice acceptors followed by poly-adenylation signals were engineered in both orientations for gene inactivation while the murine stem cell virus (MSCV) 5 LTR followed by a splice donor was engineered to over express genes. This first generation of the mutagenic transposon was termed T2/Onc (Collier et al., 2005; Dupuy et al., 2005). Since SB is a cut-andpaste transposon, it was apparent that many copies of T2/Onc would likely be needed per cell to induce cancer development in the mouse, as cancer is known to be a multihit or multi-step process. Using standard transgenesis methods of pronuclear injection of the linearized T2/Onc plasmid transposon, concatemer lines with many copies of T2/Onc were developed, ranging in copy numbers from 25 to 300 copies (Collier et al., 2005; Dupuy et al., 2005). This mutagenic transposon system was first engineered and implemented for cancer gene discovery in the authors’ laboratory and has now been used by many others for the same purpose in many types of cancer (Dupuy et al., 2009; Rahrmann et al., 2009; Starr et al., 2011). Recently, other versions of the mutagenic transposon and SB have been developed that can also be considered for implementation in SB forward genetic screens for cancer gene identification and are discussed below (Rad et al., 2010).

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Choosing a mutagenic transposon line In the last decade, many transgenic lines have been developed with different transposon concatemers for SB mutagenesis (Dupuy et al., 2009; Rad et al., 2010). The main differences among the transposon concatemer lines are copy number, chromosomal location, and mutagenic cargo. All of these variables should be considered before any screen is initiated. Most available transposon concatemer lines have similar splice acceptors and poly-adenylation signals to inactivate genes, and to date there has not been a study comparing alternative elements. However, several different promoters for overexpression of

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genes have been investigated and it has been observed that depending on the tissue/cell type of interest one promoter may be superior to another. For instance, it was found that the cytomegalovirus immediate enhancer with the chicken beta-globin promoter (CAG) is more efficient in generating solid tumors and the murine stem cell virus (MSCV) 5 LTR was superior in developing hematopoietic cancers, such as leukemia (Dupuy et al., 2009). The human phosphoglycerate kinase (hPGK) promoter was found to be an intermediate that could be used to induce both solid and blood cancers (Rad et al., 2010). If a promoter that is highly active in the tissue or cell type of interest can be found, then it may be a candidate for the mutagenic promoter. It is advisable to use two different transposon concatemer transgenic lines that reside on different chromosomes in designing a particular experiment. This is recommended because, upon analysis of CISs, the chromosome harboring the concatemer is usually discarded from the data due to the local hopping phenomenon associated with SB transposition (Keng et al., 2009; Starr et al., 2009). Thus, concatemers on different chromosomes will complement each other and allow for coverage of the entire genome. Lastly, there has not been a direct investigation comparing transposon copy number in the concatemer and the rate of gene discovery or phenotype development. However, in the authors’ experience, more copies of the mutagenic transposon can decrease the latency of the phenotype development in some instances (B. Moriarity, unpub. observ.).

Choosing a transposase source The pioneering transposon somatic mutagenesis screens in mice used a constitutively expressed version of the SB transposase under the control of the CAG or endogenous Rosa26 promoters (Collier et al., 2005; Dupuy et al., 2005). With this ubiquitous mutagenesis, mice developed a small number of different tumor types composed mostly of very rapidly developing T cell leukemia/lymphoma using the Rosa26-SB11 line and low penetrance sarcoma only in cancer predisposed p19–/– mice using the CAG-SB10 line. Since these studies, a conditional SB transposase transgenic has been developed with the transposase under the control of the Rosa26 promoter, termed Rosa26-LSL-SB11 (Dupuy et al., 2009). This allele is conditional by virtue of a floxed eGFP stop cassette (LSL) upstream of the SB coding sequence. This transgene has been successfully used in conjunction with tissue-specific-Cre recombinase (TSP-Cre) transgenes to mutagenize selected cell types. Many available TSP-Cre transgenics can be found in the Cre-XMice database (http://nagy.mshri.on.ca/cre new/index.php). Thus, if there is a TSP-Cre available for the tissue of interest, then the Rosa26-LSL-SB11 is likely the transposase source of choice. However, in the unlikely event that the Rosa26 promoter is not active in the tissue/cell type of interest, or if a TSP-Cre is not available, then another source of transposase may be necessary; a knock-in or transgenic expressing SB from a promoter known to be active in the cell type of interest may be necessary. Is a cancer predisposed genetic background necessary? In some cases, it may be necessary or desirable to use SB mutagenesis in conjunction with a predisposing genetic background. One reason to use a predisposing background is to screen for genes that cooperate with the predisposing allele, such as a fusion gene prominent in cancer where other genetic aberrations are still unknown. However, the main reason for using a predisposing background is that in some cases SB mutagenesis is not sufficient to induce the phenotype of interest. There are numerous reasons why SB mutagenesis may not be a strong enough mutagen to induce highly penetrant cancer. First, too small a target cell population undergoing mutagenesis can be problematic, as SB mutagenesis is random and therefore the more cells available to be transformed, the more likely the event will occur. Additionally, if the Rosa26 promoter is under expressed, leading to low SB expression, or the mutagenic transposon promoter is inactive or weakly

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active in the cell type of interest, the screen may not be successful. Importantly, most of these hurdles can be tested before a full screen is undertaken. For instance, the TSP-Cre of interest can be crossed to the Rosa26-LSL-βgal and the cell type of interest tested for β-gal expression by IHC (Soriano, 1999). Thus, if it is known that the phenotype of interest may suffer from one of the aforementioned hurdles, then it is advisable to use a predisposing background along with a mutagenesis-only cohort of animals.

How many mice to breed? Determining the optimal number of experimental and control animals can be one of the most difficult decisions when setting up a forward genetic screen. Many questions need to be considered. What is the predicted penetrance? Will there be background tumors that will decrease the number of tumors of interest? Will animals develop multi-focal disease? As a starting point, it is advisable to breed at least the same number of controls as experimental animals, ideally even more than experimental animals. Many of the considerations above may be difficult or impossible to predict, so it is usually best to breed more animals than needed, if possible. If there are too few tumors, then it may not be possible to assemble a meaningful list of CISs. Most published SB screens breed enough mice to obtain at least 20 tumors and sometimes as many as 200, to have enough independent tumors to acquire a statistically significant CIS list (Collier et al., 2005, 2009; Dupuy et al., 2005, 2009; Keng et al., 2009; Starr et al., 2009). Importantly, enough mice must be generated to determine if SB mutagenesis results in acceleration or spontaneous tumor development. Using the Peto-Peto-Wilson statistic for comparing Kaplan-Meier survival curves, there is 80% power to detect a two-fold difference in cancer latency with a p-value ≤0.05 if there are at least 60 animals in both the control and experimental group (Statview). The authors consider 80% power of detection acceptable and a twofold difference in cancer latency likely to be biologically meaningful. Based on this knowledge, a good starting point for any SB mutagenesis screen is to breed at least 60 control and experimental animals. NOTE: All protocols using live animals must first be reviewed and approved by an Institutional Animal Care and Use Committee (IACUC) or must conform to governmental regulations regarding the care and use of laboratory animals. BASIC PROTOCOL 1

BREEDING AND GENOTYPING A COHORT OF MICE UNDERGOING SOMATIC TRANSPOSON MUTAGENESIS This protocol outlines methods to generate and genotype a cohort of control and experimental animals necessary for a tissue specific (TSP) forward genetic screen using SB transposon-mediated mutagenesis with or without the addition of a predisposing genetic background. Tissue specificity is accomplished using the conditional R26-LSLSB11 transgenic (Dupuy et al., 2009). The predisposing background can be a canonical knock-out, overexpression allele, or ideally a conditional version of either type.

Materials

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Breeding pairs of all necessary transgenic mice (8- to 20-weeks of age) R26-LSL-SB11 (Dupuy et al., 2009) T2/Onc Concatemer (Collier et al., 2005; Dupuy et al., 2005) TSP-Cre recombinase (http://nagy.mshri.on.ca/cre new/index.php) Conditional or non-conditional predisposing transgene (optional) IACUC-approved food and water SDS lysis buffer (see recipe) Proteinase K (see recipe) Phenol (Sigma-Aldrich) Chloroform (Sigma-Aldrich) Isopropanol, ice cold Current Protocols in Mouse Biology

70% ethanol TE buffer (see recipe) 1.1× ReddyMix (Thermo Scientific) Primers (Table 1) Genomic DNA (negative and positive controls) 1% agarose gel in 1× TAE buffer 1× TAE (from 50× stock) (see recipe) 100-bp Quanti-Marker (BioExpress) Ethidium bromide (10 mg/ml) (Sigma-Aldrich) IACUC-approved animal housing facility IACUC-approved animal cages and bedding Sterile 1.5-ml microcentrifuge tubes 55◦ C shaking incubator Microcentrifuge, room temperature and 4◦ C Spectrophotometer 0.5-ml thin-walled PCR tubes Thermal cycler Gel electrophoresis apparatus and power source UV light box Gel photography equipment Table 1 Primer List

Primer

Sequence 5 -3a

R26-LSL-WT reverse

CCCCAGATGACTACCTATCCTCCC

R26-LSL-WT forward

CTGTTTTGGAGGCAGGAA

R26-LSL-SB11 reverse

CTAAAAGGCCTATCACAAAC

T2/Onc forward

CGCTTCTCGCTTCTGTTCGC

T2/Onc reverse

CCACCCCCAGCATTCTAGTT

Excision assay forward

TGTGCTGCAAGGCGATTA

Excision assay reverse

ACCATGATTACGCCAAGC

Generic Cre forward

TTCGGCTATACGTAACAGGG

Generic Cre reverse

TCGATGCAACGAGTGATGAG

Bfal linker+

GTAATACGACTCACTATAGGGCTCCGCTTAAGGGAC

Bfal linker−

P-TAGTCCCTTAAGCGGAG-AM

NlaIII linker+

GTAATACGACTCACTATAGGGCTCCGCTTAAGGGACCATG

NlaIII linker−

P-GTCCCTTAAGCGGAGCC-AM

Primary Splink IRDR right

GCTTGTGGAAGGCTACTCGAAATGTTTGACCC

Primary Splink IRDR left

CTGGAATTTTCCAAGCTGTTTAAAGGCACAGTCAAC

Primary Splink linker

GTAATACGACTCACTATAGGGC

Secondary Splink IRDR right (Illumina)

AATGATACGGCGACCACCGAGATCTACACTCTTTCCCTACACGACGCTCTTCCGATCT(N)10 AAGTGTATGTAAACTTCCGACTTCAA

Secondary Splink IRDR left (Illumina)

AATGATACGGCGACCACCGAGATCTACACTCTTTCCCTACACGACGCTCTTCCGATCT(N)10 AGGTGTATGTAAACTTCCGACTTCAA

Secondary linker (Illumina)

CAAGCAGAAGACGGCATACGAGCTCTTCCGATCTAGGGCTCCGCTTAAGGGAC

a Abbreviations: P, phosphorylation; AM, amino modifier.

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Generate experimental and control animals 1. Begin by acquiring the appropriate transgenic animals needed to perform the SB forward genetic screen and decide if a predisposing background is desired (see Strategic Planning): R26-LSL-SB11; concatemer; TSP-Cre recombinase; conditional or nonconditional predisposing transgene (optional). Provide food and water ad libitum. 2. Before beginning any breeding, determine if a predisposing background is desired. If so, it is further necessary to know if the predisposing background allele is a conditional, i.e., LoxP flanked, as this can change the breeding strategy. The misexpression of some TSP-Cre transgenics in male germ cells of the testes could cause the recombined allele to become activated in the germline, and this allele therefore becomes a body-wide non-conditional allele in the offspring. Thus, it is necessary to keep the TSP-Cre and conditional alleles separate when breeding. 3a. For screens using no predisposing background, begin by breeding R26-LSL-SB11 mice to T2/Onc mice to generate animals that are homozygous for both alleles. These mice can then be intercrossed to TSP-Cre mice to obtain experimental animals (see Fig. 1A). 3b. For screens using non-conditional predisposing background, begin by breeding R26LSL-SB11 mice to T2/Onc mice to generate animals that are homozygous for both alleles. Concurrently breed TSP-Cre mice to non-conditional predisposing allele mice to generate animals that are homozygous for the non-conditional allele. These mice can then be intercrossed to obtain experimental animals (see Fig. 1B). 3c. For screens using a conditional predisposing background, begin by breeding R26LSL-SB11 mice to the conditional predisposing mice to generate animals that are homozygous for both alleles. Concurrently breed TSP-Cre mice to T2/Onc mice to generate animals that are homozygous for T2/Onc. These mice can then be intercrossed to obtain experimental animals (see Fig. 1C). Note that it may not always be possible to maintain predisposing alleles as homozygotes due to lethality; in this case, the frequency of experimental mice will be reduced and attaining all experimental animals will require more breeding.

Extract genomic DNA from tail sample 4. Upon weaning, collect a small (∼0.5-cm) piece of tail in a 1.5-ml microcentrifuge tube for DNA extraction. Be sure to use the appropriate IACUC protocol for tail clipping specific to the institution, as protocols can vary from one institution to another.

5. Add 490 μl SDS lysis buffer and 10 μl proteinase K (10 mg/ml) to each sample and incubate overnight at 55◦ C with shaking at 200 rpm. 6. After overnight digestion, centrifuge 5 min at 20,000 × g, room temperature, to pellet remaining tail debris. 7. Transfer liquid to a fresh 1.5-ml microcentrifuge tube.

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Figure 1 (appears on following page) Outline of crosses necessary to generate experimental class animals undergoing transposon mutagenesis. (A,B) Experimental class animals for screens utilizing no predisposing background or a non-conditional background are generated by breeding animals homozygous for both R26-LSL-SB11 and the T2/Onc concatemer to animals that are heterozygous for the Cre (TSP-Cre) of interest or homozygous for the non-conditional predisposing background and heterozygous for the TSP-Cre, respectively. (C) Experimental class animals for screens utilizing a conditional predisposing background are generated by breeding animals homozygous for both R26-LSL-SB11 and the conditional predisposing background to mice homozygous for the T2/Onc concatemer and heterozygous for the TSP-Cre of interest. Current Protocols in Mouse Biology

A.

TSP-Cre

TSP-Cre

R26-LSL-SB11

R26-LSL-SB11

T2/Onc

R26-LSL-SB11

T2/Onc

T2/Onc

B.

Tg

TSP-Cre

Tg

TSP-Cre

R26-LSL-SB11

T2/Onc

R26-LSL-SB11

T2/Onc

R26-LSL-SB11

T2/Onc

Tg

C.

T2/Onc T2/Onc

TSP-Cre

Figure 1

R26-LSL-SB11 R26-LSL-SB11

TSP-Cre

R26-LSL-SB11

(legend appears on preceding page)

T2/Onc

LSL-Tg

LSL-Tg LSL-Tg

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8. Add 500 μl of 1:1 phenol/chloroform and gently invert tubes 10 to 20 times. Perform organic extraction in a properly ventilated hood.

9. Centrifuge 5 min at 20,000 × g, room temperature, to separate the aqueous and organic layers. 10. Carefully transfer the top aqueous layer (containing the DNA) to a new microcentrifuge tube and dispose of the lower organic layer to an appropriate waste receptacle. 11. Precipitate DNA by adding 0.7 vol of ice-cold isopropanol (350 μl) and mix gently by inverting tubes 10 to 20 times. 12. Centrifuge 10 min at 20,000 × g, 4◦ C, to pellet genomic DNA. 13. Remove isopropanol by decanting tubes, then add 500 μl of 70% ethanol and invert tubes 10 to 20 times. 14. Centrifuge 5 min at 20,000 × g, 4◦ C, to pellet clean genomic DNA. 15. Remove ethanol by decanting tubes and allow residual ethanol to evaporate for 5 to 10 min before resuspending DNA in 100 μl of TE buffer. 16. Incubate 30 min at 55◦ C or overnight at 4◦ C to dissolve DNA pellet. 17. Determine the DNA concentration using a spectrophotometer (Gallagher and Desjardins, 2006) and dilute to 100 ng/μl with TE buffer.

Perform genotyping PCR 18. Set up PCR reactions for each transgene as follows (prepare a mastermix of the following components without DNA by multiplying the volumes by the number of samples plus one for pipetting errors): 22 μl 1.1× ReddyMix 1 μl forward primer (10 μM) 1 μl reverse primer (10 μM) 1 μl DNA sample (100 ng/μl) Primer sequences for R26-LSL-SB11,T2/Onc, and Cre can be found in Table 1.

19. Pipet 24 μl of PCR master mix into 0.5-ml thin-walled PCR tubes and add 1 μl of diluted genomic DNA (100 ng/ μl) per tube. Run positive and negative genomic DNA controls in parallel. Loading genomic DNA should be done carefully using filtered tips and new tips should be used between samples to prevent cross contamination. Positive and negative control— depending on what transposon line, TSP-cre, and predisposing background are used (this can be highly variable)—should also be run in parallel to ensure quality genotyping. Additionally, always prepare a master mix when possible to avoid variation between reactions.

20. Amplify PCR amplicons using a thermal cycler with the following program: 1 cycle: 35 cycles: Sleeping Beauty-Based Somatic Transposon Mutagenesis in the Mouse

1 cycle:

2 min 25 sec 35 sec 1 min 5 min hold

95◦ C 95◦ C 55◦ C 72◦ C 72◦ C 4◦ C

For R26-LSL-SB11, T2/Onc, and Cre, the expected amplicon size for R26-LSL-SB11 is 420 bp and wild-type R26 is 266 bp, T2/Onc is 250 bp, and Cre is 500 bp.

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Animal 8

Animal 7

Animal 6

Animal 5

Animal 4

Animal 3

Animal 2

Animal 1

( ) Control

H2O ( ) Control Cre recombinase

R26-LSL-SB11

500 bp

420 bp WT 266 bp LSL-SB11

T2/Onc 250 bp

Figure 2 An example of PCR genotyping results for Cre recombinase, T2/Onc, and R26-LSL-SB11. Cre genotyping primers should produce a product of 482 bp. The R26-LSL-SB11 genotyping PCR utilizes a three-primer PCR to amplify both the wild-type (WT) R26 and the knock-in R26-LSL-SB11 alleles in a single reaction with wild type producing a 420-bp product and the SB11 knock-in producing a 266-bp product. T2/Onc genotyping primers should produce a product of 264 bp.

Analyze PCR results 21. Prepare a 1% agarose gel using 1× TAE and add 4 μl ethidium bromide per 100 ml agarose after boiling; use 1× TAE as the running buffer (Voytas, 2000). 22. Load PCR products directly onto the 1% gel as the 1.1× Reddymix already contains loading dye. On either end of the PCR products, load an appropriately sized DNA ladder to ensure the amplicons are of the correct size (see Fig. 2). 23. Using the appropriate power supply, run the gel at 140 V for ∼30 to 40 min, or until multiplex PCR bands are resolved sufficiently. 24. Capture an image of the gel for analysis using Polaroid film or another digital device. 25. Analyze PCR results carefully to ensure Mendelian inheritance of all transgenes has occurred. If genotyping indicates that there is non-Mendelian inheritance of any allele, it could mean that some combination of the transgenes is embryonically lethal. This could require the selection of a new Cre knock-in transgenic depending on the situation.

VERIFYING TRANSPOSASE EXPRESSION AND TRANSPOSON MOBILIZATION

BASIC PROTOCOL 2

There are two quintessential assays to determine if transposon mutagenesis is occurring in the tissue and/or cell type of interest: SB immunohistochemistry (IHC) and the transposon PCR excision assay (see Alternate Protocol). The first ensures that the SB transposase

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protein is being expressed in the cell of interest and the latter indicates that mutagenic transposons are being mobilized from the T2/Onc concatemer in the tissue of interest.

Materials Prepared slides of tissue of interest cut from paraffin blocks Citrosolv (Fisher Scientific) Ethanol (Sigma-Aldrich) Phosphate buffered saline, pH 7.5 (Fisher Scientific) Unmasking solution (Vector Laboratories) 3% hydrogen peroxide (H2 O2 ) diluted in water (Sigma-Aldrich) M.O.M. mouse Ig blocking reagent (Vector Laboratories) 1× PBST (1× PBS with 0.1% Tween 20) Anti-SB11 antibody (mouse monoclonal clone 324622) (R&D Systems) VectaStain Elite ABC reagent (Vector Laboratories) Diaminobenzidene (DAB) substrate kit (Vector Laboratories) Harris hematoxylin (Fisher Scientific) Permount (Fisher Scientific) 1-liter beakers Microwave ImmunoEdge Pen (Vector Laboratories) Humidity chamber (see recipes) 24 × 50 no. 1.5 coverslips (Thermo Scientific) Microscope Deparaffinize and prepare slides 1. Deparaffinize slides by incubating slides two times in Citrisolv for 5 min, each time. 2. Re-hydrate slides by sequential incubation in an ethanol gradient as follows: 5 min in 100% ethanol; 5 min in 100% ethanol; 5 min in 95% ethanol; 5 min in 90% ethanol; 5 min in 70% ethanol; 5 min in water; 5 min in PBS. It is acceptable to keep in PBS up to 4 hr if necessary.

3. In a 1-liter beaker, submerge slides in unmasking solution and boil slides for 30 min using a microwave set to high (this step removes cross-linking induced by formalin fixation, which frees up protein epitopes to become bound by antibody). It is extremely important to ensure that the solution level does not fall below the slides during the boiling procedure so as to prevent destruction of tissues on the slides.

4. Allow slides to cool to room temperature by placing beaker on ice. Do not allow the tissues on the slides to dry after this step. 5. Rinse slides in PBS for 5 min.

Block endogenous peroxidases 6. Block all endogenous peroxidases in the tissue by placing slides in 3% hydrogen peroxide for 10 min. 7. Rinse slides two times in PBS for 5 min, each time. 8. Using an ImmunoEdge pen, draw a boundary around the tissues of each of the slides being stained and place slides in a humidity chamber. Sleeping Beauty-Based Somatic Transposon Mutagenesis in the Mouse

From this step on, PBST is used in place of PBS to prevent the slides from drying out.

9. Place 100 μl of diluted M.O.M. Ig blocking reagent on the slide and incubate 1 hr at room temperature.

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The hydrophobic immunoedge boundary will prevent the blocking liquid from spreading across the entire slide and will contain the blocking reagent on the circled tissue only.

10. Wash slides two times in PBST for 2 min, each time.

Add primary antibody 11. Dilute primary anti-SB11 antibody in M.O.M. diluent (1:100 or 2.5 μg/ml) and place ∼100 μl of primary antibody solution on tissue sections and incubate overnight at 4◦ C or 4 hr at room temperature in a humidity chamber. Perform the necessary no-primary control for each slide—use M.O.M. diluent without primary antibody. This controls for any non-specific binding of the secondary antibody.

12. After overnight incubation, prepare VectaStain Elite ABC avidin/biotin-HRP complex reagent by adding two drops of reagent A to 2.5 ml of PBST, mix, then add two drops of reagent B and mix, allow reaction to occur 30 min before using. 13. Rinse slides two times in PBST for 2 min, each time. 14. Place 100 μl of diluted M.O.M. biotinylated anti-mouse IgG (10 μl stock into 2.5 ml of M.O.M. diluent) onto tissue sections and incubate 30 min at room temperature in humidity chamber. 15. Rinse slides two times in PBST for 5 min, each time.

Stain samples 16. Place 100 μl VectaStain Elite ABC avidin/biotin-HRP complex reagent on tissue sections and incubate 5 min. 17. Wash two times in PBST for 5 min, each time. 18. Prepare fresh DAB substrate by adding one drop buffer, two drops DAB, and one drop hydrogen peroxide in 2.5 ml water, vortexing the solution after adding each reagent. 19. Place ∼100 μl DAB substrate on tissue sections and incubate 1 to 3 min and watch for the development of brown staining. 20. Rinse slides 5 min in water.

Tumor 1

No primary control

SB antibody stained

(40 )

(40 )

Figure 3 Shown are example photomicrographs of immunohistochemical results for the SB transposase protein counterstained with hematoxylin. Staining of tumor cells expressing SB show robust brown horseradish peroxidase staining that is most pronounced in the nucleus where the protein is localized. Negative control staining lacking primary antibody should be devoid of brown horseradish peroxidase staining but still show counterstaining with hematoxylin.

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21. Dip slides in Harris hematoxylin 15 to 20 times.

Wash and mount slides 22. Gently rinse slides in a beaker under tap water for ∼3 min or until the blue stain no longer runs off the slides. 23. Dehydrate tissue sections by sequential incubation in an ethanol gradient as follows: 5 min in 70% ethanol; 5 min in 90% ethanol; 5 min in 95% ethanol; 5 min in 100% ethanol; 5 min in 100% ethanol; 5 min in CitriSolv; and 5 min in CitriSolv. 24. Mount slides in Permount and apply glass coverslip. Be careful not to produce bubbles when mounting slides.

25. View slides under microscope. Positive staining is brown and nuclear, as SB transposase is a nuclear protein. See example of positive SB staining in Figure 3. ALTERNATE PROTOCOL

TRANSPOSON PCR EXCISION As discussed in the introduction for Basic Protocol 2, this protocol is used to determine whether mutagenic transposons are being mobilized from the T2/Onc concatemer in the tissue of interest.

Materials 2× ReddyMix (Thermo Scientific) Primers (Table 1) DNA samples DNase/RNase-free water (Qiagen) Genomic DNA (negative and positive controls) 1% agarose gel in 1× TAE buffer 1× TAE (from 50× stock; see recipe) 100-bp Quanti-marker (BioExpress) Ethidium bromide (10 mg/ml) (Sigma-Aldrich) 0.5-ml thin-walled PCR tubes Thermal cycler Gel electrophoresis apparatus and power source UV light box Gel photography equipment Additional reagents and equipment for DNA extraction (see Basic Protocol 1) 1. Set up PCR reactions using 2× ReddyMix for each sample as follows:

25 μl 2× ReddyMix 1 μl forward primer (10 μM) 1 μl reverse primer (10 μM) 2.5 μl DNA sample (20 ng/μl) 20.5 μl water. Primer sequences are listed in Table 1.

Sleeping Beauty-Based Somatic Transposon Mutagenesis in the Mouse

Be sure to run positive and negative excision controls. It is advisable to analyze many tissues from at least two independent experimental and control class animals to determine all of the tissues undergoing mutagenesis. DNA can be extracted using the same phenol/chloroform extraction method used for tail clippings (see Basic Protocol 1).

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2. Amplify PCR products using the following program on a thermal cycler: 1 cycle: 35 cycles: 1 cycle:

95◦ C 95◦ C 55◦ C 72◦ C 72◦ C 4◦ C

2 min 25 sec 35 sec 90 sec 5 min Hold

The PCR excision primers found in Table 1 are specific to T2/Onc mice. If another transposon line is used, then different excision primers must be designed.

3. Prepare a 1% agarose gel using 1× TAE and add 4 μl ethidium bromide per 100 ml after boiling; use 1× TAE as the running buffer (Voytas, 2000). 4. Directly load PCR products onto the 1% gel as the 2× Reddymix already contains loading dye. On either end of the PCR products, load an appropriately sized DNA ladder to ensure the PCR products are of the correct size. 5. Using an appropriate power supply, run the gel at 140 V for ∼30 to 40 min, or until PCR excision bands are resolved sufficiently. 6. Capture an image of the gel for analysis using Polaroid film or another digital device. 7. Analyze the PCR results to ensure that the smaller 225-bp products are only apparent in the tissue of interest and that all other tissue types produce the excision negative 2.2-kb PCR product. See Figure 4 for an example excision assay.

( ) excision control

( ) excision control

Osx. 142 tumor

Osx. 88 tumor

Osx. 58 tumor 2

Osx. 58 tumor 1

Osx. 48 tumor

Osx. 47 tumor

Osx. 47 tumor SB negative

Osx. 46 tumor

Note that if there is excision occurring in undesired tissues, background tumors or phenotypes may develop, as mutagenesis will be occurring in these tissue types.

2.2 Kb

225 bp

Figure 4 An example of a PCR excision assay results from a panel of transposon mutagenesis induced tumors. Tumors positive for transposon mobilization should produce a 225-bp product. If no transposition has occurred, then a 2.2-kb product should be observed, as shown for tumor 2, which was negative for the SB transposase and developed as a background tumor in this experiment. Some tumors may be composed of a mix of cells positive and negative for transposition and will thus produce both the 225-bp and 2.2-kb products, as shown in tumor 5 and 6.

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BASIC PROTOCOL 3

IDENTIFICATION OF TRANSPOSON INSERTION SITES BY LM-PCR AND HIGH-THROUGHPUT SEQUENCING Once the phenotype of interest has been identified, samples can be collected and examined for transposon insertion sites by ligation-mediated polymerase chain reaction (LM-PCR). See overview in Figure 5 for molecular details. The method outlined below will allow for cloning of each transposon insertion site from both the left and right side of the transposon to ensure that the majority of insertions are successfully mapped to the mouse genome.

Materials Genomic DNA (see Basic Protocol 1) TE buffer (see recipe) NlaIII (New England Biolabs) BfaI (New England Biolabs) Primers (Table 1) 5 M NaCl T4 DNA ligase and 10× buffer (New England Biolabs) BamHI (New England Biolabs) Buffer no. 3 (New England Biolabs) 100× BSA 2× ReddyMix (Thermo Scientific) 10× buffer with 10 mM MgCl2 (Roche Scientific) 25 mM dNTPs (Roche Scientific) Fast-Start Taq polymerase (Roche Scientific) 2% agarose gel in 1× TAE buffer 1× TAE (from 50× stock; see recipe) 6× DNA loading buffer (see recipe) Ethidium bromide (10 mg/ml) (Sigma-Aldrich) 96-well PCR plates 37◦ C incubator 80◦ and 95◦ C heating blocks Qiagen MinElute 96 UF plates (Qiagen) Orbital shaker Gel electrophoresis apparatus and power source UV light box Gel photography equipment Spectrophotometer 1.5-ml microcentrifuge tubes Additional reagents and equipment for DNA isolation (see Basic Protocol 1) Enzyme digest DNA 1. Isolate genomic DNA from the cancer or phenotype of interest using the same methodology as isolating DNA from tails outlined in Basic Protocol 1. 2. Dilute genomic DNA to 10 ng/μl using TE buffer. 3. Set-up restriction enzyme digests of each sample with BfaI and Nlalll separately in 96-well PCR plates as follows: Sleeping Beauty-Based Somatic Transposon Mutagenesis in the Mouse

20 μl genomic DNA (10 ng/μl) 5 μl buffer no. 4 (10×) 0.5 μl BSA (100×) 1 μl NlaIII or BfaI 23.5 μl water

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TA dinucleotide

transposon (IR/DR)

genomic DNA

restriction enzyme digest

anneal linker

1° PCR

2° PCR

purify amplicons and sequence

barcode

IR/DR sequence

genomic sequence

illumina specific sequence

linker sequence illumina specific sequence

Figure 5 Flowchart outlining the molecular details at each step of the ligation-mediated PCR (LM-PCR) procedure leading to the final products that contain all the necessary elements for high-throughput Illumina sequencing to identify the genomic site of transposon integrations.

Always prepare a master mix when possible to avoid variation between reactions.

4. Incubate restriction enzyme digests overnight at 37◦ C. 5. After overnight digest, heat inactivate digests 20 min at 80◦ C.

Prepare linker primers 6. At this point, prepare linker primers for ligation to the digested genomic DNA. Mix 50 μl of both the sense (+) and anti-sense (–) linker for the BfaI and NlaIII linkers (100 μM stocks), add 2 μl of 5 M NaCl, and heat to 95◦ C for 5 min. Allow the reactions to cool to room temperature to allow the primers to anneal appropriately (∼2 hr). 7. Set up the linker ligation reaction of digested genomic DNA using the appropriate linker as follows:

10 μl restriction enzyme digested genomic DNA 1 μl left or right linker 2 μl T4 DNA ligase buffer (10×) 1 μl T4 DNA ligase 6 μl water Allow ligation to proceed overnight at 16◦ C.

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Purify ligation 8. Purify ligated DNA using Qiagen MinElute 96 UF plates following manufacturer’s instructions. Resuspend in 40 μl of water by shaking sealed/secured plate for 3 min on an orbital shaker. 9. At this point, remove the transposon concatemer with any remaining transposons by digesting with BamHI. Incubate overnight at 37◦ C after setting up the restriction enzyme digest as follows:

40 μl restriction enzyme–digested/linker-ligated/purified genomic DNA 5 μl NEB buffer no. 3 (10×) 0.5 μl BSA (100×) 1 μl BamHI 3.5 μl water This step assumes that a T2/Onc transgenic line is being used as the transposons are directly flanked by BamHI sites on either side in the concatemer. If another line was used as the source of mutagenic transposons, then a different restriction enzyme may be required to remove the residual transposon concatemer. If this step is neglected, many sequenced insertions will map to the concatemer.

10. Purify DNA using Qiagen MinElute 96 UF plates following the manufacturer’s instructions. Resuspend in 20 μl water by shaking sealed/secured plate for 3 min on an orbital shaker.

Perform primary PCR 11. Use 1 μl of the resuspended template DNA for the primary PCR as follows: 25 μl 2× ReddyMix 1 μl Splink left or right (10 μM) 1 μl Splink linker (10 μM) 1 μl DNA sample 22 μl water Use a thermal cycler with the following program: 1 cycle: 35 cycles:

1 cycle:

2 min 30 sec 30 sec 1.5 min 5 min Hold

95◦ C 95◦ C 60◦ C 72◦ C 72◦ C 4◦ C

Perform secondary PCR 12. Dilute 3 μl of the primary PCR into 222 μl water (1:75 dilution) and use 1 μl of the diluted DNA for the secondary PCR as follows:

Sleeping Beauty-Based Somatic Transposon Mutagenesis in the Mouse

1 μl primary PCR diluted 1:75 5 μl 10× buffer with MgCl2 0.4 μl dNTPs (25 mM) 1 μl nested left or right (barcoded) (10 μM) 1 μl nested linker (10 μM) 0.5 μl Fast-Start Taq polymerase 41.1 μl water

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Perform PCR on a thermal cycler with the following program: 1 cycle: 35 cycles: 1 cycle:

2 min 30 sec 45 sec 90 sec 5 min Hold

94◦ C 94◦ C 53◦ C 72◦ C 72◦ C 4◦ C

Secondary PCR, or nested PCR, must be performed for two main reasons. First, it further enriches for transposon insertion sites as there may be non-specific amplicons generated in the primary PCR. Furthermore, the secondary PCR incorporates the nucleotide barcode for high-throughput sequencing using the Illumina platform. Barcodes can be designed using various methods such as BARCRAWL (Frank, 2009). It is advisable to consult a bioinformaticist on this subject.

13. Prepare a 2% agarose gel using 1× TAE and add 4 μl ethidium bromide per 100 ml after boiling; use 1× TAE as the running buffer (Voytas, 2000). Load 10 μl of PCR product mixed with 2 μl of 6× loading buffer. 14. Using the appropriate power supply, run the gel at 140 V for ∼30 to 40 min. 15. Capture an image of the gel for analysis using Polaroid film or another digital device. See Figure 6 for an example of LM-PCR products. No distinct bands should be present, rather a smear should be observed indicating many different sized amplicons corresponding to many unique insertion sites.

Purify DNA 16. Purify DNA from the remaining PCR reaction using Qiagen MinElute 96 UF plates following the manufacturer’s instructions. Wash DNA one time by adding 50 μl water to each well after DNA has been bound to the membrane and the plate is still on the vacuum manifold. Resuspend in 30 μl water by shaking sealed/secured plate for 3 min.

Sample 8

Sample 7

Sample 6

Sample 5

Sample 4

Sample 3

Sample 2

Sample 1

Bl/6 wild-type DNA

H2O ( ) control

17. Determine the DNA concentration of each sample using a spectrophotometer (Gallagher and Desjardins, 2006) and dilute to 25 ng/μl. Combine equal amounts of DNA (usually ∼100 ng) from each sample into a single 1.5-ml microcentrifuge tube and send out for high-throughput Illumina sequencing.

Figure 6 An example of ligation-mediated PCR (LM-PCR) results from a panel of tumors induced by transposon mutagenesis. LM-PCR products should appear as smears as they contain many different sized products that correspond to many different amplified transposon-genomic DNA junction products. Wild-type mouse DNA and water should not produce PCR products of any kind. Current Protocols in Mouse Biology

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REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps.

DNA loading buffer, 6× 0.25% (w/v) bromphenol blue 0.25% (w/v) xylene cyanol 30% (v/v) glycerol in water Store indefinitely at 4◦ C Humidity chamber A humidity chamber is any enclosed box with wet paper towels, e.g., a plastic box containing four cut-off pipets, which are used to raise the slides off of the bottom of the chamber. A layer of paper towels lay on the top and bottom of the pipets. Water is poured into the chamber so some small puddles form on top of the paper towels.

Proteinase K 10 mg/ml proteinase K in water Dispense into 1-m aliquots and store up to 1 year at −20◦ C SDS lysis buffer 10 mM Tris 1 mM EDTA 1× SSC 1% SDS Store up to 3 years at room temperature TAE, 50× 242.2 g Tris base 100 ml 0.5 M EDTA, pH 8 57.1 ml glacial acetic acid Bring up to 1 liter with water Store up to 3 years at room temperature TE buffer 100 ml 1 M Tris·Cl, pH 7.5 20 ml 500 mM EDTA, pH 8.0 Bring up to 1 liter with water Store up to 3 years at room temperature

COMMENTARY Background Information

Sleeping Beauty-Based Somatic Transposon Mutagenesis in the Mouse

Forward genetic screens In the new age of human cancer genomics, there has been some thought that forward genetic screens in model organisms could be replaced with whole genome sequencing, array comparative genome hybridization (aCGH), and other genome-wide analysis technologies (Mohr et al., 2002; Kallioniemi, 2008). However, somatic mutagenesis is still an extremely powerful technique to identify genes involved

with a specific phenotype and in some instances can identify genes that genome-wide technologies could never detect in human samples. For instance, there is still not an efficient genome-wide technology to identify genes that are selected for epigenetic silencing or overexpression during tumor development in an in vivo setting. There are technologies to identify if a particular gene is methlyated or modified in some fashion, but with hundreds of genes undergoing silencing in any given

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cell it is nearly impossible to identify the silenced genes that are involved with the phenotype without some sort of functional validation. This is just one of many types of gene modulation that forward genetic screens can identify that whole genome technologies cannot currently identify. Additionally, unannotated genes or miRNA are out of the range of most whole genome analysis, whereas forward genetic screens are non-biased and are able to identify even unannotated genes or functional elements such as miRNA. Lastly, forward genetic screens can also lead to the elucidation of gene functions that have not been previously known, which is necessary in the field of biology as there are so many understudied genes (Huss et al., 2010). Forward genetic screens have been utilized for many years and have been performed in many different model organisms with a myriad of different mutagens. N-ethyl-N-nitrosourea (ENU) and retroviral mutagenesis has been particularly fruitful in many model organisms (Jonkers and Berns, 1996; Uren et al., 2005; Gondo, 2008). However, both of these mutagens have drawbacks that make them nonideal. For example, it can be difficult to identify the mutated gene of interest using ENU as it entails many back-crosses that are time consuming or impossible in some organisms, such as the mouse. Retroviral mutagenesis has been a powerful technique in the mouse and has lead to the identification of many genes involved in cancer. Retroviral mutagenesis is hampered by its predisposition to insert in actively transcribed genes and seems biased toward activation of proto-oncogenes. Retroviral infection is restricted in the cell type that it can efficiently infect, making it useless for modeling many forms of cancer, being primarily useful for studies on leukemia/lymphoma and mammary carcinoma (Mikkers and Berns, 2003). An ideal mutagen is one that randomly mutates the genome, can inactivate or over express genes, is easily trackable, and that is amenable for use in many cell/tissue types. Sleeping Beauty is a mutagen that has been shown to fulfill all of these attributes of an ideal mutagen.

Critical Parameters Genotyping Proper genotyping is arguably the most important aspect of successfully performing an SB forward genetic screen. If there are mistakes in the genotyping, it can lead to excessive

mouse breeding and lost time. The best way to avoid incorrect genotyping is to always run the appropriate controls, monitor the inheritance of alleles, and be sure that the PCR amplicons are of the correct size. Tissue collection and storage The proper collection and storage of tissue samples is a critical step in any experiment and especially in an SB forward genetic screen. Have a plan in place prior to sacrificing any mice. Upon necropsy, any tissue of interest should be processed to attain the most information possible. Ideally, the sample should be snap frozen in liquid nitrogen and stored at −80◦ C for later isolation of DNA and protein. Another portion of the sample should be formalin fixed or embedded in optimum cutting temperature (OCT) compound for later use in IHC. Lastly, if there is enough sample, a portion should be placed in an RNA preservation reagent, such as RNAlater (Sigma-Aldrich), and stored at −80◦ C. Additionally, be sure to collect and properly store any tissue that may be of interest in the future even if it is not directly the phenotype of interest. For example, in cancer gene identification, it would be ideal to collect the primary tumor and also any tissue that may harbor metastases, such as the liver, lung, and spleen. These grossly normal tissues can later be sectioned to identify micrometastases and laser capture microdissection performed to identify transposon insertion sites, as described by Rahrmann et al. (2009).

Troubleshooting Colony generation In general, generating a colony should be a straightforward task. However, there are some things that can be done to ensure success. Always use mice of prime breeding age, 8- to 12-weeks of age. Mice should produce litters within 3 to 4 weeks and pregnancy is evident by a rapid increase in weight and size of pregnant females. A deficiency of pregnant females could be due to behavioral or physical infertility. Harem breeding of male mice with two to three females can help distinguish which animals may not be breeding and new matings can then be performed with other animals. Vigilant documentation of the number of offspring can also help to identify decreases in fertility of breeding pairs. If the expected number or ratio of transgenes is non-Mendelian based on the genotypes of the breeders, then the genotyping of the breeders should be repeated.

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Genotyping Mis-genotyped mice can occur from many different problems. As a first precaution always run positive and negative control DNA with every set of genotyping PCR reactions. If there are no bands present in any of the PCR reactions, but there is a clear DNA ladder, it could be due to many factors, such as too little DNA or degraded DNA, inappropriate PCR program, or expired or degraded reagents. Requantify genomic DNA and check the quality on an agarose gel, check the PCR program, and if the problem persists, replace all reagents and repeat the genotyping PCR. On the other hand, if all of the PCR reactions exhibit bands, then a reagent or piece of equipment is likely contaminated. Prepare fresh reagents from new stock and retry genotyping, if the problem still persists, the genomic DNA could be contaminated and new DNA should be extracted from the animals. Immunohistochemistry (IHC) As there are many steps involved in IHC there are also many potential problems that can occur. Ensuring that all reagents are fresh, including ethanol gradients, citrisolv, antibodies, and hematoxylin, will provide the best opportunity for success. Always be sure to stain negative and positive control tissues as well as the no primary antibody controls for every tissue that is being stained for SB protein expression. If there is no positive staining on any slides, check that all steps were performed and in the correct order. If there is too much background staining, it may be advisable to decrease the time that the DAB substrate is applied to the slides. If the hematoxylin staining is too robust, then the number of dips can be reduced to as few as five, especially if the hematoxylin is fresh.

Sleeping Beauty-Based Somatic Transposon Mutagenesis in the Mouse

PCR excision assay Many of the troubleshooting tips for the PCR excision assay are the same as for genotyping PCR troubleshooting, see above. In some cases, if the diluted DNA concentrations are not accurate or if the thermal cycler is not calibrated precisely, there may be faint but visible non-specific bands; these bands can generally be ignored if the bands of interest are robust and of the appropriate sizes. However, in most cases, only the upper and lower bands are observed if the protocol is performed correctly. LM-PCR Due to the many steps and complexities involved with LM-PCR, there are also many

places to make potential mistakes. The best way to ensure success when performing LMPCR is to include negative and positive controls, pay close attention to what is being done at each step, and be sure not to switch the left and right side cloning reactions at any point in the process. If proper controls are not performed throughout, it is not possible to tell if the final results are correct. It is of utmost importance to assemble each reaction with the appropriate amounts and types of reagents to ensure successful cloning of insertion sites. If the left and right side reactions are switched at any point after the initial restriction enzyme digest, then the procedure will fail as each side has specific linkers and PCR primers.

Anticipated Results Colony generation Once established, breeding pairs should produce litters every 19 to 21 days. The litter sizes should range from five to ten, depending on the age of the breeding pair and genetic background. All transgenic alleles should be inherited in a Mendelian fashion. Genomic DNA isolation Genomic DNA extracted from tails at weaning using phenol/chloroform should yield high-quality DNA with a 260/280 ratio varying from 1.7 to 2.0 and have a concentration between 50 ng/μl and 500 ng/μl. DNA isolated from tissue should also be of high quality but may have a much higher or lower DNA concentration depending on the cellularity of the tissue from which DNA is being extracted and also the size of the starting material. Immunohistochemistry (IHC) Positive staining for SB proteins should appear as dark punctate staining in the nucleus, as this is where the transposase functions. There should not be diffuse staining or high background if the protocol outlined above is followed properly. PCR excision assay The positive control should have only the excision band at 225 bp and the negative control should have only the upper 2.2-kb excision band. Samples under investigation could have either band and in some cases may have both bands, indicating that there was a mixture of transposition positive and negative cells in the tissue from which the DNA was isolated or there is a low rate of transposition and the concatemer of mutagenic transposons has not been depleted.

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LM-PCR In most cases, LM-PCR should yield thousands of unique transposon insertion sites. In the authors’ experience, one lane of an Illumina sequencing run should yield between 20 and 50 million sequence reads. These data need to be filtered and distilled by a knowledgeable bioinformaticist to acquire a meaningful catalogue of non-redundant insertion sites from which a list of common insertion sites can be calculated to identify candidate genes of the phenotype of interest.

Time Considerations Colony generation The time to generate a cohort of animals depends on many factors including approval of the institution to perform mouse experiments, obtaining all required transgenics, and quarantine time before breeding can begin. Once all of these tasks are finished, it should take between 6 and 8 weeks to begin a small colony of breeding pairs to produce experimental and control animals. The more transgenes needed per animal and the total number of experimental and control animals needed both increase the time to acquire a complete cohort of animals necessary to perform a successful forward genetic screen. It may take 6 months to 1 year to finish breeding a cohort of animals. Genomic DNA isolation The time necessary to perform genomic DNA isolation is dependent on the number of samples to be isolated. It should take ∼1.5 to 2 hr to extract DNA from ten samples. Genotyping Genotyping should take ∼15 min for all PCR components to thaw on ice before setting up PCR reactions. For ten samples, with controls, it should take ∼15 to 20 min to set up the reactions. PCR times vary on the number and time of each cycle in the PCR program but are generally <3 hr. Resolving the PCR amplicons by gel electrophoresis should take an additional 30 to 60 min. Immunohistochemistry (IHC) IHC is typically a 2-day procedure and requires ∼4 hr per day, so in total it requires ∼8 hr of time assuming <24 slides are processed at once. If more slides are involved, then the protocol could easily take 2 full days to complete. However, there are a lot of incubation steps that allow for multiple procedures to be carried out in tandem.

PCR excision assay The PCR excision assay should take a similar amount of time as genotyping. LM-PCR Since LM-PCR is a multi-day procedure, it can be a long procedure, especially if there is a large number of samples being processed. Assuming there are <20 samples, the primary restriction digests should take ∼30 min to set up. After the overnight digestion, heat inactivation requires 20 min and the linker annealing preparation takes ∼2.5 hr. Preparing the ligation reaction with the annealed linkers should take ∼30 min. After the overnight ligation, cleaning up the ligated fragments with the mini-elute plates takes ∼30 min including the resuspending of DNA. The secondary digest should also take ∼30 min to set up. After the secondary overnight digest, it should take another 30 min to clean up the DNA. The primary PCR should take ∼30 min to set up and the PCR cycle is ∼2 hr. Secondary PCR takes a similar amount of time with 10 min added to dilute the primary PCR in water. Cleaning up the DNA from the secondary PCR takes ∼30 min and running a small portion of the secondary PCR on an agarose gel will take 30 to 60 min. Overall, the process should take ∼10 to 12 hr spread out over 4 days. Up to 400 samples can be processed at a time over the 4 days.

Conflict of Interest Statement David A. Largaespada is a consultant to and co-founder of a biotechnology company called Discovery Genomics, Inc. (DGI) which is pursuing transposon technology for gene therapy. However, none of the methods or work described in this manuscript are related to DGI goals and DGI resources or personnel were not used.

Literature Cited Collier, L.S., Carlson, C.M., Ravimohan, S., Dupuy, A.J., and Largaespada, D.A. 2005. Cancer gene discovery in solid tumours using transposon-based somatic mutagenesis in the mouse. Nature 436:272-276. Collier, L.S., Adams, D.J., Hackett, C.S., Bendzick, L.E., Akagi, K., Davies, M.N., Diers, M.D., Rodriguez, F.J., Bender, A.M., and Tieu, C. 2009. Whole-body sleeping beauty mutagenesis can cause penetrant leukemia/lymphoma and rare high-grade glioma without associated embryonic lethality.Cancer Res. 69:8429. Dupuy, A.J., Akagi, K., Largaespada, D.A., Copeland, N.G., and Jenkins, N.A. 2005. Mammalian mutagenesis using a highly mobile

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somatic sleeping beauty transposon system. Nature 436:221-226. Dupuy, A.J., Rogers, L.M., Kim, J., Nannapaneni, K., Starr, T.K., Liu, P., Largaespada, D.A., Scheetz, T.E., Jenkins, N.A., and Copeland, N.G. 2009. A modified sleeping beauty transposon system that can be used to model a wide variety of human cancers in mice. Cancer Res. 69:8150. Frank, D.N. 2009. BARCRAWL and BARTAB: Software tools for the design and implementation of barcoded primers for highly multiplexed DNA sequencing. BMC Bioinformatics 10:362. Gallagher, S.R. and Desjardins, P.R. 2006. Quantitation of DNA and RNA with absorption and fluorescence spectroscopy. Curr. Protoc. Mol. Biol. 76:A.3D.1-A.3D.21. Gondo, Y. 2008. Trends in large-scale mouse mutagenesis: From genetics to functional genomics. Nat. Rev. Genet. 9:803-810. Huss, J.W., Lindenbaum, P., Martone, M., Roberts, D., Pizarro, A., Valafar, F., Hogenesch, J.B., and Su, A.I. 2010. The gene wiki: Community intelligence applied to human gene annotation. Nucleic Acids Res. 38:D633. Ivics, Z., Hackett, P.B., Plasterk, R.H., and Izsv´ak, Z. 1997. Molecular reconstruction of sleeping beauty, a Tc1-like transposon from fish, and its transposition in human cells. Cell 91:501-510. Jonkers, J. and Berns, A. 1996. Retroviral insertional mutagenesis as a strategy to identify cancer genes. Biochim Biophys Acta 1287:29-57. Kallioniemi, A. 2008. CGH microarrays and cancer. Curr. Opin. Biotechnol. 19:36-40. Keng, V.W., Villanueva, A., Chiang, D.Y., Dupuy, A.J., Ryan, B.J., Matise, I., Silverstein, K.A.T., Sarver, A., Starr, T.K., and Akagi, K. 2009. A conditional transposon-based insertional mutagenesis screen for genes associated with mouse hepatocellular carcinoma. Nat. Biotechnol. 27:264-274.

Mikkers, H. and Berns, A. 2003. Retroviral insertional mutagenesis: Tagging cancer pathways. Adv. Cancer Res. 88:53-99. Mohr, S., Leikauf, G.D., Keith, G., and Rihn, B.H. 2002. Microarrays as cancer keys: An array of possibilities. J. Clin. Oncol. 20:3165. Rad, R., Rad, L., Wang, W., Cadinanos, J., Vassiliou, G., Rice, S., Campos, L.S., Yusa, K., Banerjee, R., and Li, M.A. 2010. PiggyBac transposon mutagenesis: A tool for cancer gene discovery in mice. Science 330:1104. Rahrmann, E.P., Collier, L.S., Knutson, T.P., Doyal, M.E., Kuslak, S.L., Green, L.E., Malinowski, R.L., Roethe, L., Akagi, K., and Waknitz, M. 2009. Identification of PDE4D as a proliferation promoting factor in prostate cancer using a sleeping beauty transposon-based somatic mutagenesis screen. Cancer Res. 69:4388. Soriano, P. 1999. Generalized lacZ expression with the ROSA26 cre reporter strain. Nat. Genet. 21:70-71. Starr, T.K., Allaei, R., Silverstein, K.A.T., Staggs, R.A., Sarver, A.L., Bergemann, T.L., Gupta, M., O’Sullivan, M.G., Matise, I., and Dupuy, A.J. 2009. A transposon-based genetic screen in mice identifies genes altered in colorectal cancer. Science 323:1747. Starr, T.K., Scott, P.M., Marsh, B.M., Zhao, L., Than, B.L.N., O’Sullivan, M.G., Sarver, A.L., Dupuy, A.J., Largaespada, D.A., and Cormier, R.T. 2011. A sleeping beauty transposonmediated screen identifies murine susceptibility genes for adenomatous polyposis coli (apc)dependent intestinal tumorigenesis. Proc. Natl. Acad. Sci. 108:5765. Uren, A.G., Kool, J., Berns, A., and Van Lohuizen, M. 2005. Retroviral insertional mutagenesis: Past, present and future. Oncogene 24:76567672. Voytas, D. 2000. Agarose gel electrophoresis. Curr. Protoc. Mol. Biol. 51:2.5A.1-2.5A.9.

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Assessment of Circadian and Light-Entrainable Parameters in Mice Using Wheel-Running Activity Gareth T. Banks1 and Patrick M. Nolan1 1

Neurobehavioural Genetics, MRC Harwell, Harwell Science and Innovation Campus, Oxfordshire, United Kingdom

ABSTRACT In most organisms, physiological variables are regulated by an internal clock. This endogenous circadian (∼24-hr) clock enables organisms to anticipate daily environmental changes and modify behavioral and physiological functions appropriately. Processes regulated by the circadian clock include sleep-wake and locomotor activity, core body temperature, metabolism, water/food intake, and available hormone levels. At the core of the mammalian circadian system are molecular oscillations within the hypothalamic suprachiasmatic nucleus. These oscillations are modifiable by signals from the environment (so called zeitgebers or time-givers) and, once integrated within the suprachiasmatic nucleus, are conveyed to remote neural circuits where output rhythms are regulated. Disrupting any of a number of neural processes can affect how rhythms are generated and relayed to the periphery and disturbances in circadian/entrainment parameters are associated with numerous human conditions. These non-invasive protocols can be used to determine whether circadian/entrainment parameters are affected in mouse mutants or treatment groups. C 2011 by John Wiley & Sons, Inc. Curr. Protoc. Mouse Biol. 1:369-381  Keywords: circadian r light entrainment r period r phase r amplitude r constant conditions

INTRODUCTION Circadian rhythms are important biological properties of all organisms, allowing for alterations in the functionality of numerous physiological systems in anticipation of or in response to environmental change. Although rhythms are generally extremely robust, a number of genetic and pharmacological factors have been shown to interfere with their periodicity, phase, and amplitude and the physiological consequences of these disturbances are likely to affect the function of many of the body’s organs. Moreover, several neurological, neurosensory, and neurobehavioral disorders in humans have associated rhythm disturbances. To avoid invasive procedures in mice, circadian system measurements are generally assessed by measuring an easily-identifiable component of the output pathway such as locomotor (wheel-running) activity under controlled lighting conditions. These measures can be used to study the effects of a specific treatment or knockout of a specific gene or can be used as a forward genetic screen to identify novel factors influencing circadian activity. The equipment and procedures used to perform basic circadian wheel running experiments and the significance of parameters that can be measured in such investigations are described in this unit. The Basic Protocol provides information on how to conduct a broad screen, which assesses the general wheel-running activity of the animal, its ability to entrain to external environmental cues (light), and the free-running period under constant conditions (constant darkness and constant light). Support Protocol 1 is a modification of the Basic Protocol to investigate the light input pathway of the animal by measuring its Current Protocols in Mouse Biology 1: 369-381, September 2011 Published online September 2011 in Wiley Online Library (wileyonlinelibrary.com). DOI: 10.1002/9780470942390.mo110123 C 2011 John Wiley & Sons, Inc. Copyright 

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responsiveness to discreet light pulses. Support Protocol 2 tests the ability of an animal to entrain under different cycling conditions.

STRATEGIC PLANNING The most important consideration for wheel running monitoring of circadian activity is the light-controlled environment into which the animals can be placed. Light-tight, ventilated chambers that can house ten home cages (see Fig. 1) are used here, but any chamber into which the wheel-running cage can be placed can be used provided it does not allow light leakage from the outside. The doors, vents, and any holes for electronic cable access should be bordered with felt or rubber to maintain a light-tight seal. Chambers must also contain an internal light source (see Critical Parameters) that can be controlled by an external timer. Wherever possible, the internal walls of the chambers should be matte black. This prevents reflection artifacts that otherwise give inconstant illumination across the chamber. In addition, this also reduces the levels of anxiety of the mice that can lead to increased variability of results. If possible, it is advisable to include a light monitor in the circadian chamber. This allows the lighting conditions to be checked without the need to open the chamber and disturb the mice. To test light leakage into the chamber, photosensitive paper can be placed into the chamber, the chamber is then sealed (with the chambers internal light left off) and the lights in the room left on for 24 hr. After 24 hr, the photosensitive paper is developed to ascertain any light leakage.

Figure 1 Typical set-up for recording wheel-running activity in mice showing light-controlled circadian chambers and wheel running cages. (A) A full circadian chamber, open to reveal wheel running cages. (B) Close up of wheel running cages within the circadian chamber. (C) Singly housed mouse within a wheel running cage.

BASIC PROTOCOL

CIRCADIAN ACTIVITY IN LIGHT/DARK CYCLES AND CONSTANT CONDITIONS This protocol gives a measure of the circadian activity of an animal in light/dark cycles, in constant darkness, and in constant light. Many of these protocols are based on the original observations by Daan and Pittendrigh (1976a,b,c,d,e) on wheel-running behavior in rodents.

Materials

Assessment of Circadian and LightEntrainable Parameters in Mice

Mice (unless chambers being used are equipped with individually ventilated cages, IVC, all animals should be same sex; young adult mice between 8 and 20 weeks; cohorts of at least ten mice per genotype or treatment group) Cages with running wheels, bedding, but no other environmental enrichment (running wheels equipped with system to quantify number of revolutions, e.g., available from http://www.coulbourn.com; http://www.panlab.com; http://www.techniplastuk.com; http://www.tse-systems.com;

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http://www.lafayetteinstrument.com) or alternatively, use microswitches or magnets to custom make activity monitoring running wheels for any standard home cage (Jud et al., 2005; Siepka and Takahashi, 2005), individual microswitches are connected to a data-collection computer via cabling connected to data acquisition boards Circadian light-tight chambers (see Strategic Planning; light set at ∼150 lux and estimated using a lux meter; air flow, temperature, and humidity maintained according to institutional recommendations) Dedicated room for housing circadian monitoring system (a set of double doors useful to prevent light from accidentally entering circadian chambers during cage checking; cover all potential light sources within the room, e.g., power monitor lights, with light-proof tape; cover overhead lighting with light filters, e.g., Kodak no. 11, to check mice daily in darkness without using infra-red goggles) Data collection computer and software to record wheel running activity (e.g., ClockLab, http://www.actimetrics.com/ClockLab/; VitalView, http://www.minimitter.com/vitalview software.cfm; The Chronobiology Kit, http://www.query.com/chronokit/; or Med Associates SOF-860, http://www.med-associates.com/activity/wireless.htm) Data collection hardware (National Instruments, cat. no. PCI-6023E) Data analysis computer (e.g., for ClockLab, an additional license for MATLAB software, The Mathworks, is required for data analysis; upload of data to the data analysis computer is automated to occur every 2 hr) Data collection 1: Entrain mice to 12:12 hr light/dark cycles 1. Set the lighting timer in the circadian chamber to generate a cycle of 12 hr of light followed by 12 hr of darkness (e.g., set lights to turn on at 7 a.m. and turn off at 7 p.m.). Set the light cycle so that it matches that of the animal house to reduce the amount of time it will take an animal to adjust and entrain to the circadian cabinet. Use of filtered lighting in a room to observe mice can be tested in advance to determine whether it causes phase shifts in the animal’s activity (see Support Protocol 1).

2. Singly house each animal to be tested in a cage containing a circadian running wheel. Use the data collection software to assign animal IDs to each cage. The data collection computer is held in a room adjacent to the circadian monitoring room.

3. Place the cages into the circadian cabinet and (if necessary) connect the running wheels as appropriate. When all cages are connected, close the cabinet. Load further cabinets as necessary until all mice are housed. Unless IVC cages are being used, all mice in the same coffin should be of the same gender.

4. Begin computer recording of wheel running activity (refer to appropriate program instructions). Use a data acquisition board to relay wheel-revolution data to the data collection computer. 5. Record activity data from these mice for 7 to 14 days. During this time, perform daily visual checks of mice and top off food and water as necessary (in accordance with local institutional recommendations). However, during these checks, minimize disruption of mouse behavior. Change cages on the final day of the light/dark cycle and again once animals have been switched to a constant light environment (see step 8 below).

Data collection 2: Expose to constant environmental conditions 6. Following entrainment to a 12:12 hr light/dark cycle, reset the lighting timer to leave the animals in a state of constant darkness. To minimize the effect of phase changing, do not interrupt a light cycle to place the animals in constant darkness. Instead set the timer so that the lights do not turn on at their next cycle. For example, if a 12-hr

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light/dark cycle is run with lights on at 7 a.m. and off at 7 p.m., then set the timer so that the lights remain off when they are due to switch on at 7 a.m. 7. Record data from the mice in constant darkness for 10 to 14 days (as per steps 4 and 5 above). If the data collection program used does not include a light monitor within the coffin, record when the period of constant darkness begins. As per step 5 above, perform daily checks on the mice. To minimize disruptions to mouse activity, use infrared cameras or red-filtered light sources to visually inspect the mice. 8. Following activity monitoring in constant darkness, reset the lighting timer to leave the animals in a state of constant light. Change cages (as per step 5 above). Record data from mice for 10 to 14 days (as per steps 4 and 5 above). If the data collection program does not include a light monitor within the coffin, record when the period of constant light begins. As per step 5 above, perform daily checks on the mice with minimum disruption to their routines. 9. When data collection is complete, remove mice from the circadian cabinets.

Analyze data 10. Once all wheel running data has been collected, it can be analyzed using appropriate analysis software (e.g., Clocklab or equivalent). Refer to software manufacturer’s instructions for details of data analysis using specific software. However, see below for common analysis measures and where it is appropriate to apply them: a. Actogram: Circadian activity rhythms are frequently presented as a specialized activity over a time graph known as an actogram (Fig. 2A). By convention, the left-hand side of an actogram is coincident with the time when lights are switched on and this is referred to as zeitgeber time 0 (ZT0), similarly in a 12:12 light/dark cycle, ZT12 is the time when lights are switched off. In constant conditions, time is referred to as circadian time (CT), where CT12 is coincident with the onset of the animal’s active phase. Circadian hours are subsequently calculated as 1/24th of the animal’s free-running period, tau (see below for definition). In constant conditions, CT0 to CT12 is referred to as subjective day and CT12 to CT24 as subjective night. In standard actograms each horizontal line represents 1 day of activity, while vertical lines plotted within the horizontal lines show wheel-running activity within that day. Activity is generally collected in time bins (e.g., 6 min). The height of the vertical lines represents the level of activity (number of wheel

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Figure 2 Actograms. (A) Standard actogram, showing wheel running activity over 2 days in a 12:12 hr light/dark cycle followed by 2 days in constant darkness. Each day’s data is presented beneath that of the previous day. Each horizontal line represents 24 hr of activity and activity is represented by vertical bars. Actograms are shaded where lights are off. (B) Double-plotted actogram of the same data shown in A. Each horizontal line represents 48 hr of activity and each day’s data is presented both beneath and to the right of the previous day.

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revolutions) during a particular time bin. Periods of darkness are represented by shading on the graph. Actograms provide a clear visual representation of the wheel running activity of a mouse in different conditions. Gross phenotypes can easily be spotted before a full analysis needs to be performed. Briefly, an animal’s activity should be predominantly nocturnal during the light/dark phase of the protocol, with the activity onsets coincident with lights-off. A vertical line can be drawn through activity onsets during this phase. During constant darkness, activity onsets usually occur a little earlier each day whereas during constant light activity onsets are a little later each day. A conventional modification of the standard actogram is the double-plotted actogram. This aligns the same actogram such that subsequent days are plotted both to the right and below the previous one. Double plotted actograms make activity cycles easier to visualize (Fig. 2B). b. Tau (τ ): The circadian period or tau (τ) is a measure of an animal’s internal circadian clock (Fig. 3). Normally, this is entrained by external cues (zeitgebers) such as light/dark cycles. However, if an animal is placed in constant conditions (i.e., without entrainment to external cues), it will free run at a period set by its internal clock. This can be estimated by fitting a line through activity onsets on successive days. Generally, the last 10 days of a particular phase of the protocol are used to measure τ. Standard software programs generally calculate this automatically using one of a number of mathematical methods (e.g., Fourier analysis or Chi-square periodogram). The length of τ for mice in constant darkness (τDD ) is typically ∼23.5 hr; however, this varies depending on the mouse strain being tested (Shimomura et al., 2001). In constant lighting conditions, a mouse will

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Figure 3 Data generated using the Basic Protocol. (A) Actogram showing mouse wheel running data using the conditions outlined in the Basic Protocol. Actograms are shaded where lights are off. Note the activity relative to lights-off in the first part of the protocol, the period shortening in constant darkness and period lengthening in constant light. (B) Graph showing average wheel running counts in the different lighting conditions of the Basic Protocol. (C) Actogram showing splitting in the constant light phase of the screen. Actograms are shaded where lights are off.

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c.

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SUPPORT PROTOCOL 1

express a consant light (τLL ) of >24 hr. This follows the Aschoff rule that states that under constant light the circadian period of a nocturnal animal will increase (Aschoff, 1981). Period lengthening in constant light (τ LL -τ DD ): Period lengthening in constant light (τLL -τDD ) can give a measure of an animal’s ability to entrain to light as a zeitgeber. In a wild-type mouse this value will be between 1 and 3 hr. Animals with deficits in the phototransduction system (e.g., melanopsin null mouse; Panda et al., 2002) will show a reduced period lengthening relative to wild-type. Quantitation of wheel running activity: Average counts of wheel rotations give a measure of the activity of an animal and can thus indicate hypo- or hyper-activity. This can be of note when considering the links between circadian and metabolic systems (e.g., Green et al., 2007). In addition, while in the light:dark phase of the protocol, a comparison of the levels of wheel-running activity in the light (inactive phase) and the dark (active phase) can give an indication of whether a cohort displays alterations in sleep behavior. However, it should be noted that wheel running activity shows high variability, even within littermates (Church et al., 2010). This feature should be used as a primary indication only. Average duration of wheel-running bouts has also been used to study deficits in, e.g., dystrophic mouse mutants (Hara et al., 2002). Alpha: A circadian rhythm can be divided into an active phase and a rest phase. The duration of the active phase (α) can be ascertained by measuring the average time elapsed between activity onset and offset. In light/dark cycling conditions α is often defined by the light/dark cycle. These measures can be taken in constant conditions, but will differ between constant dark and constant light conditions. Amplitude: Amplitude is the difference between the peak (or trough) and the mean value of a rhythm. Rhythm amplitude is often low in mutants where τ is extremely short or extremely long. Amplitude can be calculated using the same mathematical fitting methodology as per τ (see τ above). Acrophase: The mid-point or peak in the activity phase of an animal is known as the acrophase. Phase angle of entrainment: Phase angle of entrainment is the time difference between the rhythm of the entraining zeitgeber (i.e., the onset of light) and the onset of activity. This gives a measure of how responsive an animal is to the external zeitgeber.

MASKING AND PHASE SHIFTING This modified protocol provides a measure of how the extent and phase of an animal’s activity is affected by discrete changes in light input throughout the rest-activity cycle (Fig. 4). For materials, see Basic Protocol.

Data collection 1: Perform masking during light/dark cycles 1. Place mice in circadian chambers set to a 12-hr light/dark cycle following Basic Protocol, steps 1 to 5. Record the activity of these mice for 5 days to ascertain that they have entrained to the light cycle. Assessment of Circadian and LightEntrainable Parameters in Mice

2. On day 6, expose the mice to a 3-hr long light pulse starting 2 hr into the dark phase of the cycle, ZT14 (e.g., for a 7 a.m. to 7 p.m. lights-on cycle, begin a 3-hr light pulse at 9 p.m.). Continue to record activity throughout this time. 3. Return mice to the 12-hr light/dark cycle (without the light pulse) for 5 days and again ascertain that they have entrained to the light cycle.

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A

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Figure 4 Data generated from Support Protocol 1. (A) Actogram showing responses to light and dark pulses. Actograms are shaded where lights are off. Note the acute suppression of wheel-running activity by light (masking). Light pulses can also result in sustained effects on the phase of activity. In this example, the mouse shows a delayed phase of activity for 6 days after the light pulse. (B) Actogram showing the phase delay in the onset of activity in constant darkness following a 15-min light pulse applied at CT16 (light pulse is indicated by an * on the actogram). (C) Actogram showing phase advance in the onset of activity in constant darkness following a 15-min light pulse applied at CT23 (light pulse is indicated by an * on the actogram). (D) Representation of a phase response curve. Note the time window during subjective day (CT02-CT12) where activity phase is not affected by light pulses.

4. Expose the mice to a 3-hr long dark pulse starting 4.5 hr into the light phase of the cycle, ZT04.5 (e.g., for a 7 a.m. to 7 p.m. lights on cycle, begin a 3-hr dark pulse starting at 11:30 a.m.). Continue to record activity throughout this time. 5. Return the mice to the 12-hr light/dark cycle (without the dark pulse) for 5 days.

Data collection 2: Perform phase shifting 6. Place animals in constant dark conditions following Basic Protocol, step 6. Record activity data from these animals for at least 4 days. Based on the onsets of activity (equivalent to CT12, see Basic Protocol, step 10a) over this 4-day period, one can use computer software to predict CT12 for day 5. From this data, calculate CT16 for the animal (i.e., predicted CT12 plus 4[τ/24]). 7. On day 5 of constant darkness, expose the mice to a 15-min light pulse at CT16. Following this, return the mice to constant dark conditions. Record data in constant darkness for 7 days. 8. From the latest recorded data, calculate the CT23 for each mouse as per Basic Protocol, step 6. Expose the mice to a 15-min long light pulse (i.e., lights on) at CT23. Following this, return the mice to constant dark conditions. Record data in constant darkness for 7 days.

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9. Repeat steps 6 through 8 for any number of time points. 10. When data collection is complete, remove mice from the circadian cabinets.

Analyze data 11. Once all wheel running data has been collected, it can be analyzed by appropriate analysis software (e.g., Clocklab or equivalent), as in Basic Protocol, step 10. For analysis: a. Masking: The inclusion of a light pulse in the dark phase will result in acute suppression of mouse activity; a phenomenon known as masking. To calculate the masking effect, compare the activity of the mice during each hour of the light pulse to the activity at the same time on the two preceding days (i.e., activity during which there is no light pulse). b. Dark pulse: Unlike masking (wherein the pulse causes a change in the activity of an animal), the inclusion of a dark pulse during the light phase is not expected to elicit a change in activity. This is because the pulse occurs during the inactive phase of the animal’s activity cycle. If the pulse increases the activity of an animal during its rest phase, it may indicate a poor propensity for sleep. To calculate the effect of the dark pulse, compare the activity of the mice during each hour of the dark pulse to the activity at the same time on the two preceding days (i.e., activity during which there is no dark pulse). c. Phase shifting: For mice in constant dark conditions, light pulses in the early subjective night (e.g., CT16) will cause a delay in the onset of activity (phase delay), whereas light pulses in the late subjective night (e.g., CT 23) will cause an advance in the onset of activity (phase advance). Phase changes in response to light pulses can be determined using computer software. Best fit lines are calculated for activity onsets prior to the light pulse and following the light pulse. The change in phase (φ) is the time difference between the two fitted lines. d. Phase response curve (PRC): An extension of the phase shifting protocol is to generate a phase response curve. This graph visualizes how the circadian oscillator reacts to light pulses at different CTs throughout the circadian cycle. To generate a phase response curve (PRC), modify steps 6 to 8 to calculate φ to light pulses across a range of CTs (every 2 hr). Plot this data as CT against φ to generate the PRC. For further details, see Anticipated Results. SUPPORT PROTOCOL 2

T-CYCLES AND REENTRAINMENT This modified protocol shifts light/dark cycles to measure the entraining ability of a mouse to external conditions (Fig. 5). A T-cycle is defined as an entraining cycle with a period differing from that of the usual 24-hr period. The protocol below describes running T-cycles of 22 and 26 hr (11:11 and 13:13 hr light/dark cycles, respectively). These are the limits of entrainable T-cycles for mice (Jud et al., 2005). However the T-cycle period can be modified as required. For materials, see Basic Protocol.

Assessment of Circadian and LightEntrainable Parameters in Mice

Data collection 1: Run T-cycles 1. Place mice in circadian chambers set to a 12-hr light/dark cycle following Basic Protocol, steps 1 to 5. Record the activity of these mice for ≥5 days and confirm that they have entrained to the light/dark cycle. 2. Modify the light/dark cycles to a 22-hr T-cycle (i.e., 11 hr of light and 11 hr of dark). Monitor the activity of the mice for 10 to 14 cycles to establish whether the animals entrain to the T-cycle.

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A

B 12-hr light/dark cycle

12-hr light/dark cycle

11-hr light/dark cycle

11-hr light/dark cycle

am

1

pm pm 7

1

am am 7

1

pm pm 7

1

7

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Figure 5 Data generated from Support Protocol 2. (A) Actogram showing activity during a 12:12 followed by an 11:11 hr light/dark T-cycle. Actogram is double-plotted on a 24-hr cycle. Actograms are shaded where lights are off. (B) Actogram plotted using the same data as for A above, but the actogram is double plotted on a 22-hr cycle. Actograms are shaded where lights are off. (C) Actogram showing re-entrainment following 6 hr advance and delay shifts in the light/dark cycle. Actograms are shaded where lights are off.

If necessary, monitor animals for longer to fully establish entrainment.

3. Return animals to a 12-hr light/dark cycle for 5 to 10 days, allowing them to re-entrain to the 12-hr light/dark regime. 4. Modify the light/dark cycles to a 26-hr T-cycle (i.e., 13 hr of light and 13 hr of dark). Monitor the activity of the mice for 10 to 14 cycles to establish whether the animals entrain to the T-cycle. If necessary, animals can be monitored for longer to fully establish entrainment.

5. Return the animals to a 12-hr light/dark cycle for 5 to 10 days, allowing them to re-entrain to the 12-hr light/dark regime.

Data collection 2: Re-entrain following a 6-hr shift 6. Once the animals have re-entrained to the 12-hr light/dark cycle, shift the phase of the light/dark cycle to delay by 6 hr (e.g., if the original 12-hr light/dark cycle ran with lights on at 7 a.m. and off at 7 p.m., shift to lights on at 1 p.m. and lights off at 1 a.m.). 7. Leave the animals to re-entrain to the shifted light/dark cycle. 8. Once the animals have entrained to the delayed light/dark cycle, return the animals to the original 12-hr light/dark regime (e.g., if the original light/dark cycle was lights on at 7 a.m. and off at 7 p.m., return the lighting regime to these times). Leave the animals to re-entrain to the shifted light/dark cycle. 9. When the mice have re-entrained, remove them from the circadian cabinets.

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Analyze data 10. Once all wheel running data has been collected, it can be analyzed by appropriate analysis software (e.g., Clocklab or equivalent), as in Basic Protocol, step 11. Both T-cycle and re-entrainment measure an animal’s ability to re-entrain to changing environmental cues. For the T-cycle experiment, most software allows one to re-plot actograms to, e.g., a 22-hr rather than a 24-hr cycle. Once this is done, one of the following three behaviors will be evident: (1) the animal entrains to the new T-cycle with activity onsets coincident with lights-off; (2) the animal entrains to the new T-cycle but with an altered phase angle of entrainment; or (3) the animal fails to re-entrain. For the re-entrainment experiment, determine the number of days it takes an animal to re-entrain following the 6-hr shift in the phase of the light/dark cycle. COMMENTARY Background Information

Assessment of Circadian and LightEntrainable Parameters in Mice

The biochemistry, physiology, and behavior of a great many organisms demonstrate robust oscillations and cycles. We are most familar with the cyclic activity of mammals (e.g., sleep/wake cycles or the daily fluctuations in temperature and hormone levels); however, such rhythmic activity has been demonstrated across a diverse number of other organisms, from the activity and photosensitivity of snails (Block and Davenport, 1982) to the spore formation of bread molds (Lakin-Thomas et al., 2001). The majority of these cycles show a period length of ∼24 hr (1 day), and hence these cycles are known as Circadian (in latin, circa meaning about and diem meaning day) rhythms. Although the period of these cycles is set (or entrained) by environmental cues such as light, temperature, or food, circadian oscillators are self sustaining, and will continue cycling in the absence of external cues. These internal clocks allow the organism to anticipate changes to the environment and therefore to adjust accordingly. The importance of these circadian oscillators is evidenced by the large number of processes that they affect. Gene expression, hormone levels, and a variety of different behaviors are all known to be regulated in a circadian fashion and it is therefore not surprising that circadian disruptions have been implicated in a number of psychiatric and metabolic conditions (Green et al., 2007; Roybal et al., 2007). Because of the importance of these processes the study of circadian rhythms has become a major research area in biological science. The study of whole-organism circadian behaviors presents different challenges depending on the organism being studied. Although nocturnal, the laboratory mouse is a good model for circadian analysis since the underlying neuronal circuitry and genes are conserved

between mice and other mammals, including humans. In addition, the availability of genetically modified mice is an ever expanding resource giving the research community a wide range of mouse based models with which to dissect the processes controlling circadian activity: gene knock outs can be used to assess the contribution of specific proteins to circadian processes (e.g., Bunger et al., 2000), while chemical mutagens such as N-ethyl-Nnitrosourea (ENU) can be used to identify novel circadian genes in a forward genetics approach (Bacon et al., 2004; Godinho et al., 2007). The most widely used method to study circadian activity in mice is to monitor wheel running activity. This utilizes the fact that mice, in cages with free access to running wheels, will preferentially spend a significant proportion of their active phase running on the wheels. Thus, circadian wheel running screens measure the voluntary movements of a mouse in response to different environmental conditions. Since the wheel running activity of the mice is recorded by computer, such screens benefit from being automated and are more robust than many other behavioral assays. An alternative to wheel running that also measures voluntary activity is to measure infrared beam beaks. Here, the movement of the mouse within the cage breaks infrared beams, and these breaks are recorded. Positioning of the beams can also give a measure of feeding and drinking behavior, and therefore beam break cages are often used in metabolic studies. Although both beam breaks and wheel running can be used to assess circadian activity, the basic equipment for wheel running screens is cheaper and simpler to maintain. An alternative to measures of voluntary activity is direct video monitoring. This can give more accurate measurements since all activity will be monitored, not just that leading to wheel running or beam breaks. However,

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video tracking also produces large output files, meaning data storage can become problematic.

Critical Parameters Mouse strain and age Circadian activity has been reported to differ across mouse strains (Schwartz and Zimmerman 1990; Hofstetter et al., 2007), with certain strains being more amenable for circadian analysis than others. For example, the BALB/cJ strain shows high variance in activity and free running circadian period, whereas the C57BL/6 strain is far more consistent in the same parameters (Shimomura et al., 2001). Therefore, when comparing two cohorts of mice, the genetic background of the cohorts should be as closely matched as possible. The best comparison is between littermates, so if possible use siblings for experimental and control groups. In addition, age related changes in circadian phenotype have been reported in mice. Older (16-month-old) mice show a delayed response to light, fragmented activity bouts and increased variability in their rhythms (Valentinuzzi et al., 1997). Older mice therefore make poor candidates for circadian analysis. Mouse disturbance To accurately measure circadian rhythms, the mice being tested should be isolated from external disturbance as much as possible. The room in which circadian chambers are housed should be kept as quiet as possible (external noise can also be kept to a minimum by sound proofing the chambers if necessary). Temperature and air pressure must be maintained at a reasonable consistency. However, it should be noted that light is a more powerful zeitgeber than temperature or air pressure, so minor fluctuations in these conditions can be neglected. Since light is the primary zeitgeber in most conditions, the room should be maintained either in darkness or illuminated using red light filters to minimize accidental light exposure to the animals. Daily animal checks should be performed with as little disruption to the animals as possible. Unless the investigation specifically demands it, food and water should be accessible ad libitum to prevent entrainment to feeding regimes. Light source and intensity A white light source generating a light intensity of ∼150 lux is standard for circadian entrainment. This can be achieved through ei-

ther fluorescent light bulbs or LED bulbs or panels. We use fluorescent tube bulbs, which give some slight variation in light intensity across the chamber, but not to any significant level. One consideration to make when choosing a light source is the heat produced by the lighting. If necessary, an externally placed heat sink may be required for certain sources. A modification to the protocols above can be made by equipping the circadian coffins with a variable light source. Entrainment, masking, and phase shifting are modulated by light intensity and thus the screen can be modified to include changes to the intensity of the light cue. For example, in contrast to the high intensity light pulses used in the masking protocol above, a low intensity light pulse in the dark phase will increase the activity of mice—a phenomenon known as positive masking (Mrosovsky et al., 1999).

Anticipated Results A typical actogram generated by a wildtype C57BL/6J following the Basic Protocol outlined above is shown in Figure 3A. The animal shows good entrainment to the initial 12-hr light/dark regime (as demonstrated by a strong corroboration between the offset of light and the onset of activity), a τ of <24 hr in constant darkness and a τ > 24 hr in constant light. The active phases (α) are also clearly observable on the actogram. The wheel running activity across the different lighting regimes is plotted in Figure 3B. While the total activity is highest for animals in the light/dark regime, ∼95% of this is accounted for by activity in the dark phase of the cycle. Of the three lighting conditions, constant darkness generates the highest activity levels. This is in contrast to constant light, which generates the lowest activity levels. It should also be noted that, after prolonged periods in constant light, disturbances to the circadian period such as arrhythmia or splitting (wherein the rhythm dissociates or ‘splits’ into two separate components) may be observed (see Fig. 3C). Actograms generated following the masking section of Support Protocol 1 are shown in Figure 4A. The activity changes following both light and dark pulses are assessed by the comparison of the activity during the pulse to the activity at the same time points on the previous days. In the masking component, the light pulse in the dark phase of the cycle suppresses activity as the light cue triggers the onset of the rest phase. In contrast, the dark pulse is applied during the inactive phase of the cycle, and thus the dark pulse fails to alter

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the activity of the mouse. Changes to masking behavior are indicative of defects in light input or light response. Phase shifting actograms are shown in Figure 4B-C. Here, a light pulse at CT16 causes a delay in the onset of activity (Fig. 4B), whereas a light pulse at CT23 causes an advance in the onset of activity (Fig. 4C). Expanding the CT range at which the light pulse occurs allows the generation of a phase response curve—an example of which is shown in Figure 4D. The actogram in Figure 5A, double-plotted on a 24-hr scale, shows the effect of switching from a 24-hr to a 22-hr T-cycle. The ability of an animal to entrain to a T-cycle depends upon its ability to accelerate or decelerate its internal clock. For example in the 22-hr T-cycle shown in Figure 5A, the animal must constantly accelerate its internal rhythm. Whether an animal can entrain to T-cycles and how long this entrainment can be maintained will depend upon the light input pathways of that animal and upon the stability of its internal clock. An alternative actogram presenting the same data is shown in Figure 5B. Here the actogram is double-plotted on a 22-hr scale, rather than the 24-hr scale normally used. This presentation format can be used to more clearly demonstrate an animal’s ability to entrain to the 11-hr light/dark cycle. In this example, the animal appears to entrain but with a delayed phase angle of entrainment. Re-entrainment is shown on the actogram in Figure 5C. As described in Support Protocol 2, the initial light/dark cycles run with lights on at 7 a.m. and lights off at 7 p.m. Following 3 days of this regime, the lights on/off times are delayed by 6 hr. The animal takes 3 days to adjust its rhythms and entrain to this shifted lighting regime. The timing for lights on/off are then shifted back to the original 7 a.m./ 7 p.m. regime and the animal takes 6 days to adjust its rhythms and re-entrain to the original lighting regime. As with the T-cycle protocol above, the ability of an animal to re-entrain depends upon light input pathways and the stability of its internal clock.

Time Considerations

Assessment of Circadian and LightEntrainable Parameters in Mice

The full circadian screen outlined above can take 30 to 40 days to complete. However, the protocols outlined above can be easily adapted and modified according to need and time constraints. For example, many circadian screens omit the analysis in constant light, leaving just the entrainment and constant dark-

ness measurements. In addition, it is possible to combine different parts of the analysis. For example, in the authors’ screens, the authors combine the Basic Protocol with the masking section of Support Protocol 1 by including a masking pulse in the initial light/dark cycling performed at the beginning of the screen. It is also worth noting that, although running a complete screen will take several days, the majority of the experiment is fully automatic, requiring little input from the investigator during the screen. Although it is possible to shorten some of the steps, decreasing the time in a new lighting regime to <7 days is not recommended. The first 2 to 3 days in which an animal is placed in a novel lighting environment will be stressful to the animal, and therefore this may affect their wheel running activity. In addition to this, it is advisable to take data from a minimum of 5 to 7 days to minimize the variability of the measurement. However, as noted above, this is the minimum time. It is recommended that animals are allowed to respond to lighting regimes for at least 10 to 14 days, allowing for more subtle phenotypes that may only become apparent after additional time in the lighting condition.

Literature Cited Aschoff, J. 1981. Free running and entrained circadian rhythms. In Handbook of Behavioural Neurobiology: Biological Rhythms (J. Aschoff, ed.) pp. 81-89. Plenum, New York. Bacon, Y., Ooi, A., Kerr, S., Shaw-Andrews, L., Winchester, L., Breeds, S., Tymoska-Lalanne, Z., Clay, J., Greenfield, A.G., and Nolan, P.M. 2004. Screening for novel ENU-induced rhythm, entrainment and activity mutants. Genes Brain Behav. 3:196-205. Block, G.D. and Davenport, P.A. 1982. Circadian rhythm in Bulla gouldiana: Role of the eyes in controlling motor behaviour. J. Exp. Zoo. 224:57-63. Bunger, M.K., Wilsbacher, L.D., Moran, S.M., Clendenin, C., Radcliffe, L.A., Hogenesch, J.B., Simon, M.C., Takahashi, J.S., and Bradfield, C.A. 2000. MOP3 is an essential component of the master circadian pacemaker in mammals. Cell 103:1009-1017. Church, C., Moir, L., McMurray, F., Girard, C., Banks, G.T., Tebul, L., Wells, S., Bruning, J.C., Nolan, P.M., Ashcroft, F.M., and Cox, R.D. 2010. Overexpression of Fto leads to increased food intake and results in obesity. Nat. Genet. 42:1086-1092. Daan, S. and Pittendrigh, C.S. 1976a. A functional analysis of circadian pacemakers in nocturnal rodents. I. The stability and lability of spontaneous frequency. J. Comp. Physiol. A 106:223252.

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Daan, S. and Pittendrigh, C.S. 1976b. A functional analysis of circadian pacemakers in nocturnal rodents. II. The variability of phase response curves. J. Comp. Physiol. A 106:253-266. Daan, S. and Pittendrigh, C.S. 1976c. A functional analysis of circadian pacemakers in nocturnal rodents. III. Heavy water and constant light: Homeostasis of frequency. J. Comp. Physiol. A 106:267-290. Daan, S. and Pittendrigh, C.S. 1976d. A functional analysis of circadian pacemakers in nocturnal rodents. IV. Entrainment: Pacemaker as clock. J. Comp. Physiol. A 106:291-331. Daan, S. and Pittendrigh, C.S. 1976e. A functional analysis of circadian pacemakers in nocturnal rodents. V. Pacemaker structure: A clock for all seasons. J. Comp. Physiol. A 106:333-355. Godinho, S.I.H., Maywood, E.S., Shaw, L., Tucci, V., Barnard, A.R., Busino, L., Pagano, M., Kendall, R., Quwailid, M.M., Romero, M.R., O’Neill, J., Chesham, J.E., Brooker, D., Lalanne, Z., Hastings, M.H., and Nolan, P.M. 2007. The after-hours mutant reveals a role for Fbxl3 in determining mammalian circadian period. Science 316:897-900. Green, C.B., Douris, N., Kojima, S., Strayer, C.A., Fogerty, J., Lourim, D., Keller, S.R., and Besharse, J.C. 2007. Loss of nocturnin, a circadian deadenylase, confers resistance to hepatic steatosis and diet-induced obesity. Proc. Natl. Acad. Sci. U.S.A. 104:9888-9893. Hara, H., Nolan, P.M., Scott, M.O., Bucan, M., Wakayama, Y., and Fischbeck, K.H. 2002. Running endurance abnormality in mdx mice. Muscle Nerve 25:207-211. Hofstetter, J.R., Svihla, D.A., and Mayeda, A. 2007. A QTL on mouse chromosome 12 for the genetic variance in free running circadian period between inbred strains of mice. J. Circadian Rhythms 5:7. Jud, C., Schmutz, I., Hampp, G., Oster, H., and Albrech, U. 2005. A guideline for analysing circadian wheel-running behaviour in rodents un-

der different lighting conditions. Biol. Proced Online 7:101-116. Lakin-Thomas, P.L., Gooch, V.D., and Ramsdale, M. 2001. Rhythms of differentiation and diacylglycerol in Neurospora. Philos. Trans. R Soc. Lond. B Biol. Sci. 356:1711-1715. Mrosovsky, N., Foster, R.G., and Salmon, P.A. 1999. Thresholds for masking responses to light in three strains of retinally degenerate mice. J. Comp. Physiol. 184:423-428. Panda, S., Sato, T.K., Castrucci, A.M., Rollag, M.D., DeGrip, W.J., Hogenesch, J.B., Provencio, I., and Kay, S.A. 2002. Melanopsin (Opn4) requirement for normal light-induced circadian phase shifting. Science 298:22132216. Roybal, K., Theobold, D., Graham, A., DiNieri, J.A, Russo, S.J., Krishnan, V., Chakravarty, S., Peevey, J., Oehrlein, N., Birnbaum, S., Vitaterna, M.H., Orsulak, P., Takahashi, J.S., Nestler, E.J., Carlezon, W.A., and McClung, C.A. 2007. Mania-like behavior induced by disruption of CLOCK. Proc. Natl. Acad. Sci. U.S.A. 104:6406-6411. Schwartz, W.J. and Zimmerman, P. 1990. Circadian timekeeping in BALB/c and C57BL/6 inbred mouse strains. J. Neurosci. 10:3685-3694. Shimomura, K., Low-Zeddies, S.S., King, D.P., Steeves, T.D.L., Whiteley, A., Kushla, J., Zemenides, P.D., Lin, A., Vitaterna, M.H., Churchill, G.A., and Takahashi, J. 2001. Genome-wide epistatic interaction analysis reveals complex genetic determinants of circadian behaviour in mice. Genome Res. 11:959980. Siepka, S.M. and Takahashi, J.S. 2005. Methods to record circadian rhythm wheel running activity in mice. Methods Enzymol. 393:230-239. Valentinuzzi, V.S., Scarbrough, K., Takahashi, J., and Turek, F.W. 1997. Effects of aging on the circadian rhythm of wheel-running activity in C57BL/6 mice. Am. J. Physiol. 273:R1957R1964.

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Behavioral Measures of Pain Thresholds Michael S. Minett,1,2 Kathryn Quick,1 and John N. Wood1,2 1

Molecular Nociception Group, Wolfson Institute for Biomedical Research, University College London, London, United Kingdom 2 London Pain Consortium, Kings College London, London, United Kingdom

ABSTRACT Pain afflicts a fifth of the population, and animal models have proven useful in target validation and analgesic drug development. Thresholds to pain are tested by applying a sensory stimulus, such as heat or pressure, and observing the resulting withdrawal behavior. Sensitized pain models involve provoking an inflammatory response or damaging the nerves themselves, and testing the changes in pain threshold. In this article, mouse models of acute mechanical and thermal pain and inflammatory, visceral, and neuropathic pain are discussed. These behavioral measures can be used to phenotype transgenic mice for target validation and mechanistic studies, as well as to screen potential analgesic C 2011 by John Wiley & Sons, Inc. compounds. Curr. Protoc. Mouse Biol. 1:383-412  Keywords: nociception r reflex r pain r neuropathic r inflammatory r visceral

INTRODUCTION Animal models of pain have been critical for our current understanding of the underlying mechanisms of pain and the development of pharmacological interventions. All human analgesic drugs are active in rodent models of pain, although the converse is not necessarily true—for example, NK1 antagonists are analgesic in rodents but not man. Nonetheless, animal models have proven important in both target validation and understanding mechanisms underlying pain sensation. These models explore various aspects of pain mainly through the interpretation of physiological responses, such as withdrawal reflexes or more complex escape behaviors. The behavioral responses occur in reaction to thermal, mechanical, and chemical stimuli. Sensory stimuli have a dynamic range—for example, mechanical sensation ranges from light touch to noxious pressure, and thermal sensation ranges from noxious cold through warm temperatures up to noxious heat. The thresholds to these different types of acute stimuli can be tested using the protocols described here. Acute pain is a vitally important signaling system that warns us of imminent and/or actual tissue damage. However, chronic pain has no biologically relevant purpose and dramatically reduces quality of life and productivity. The cost of pain is estimated at over £13 billion per year in the UK (British Pain Society, http://www. britishpainsociety.org/bps nl sum 2009.pdf) and over $100 billion per year in the U.S. (Mayday Fund, http://www.maydaypainreport.org/). There are a number of chronic pain models outlined in this unit that model some of the different types of pain experienced in humans—inflammatory and neuropathic. When inflammation or injury is induced using inflammatory, visceral, or neuropathic pain models, the thresholds to sensory stimuli change—producing hyperalgesia, an enhanced response to a normally painful stimulus—and allodynia, a painful response to a normally innocuous stimulus. The most commonly used animal in pain research has been the laboratory rat (Mogil et al., 2001), but more recently the application of these pain models to mice with targeted gene mutations has proved to be a powerful tool. However, the behavioral responses of rats and mice can differ; as Mogil et al. (2001) succinctly explain, “mice are not small Current Protocols in Mouse Biology 1: 383-412, September 2011 Published online September 2011 in Wiley Online Library (wileyonlinelibrary.com). DOI: 10.1002/9780470942390.mo110116 C 2011 John Wiley & Sons, Inc. Copyright 

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rats.” It is important to bear this in mind if applying an established rat model to mice for the first time. We describe here the steps required to characterize sensory thresholds of mice and model inflammatory, neuropathic, and visceral pain states. The first section contains protocols for establishing acute pain thresholds to thermal stimuli, using tail flick (see Basic Protocol 1 and Alternate Protocols 1 and 2), hot and cold plate test (see Basic Protocols 2 and 3), Hargreaves’ apparatus (see Basic Protocol 4 and Support Protocol 1), and acetone evaporation (see Basic Protocol 5); the second section provides protocols for testing threshods to mechanical stimuli using von Frey hairs (see Basic Protocol 6, Support Protocol 2, and Alternate Protocol 3) and the Randall-Selitto apparatus (see Basic Protocol 7 and Alternate Protocol 4). Additional sections outline animal models of pain split into three types—inflammatory pain (see Basic Protocol 8 and Alternate Protocol 4), visceral pain (see Basic Protocol 9 and Support Protocol 3), and neuropathic pain (Basic Protocol 10 and Alternate Protocols 5 and 6).

STRATEGIC PLANNING While planning an experiment, it is important to consider the following: issues.

Ethical considerations Before beginning any work with mice, proper training and protocol approval must be obtained according to the European Directive 86/609/EEC, Animals (Scientific Procedures) Act 1986, or equivalent legislation in other parts of the world, and the study must conform to government regulations. Also, it is advisable to consult the International Association for the Study of Pain ethical guidelines (http://www.iasp-pain.org) for more details and advice. Also see the relevant article in Current Protocols in Mouse Biology, Baertschi and Gyger (2011). Cut-off time The nature of nociceptive testing means that it is important to limit the risk of tissue damage by limiting unnecessary exposure to noxious stimuli. This is especially true when working with transgenic mice and analgesics, where nociceptive thresholds may be altered. It is recommended to conduct a literature search for previous findings for your specific mouse strain. If this is unavailable, a cut-off of three times the average withdrawal response of na¨ıve mice is desirable. Motor function It is critical to demonstrate that the motor function and coordination of any mice undergoing nociceptive testing are intact. A loss in motor function and coordination may alter withdrawal latencies or other nociceptive behaviors. Operator blinding The test operator should be blinded to the test groups (e.g., genotype, treatment etc). This ensures unbiased results. Pain behaviors Throughout this chapter, “pain behaviors” or “nocifensive behaviors” are referred to frequently as outcome measures of many of the tests. Common pain behaviors are: Behavioral Measures of Pain Thresholds

Flinching Licking Shaking

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Biting Jumping Stretching or squashing the abdomen Guarding of hindpaw Change in posture. It is essential to become practiced in the “normal” behavior of a mouse, so that any differences in behavior during a nociceptive test can be noted. It is also important to note if anything may be affecting pain behavior, such as grooming, sleeping, defecating, or distraction by changes in noise or light levels.

Controls As with all scientific experiments, proper controls are essential. Always test a group of control mice (e.g., wild-type or non-treated) at the same time as the test group. When administering drugs or compounds, it is good practice to use an appropriate vehicle control. Similarly, with surgical models, it is important to perform sham surgery on littermates alongside any real surgery. This can indicate if any other factors have contributed to any changes in nociceptive behavior, Baseline pain measures Before inducing any of the inflammatory, visceral, or neuropathic pain models, determine which pain measures are to be investigated and obtain consistent baseline measurements. It is recommended that at least two baseline studies be performed. TESTING THRESHOLDS TO THERMAL STIMULI Heating the skin to temperatures above 45◦ C is generally considered noxious, and is the most commonly used method for assessing nociception. When using a noxious heat stimulus, it is critical to limit exposure time in order to avoid tissue damage.

Radiant Heat Tail-Flick This protocol describes the Radiant heat tail-flick test developed 70 years ago by D’Amour and Smith (1941). The tail-flick was one of the first, and subsequently most commonly used, nociceptive tests. The D’Amour and Smith technique involves application of focused beam of radiant heat and measuring the flexor withdrawal reflex latency. It is a robust thermal nociception assay that is stable during repeated tests.

BASIC PROTOCOL 1

Materials Mice of optimal age (6 to 8 weeks) Mouse restrainer (consisting of base plate, body tube, and head cover) Infrared heat source with built-in timer and motion detector. (e.g., Tail Flick Analgesia Meter #33; IITC Life Science, http://www.iitcinc.com/) Infrared heat-flux radiometer (Ugo Basile, cat. no. 37300), for calibrating and maintaining apparatus Habituation 1. Clean the mouse restrainer. 2. Place a single subject mouse into the restrainer (Fig. 1). It is recommended to use a restrainer with a removable base plate. Remove the base plate and cover the tube of the restrainer with tissue to make an ideal hiding place for a mouse. Next, place the test subject mouse next to the tube opening and gently tug on its tail to trigger escape behavior, encouraging the mouse to enter the restrainer tube. If the mouse

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Figure 1

Mouse in a restrainer.

repeatedly places its head within the tube only to withdraw it again, this may indicate that the bore is too narrow and a larger restrainer is required.

3. Adjust head cover for body length of mouse. Adjust the head cover so that the mouse’s movements are minimized without causing unnecessary distress.

4. Check tail for evidence of injury (e.g., from fighting). If the tail is significantly injured, consider using another mouse or testing the injured mouse at a later time.

5. Place restrainer on top of the tail-flick apparatus and leave mouse until settled (average 10 min). A cover can be placed over the restrainer to help calm the animal, but this must be removed before the test so the animal can be seen fully.

Testing 6. While the radiant heat light source is in the “idle” state, guide the tail over the test head so the stimulus spot is midway along the tail. Location of the stimulus spot used on the tail may vary; the most important point is to keep the stimulus site constant within your experiment. If tail color is not constant over the whole tail or not the same for all subjects, it is suggested to use the hot water immersion protocol (Alternate Protocol 1), since tail color has been shown to alter the response latency of this test (Vetulani et al., 1988).

7. Switch the radiant heat light source to the high-intensity setting and leave in place until nociceptive reflex/withdrawal behavior (e.g., a sudden twitch of the tail) is observed. The user interface, controls, and feature of the tail-flick apparatus vary between manufacturers and models. Please consult user manual for exact details

8. Once a response time has been recorded, repeat the test twice more. The mouse should be allowed 5 min to recover each time, and the next stimulus spot should be 10 mm to either side of the original stimulus spot. Behavioral Measures of Pain Thresholds

Threshold calculation 9. Use the multiple test results to calculate an average response time for each mouse.

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Hot Water Immersion Tail-Flick Ben-Bassat et al. (1959) describe an alternative version of the above protocol using a hot-water bath (typically 46◦ to 52◦ C) instead of an infrared radiant heat source. This protocol minimizes the effects of tail color or stimulation site, which can alter heattransfer properties. Also, this version of the tail flick test is particularly useful if a lab has no specialist in analgesia assay equipment.

ALTERNATE PROTOCOL 1

Additional Materials (also see Basic Protocol 1) Mice of optimal age (6 to 8 weeks) Water bath Thermometer (accurate to 0.5◦ C) Timer Habituation 1. Habituate the mouse to the apparatus as described in Basic Protocol 1, steps 1 to 5. Testing 2. Heat water bath to 46◦ to 52◦ C. Make sure temperature is stable. Start with the lower end of the temperature range, and adjust it as necessary so that withdrawal latency occurs within a reasonable amount of time for the majority of the mice.

3. Gently guide the distal half of the tail into the water bath and time the withdrawal latency. This is where it can be useful to have an assistant to handle the mouse while the observer operates the timer. It is suggested that the experimenter with the most mouse husbandry experience restrain and handle the mice.

4. Allow the mouse 60 sec to recover before repeating test. Each mouse can be tested at least three times per session.

Threshold calculation 5. Use the multiple test results to calculate an average response time for each mouse. Cold Water Immersion Tail-Flick Pizziketti et al. (1985) describe an alternative version of Alternate Protocol 1 using a coldwater bath (0◦ to 5◦ C) instead of a hot water bath. The protocol is identical to the radiant heat tail-flick with the exception that the distal half of the tail is immersed in cool water. It is possible to test subzero (as low as −18◦ C) temperatures by using other fluids such as ethanol. However, this increases risk of tissue damage, and, therefore, cut-off times should be reduced accordingly. The needed materials are as tabulated for Alternate Protocol 1. Habituation, testing, and threshold calculation are as described for Alternate Protocol 1.

Hot Plate: Nociceptive Response to Noxious Heat This protocol describes the steps required to assess behavioral responses to temperatures up to 55◦ C. The most commonly used version of the hot plate test was originally described by Woolfe and Macdonald (Macdonald et al., 1946) and later modified by Eddy and Leimbach (1953). In contrast to the tail flick test (Basic Protocol 1), which is a spinal response, the hot plate test seems to represent a supraspinal thermal assay (discussed in more detail in the Commentary).

Materials Mice of optimal age (6 to 8 weeks) Hot plate apparatus with enclosure (e.g., Hot/Cold Plate; Ugo Basile, cat. no. 35100) Surface thermometer, for calibrating and maintaining apparatus (optional)

ALTERNATE PROTOCOL 2

BASIC PROTOCOL 2

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Habituation 1. Set plate temperature to match that of the test room and clean test plate surface. 2. Place a single subject mouse into the test chamber and leave to explore undisturbed for 15 min. Exposure to novel environments may mask or alter withdrawal threshold

3. Remove mouse and clean any urine or feces from the plate surface. Urine or fecal matter on the hindpaws may alter the heat transfer properties of the hot plate.

4. Set the hot plate to the test temperature. This is typically between 50o and 56◦ C.

Testing 5. Place the habituated mouse onto the hot plate, once temperature has been reached. 6. Measure the time latency to first nocifensive behavior (e.g., hindpaw lift or jumping). Using forepaw behaviors is NOT recommended, since the forepaws are commonly involved in grooming and exploratory behaviors. Similarly, a simple hindpaw lift may be due to grooming and exploration.

7. Quickly remove mouse from test surface to avoid unnecessary suffering and/or tissue damage. Repeat testing within one session is NOT recommended. Repeated exposure to the hotplate test at noxious or even innocuous temperatures can decrease behavioral response latencies (Mogil et al., 2001).

Preparation of next mouse 8. Place tested mouse into a separate cage from untested mice. Mice can communicate via a variety of olfactory and ultrasound systems, which can be a source of anxiety and fear (Gray, 1978).

9. Return the test plate temperature to that of the room and clean surface in preparation.

Multiple temperatures 10. If testing same mice at multiple temperatures, then separate the tests by at least 24 hr to avoid the effects of stress and repeated testing on latency times (detail above). BASIC PROTOCOL 3

Cold Plate: Noxious Cold This protocol describes the steps to assess behavioral responses to noxious cold. A number of different versions of the cold plate test have been described, such as Lee et al. (1999), Zimmermann et al. (2007), and Abrahamsen et al. (2008). However, all versions involve observing hindpaw nociceptive behaviors of mice placed onto a surface maintained at a temperature between 0◦ to 4◦ C. Unlike nocioceptive responses to noxious heat, noxious cold stimuli do not consistently induce nocioceptive responses within well established time frames. Therefore, it is useful to record the number of nocioceptive responses within a fixed period, as well as the latency to the first response.

Materials

Behavioral Measures of Pain Thresholds

Mice of optimal age (6 to 8 weeks) Cold plate apparatus with enclosure (e.g. Hot/Cold Plate; Ugo Basile, cat. no. 35100) Surface thermometer, for calibrating and maintaining apparatus (optional)

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Habituation 1. Set plate temperature to match that of the test room, and clean test plate surface. 2. Place a single subject mouse into the test chamber and leave to explore undisturbed for 15 min. Exposure to novel environments may mask or alter behavioral response.

3. Remove mouse and clean any urine or feces from the plate surface. Urine or fecal matter on the hindpaw may alter the heat transfer properties of the cold plate.

4. Set the cold plate to the test temperature.

Testing 5. Place the habituated mouse onto the cold plate, once temperature has been reached. 6. Measure the time latency to first nocifensive behavior (e.g., hindpaw fluttering/licking). Using forepaw behaviors is NOT recommended since the forepaws are commonly involved in grooming and exploratory behaviors. Similarly, a simple hindpaw lift may be due to grooming and exploration

7. Measure the amount of time the mouse spends displaying nociceptive behaviors for 5 min. 8. Quickly remove mouse from the test surface to avoid unnecessary suffering and/or tissue damage.

Preparation of next mouse 9. Place tested mouse into a separate cage from untested mice. Mice can communicate via a variety of olfactory and ultrasound systems, which can be a source of anxiety and fear (Gray, 1978).

10. Return the test plate temperature to that of the room and clean surface in preparation.

Hargreaves’ Test: Withdrawal Threshold to Noxious Heat This protocol describes the steps required to assess noxious heat thresholds to radiant heat. The Hargreaves test was originally described as a method for measuring thermal nociception in cutaneous hyperalgesia, in response to carrageenan-induced inflammation (Hargreaves et al., 1988). However, it can also be used for comparative phenotyping of transgenic mice, measuring analgesic drug efficacy etc.

BASIC PROTOCOL 4

Materials Mice of optimal age (6 to 8 weeks) Hargreaves’ apparatus, infrared or light (IITC Life Science, cat. no. 390; http://www.iitcinc.com/; Ugo Basile, cat. no. 37370) Raised glass pane with clear plastic enclosures Optional: Infrared heat-flux radiometer (Ugo Basile, cat. no. 37300), for calibrating and maintaining apparatus Habituation 1. Design an identification key, assigning each subject mouse to a specific test compartment. If the test group (e.g., genotype, dose, control etc.) of subject mice is known, the above task cannot be performed by the test operator, as it may lead to test bias.

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2. Place the subject mice into the assigned test compartments, with a piece of tissue to absorb any urine that may collect within the test compartment. 3. Leave subject mice to explore test compartments, undisturbed for at least 1 hr. Exposure to novel environments may mask or alter withdrawal thresholds

4. Check that exploratory behavior has stopped or decreased to a minimum. If not, leave undisturbed for an additional 30 min before rechecking. It may be necessary to repeat this more than once.

5. Remove any tissue fragments from the test compartments. 6. Use a fresh tissue to remove as much urine and feces as possible from the glass floor of each test compartment. Slow and gentle movements will minimize any disturbance to the subject mice. If the mouse’s paw is submerged in substantial pool of urine, it may alter the Hargreaves machine’s ability to constantly heat the plantar surface of the paw. Any fecal matter smeared on the glass floor may also have a similar effect.

7. Leave the mice to resettle for any additional 5 to 10 min following cleaning. The test operator should remain in the room during this period. Remaining in the room during this period familiarizes the subject mice with the test operator’s presence, meaning the test operator is less of a novel event during the test.

Testing 8. While the radiant heat light source is in the “idle” state, use the guide mirror on top of the test head to position the stimulus spot over the plantar surface of the left hindpaw. The majority of studies use the hindpaw. Choosing either left or right is arbitrary; however, it is advisable to consistently use the same hindpaw for all animals and repeats during a test.

9. Switch the radiant heat light source to the high intensity setting and leave in place until a nociceptive reflex/withdrawal behavior is seen. The user interface, controls, and features of the Hargreaves apparatus vary between manufacturers and models.

10. If the behavioral response is ambiguous (e.g., the mouse engages in grooming behavior or the movement is triggered by a sudden noise), then disregard the reading. Otherwise, record reading and move on to the next mouse. Allow the mouse to recover by ensuring at least 2 min between each stimulus on the same mouse. It is important that the mouse be still but not asleep when testing. Similarly, the mice should not be grooming, as this may influence the response (Callahan et al., 2008).

11. Once a response time has been recorded for each mouse, return to the first mouse and repeat previous step. Several response times (three is common practice) can be recorded from each mouse within one session without causing harm and thereby altering the response. 12. Upon completion of test, use the identification key to double check that the response latency time corresponds to the correct mouse.

Threshold calculation 13. Use the multiple test results to calculate an average response for each mouse. Behavioral Measures of Pain Thresholds

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Alternative Measurement for Hargreaves’ Apparatus Paw withdrawal latency can also be supplemented by additional behavioral outcome measures:

SUPPORT PROTOCOL 1

Velocity of paw withdrawal (ordinal score of 0 if movement complete within 1 sec, or a score of 1 if persisted beyond 1 sec). Time spent licking affected paw following withdrawal. Duration of hindpaw withdrawal/lift until affected paw is returned to glass surface. These additional behavioral measures can be seen long after the paw withdrawal latency has return to baseline, following induced hyperalgesia (Hargreaves et al., 1988).

Acetone Evaporation Test: Innocuous Cooling and Cold Allodynia 10◦

15◦ C

This protocol describes the steps required to assess cold sensitivity in the to range, which is usually considered innocuous. The application of acetone causes a rapid decrease in temperature (of ∼10◦ C). This can be used to examine sensitivity to cooling, or more commonly as a measure of cold allodynia following an inflammatory or neuropathic pain model (see relevant sections below).

BASIC PROTOCOL 5

Materials Mice of optimal age (6 to 8 weeks) Acetone Test compartments with a mesh floor (e.g., Mesh Stand; IITC Life Science, cat. no. 410; http://www.iitcinc.com) Acetone applicator (blunting two 33-G needles and inserting them in either end of a 10-cm length of thin tubing, and using a 5-ml syringe, have approved successful previously; Fig. 2) Two timers

Figure 2

Acetone applicator.

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Habituation 1. Design an identification key, assigning each subject mice to a specific test compartment. If the test group (e.g., genotype, dose, control etc.) of subject mice is known, the above task cannot be performed by the test operator, as it may lead to test bias.

2. Place the subject mice into the assigned test compartments. 3. Leave subject mice to explore test compartments, undisturbed for 1 hr. Exposure to novel environments may mask or alter withdrawal thresholds

4. Check exploratory behavior has stopped or decreased to a minimum. If not, leave undisturbed for an additional 30 min before rechecking. It may be necessary to repeat this more than once.

5. Once settled, have the test operator sit in the room for 5 min before starting the test. Remaining in the room during this period familiarizes the subject mice with the test operator’s presence, meaning that the test operator is less of a novel event during the test.

Testing 6. Gently apply pressure to the syringe to convex the acetone meniscus at the open end of the needle (see Fig. 2). 7. Apply the acetone to the center of the hindpaw and immediately start your first timer. It is important that the mouse be still but not asleep when testing. Similarly, the mice should not be grooming, as this may influence the response (Callahan et al., 2008).

8. Use your second timer to measure the amount of time mice displays nocifensive behavior (e.g., hindpaw fluttering/licking). It is recommended that two people perform this test, one person to operate the timers and a second applying the acetone.

9. Repeat this at least three times with approximately 1 min between each application.

Threshold calculation 10. Use the multiple test results to calculate an average response for each mouse. TESTING THRESHOLDS TO MECHANICAL STIMULI Pain can result from a range of noxious stimulus modalities. Mounting evidence shows that these pain modalities diverge at the primary sensory neurons as well as at the spinal and supraspinal levels. Therefore it is critical to apply a full range of tests when characterizing nociceptive responses. BASIC PROTOCOL 6

Behavioral Measures of Pain Thresholds

Von Frey Test: Light Touch Perception Threshold This protocol describes the steps required to assess touch thresholds in mice. Touch thresholds in humans and animals have been determined using von Frey hairs since the 1890s. The calibrated monofilaments are most commonly made of nylon and apply a consistent accurate force when used correctly. The up-down method for obtaining the 50% threshold using von Frey hairs was described by Chaplan et al. (Chaplan et al., 1994). This method makes use of the statistical formula described by Dixon in 1980 to determine the LD50 (Dixon, 1980).

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Materials Mice of optimal age (6 to 8 weeks) Test compartments with a mesh floor (e.g., Mesh Stand; IITC Life Science, cat. no. 410; http://www.iitcinc.com) Calibrated set of von Frey hairs (e.g., Stoelting Touch Test Sensory Evaluator) Habituation 1. Design an identification key, assigning each subject mice to a specific test compartment. If the test group (e.g., genotype, dose, control etc.) of subject mice is known, the above task cannot be performed by test operator as it may lead to test bias.

2. Place the subject mice into the assigned test compartments. 3. Leave subject mice to explore test compartments, undisturbed for at least 1 hr. Exposure to novel environments may mask or alter withdrawal thresholds.

4. Check exploratory behavior has stopped or decreased to a minimum. If not, leave undisturbed for an additional 30 min before rechecking. It may be necessary to repeat this more than once.

5. Once settled the test operator should sit in the room for 5 min before starting the test. Remaining in the room during this period familiarizes the subject mice with the test operator’s presence, meaning the test operator is less of a novel event during the test.

Figure 3 Mouse paw. Circle indicates the area of the plantar surface of the paw to be stimulated (e.g., with von Frey hair), as well as the site for intraplantar injection. Scale bar = 1 mm.

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Testing 6. Apply the first von Frey hair, perpendicular to the paw (Fig. 3), until it buckles slightly, and hold for 3 sec. If a response (pain behavior) is seen, mark the first column of the scoring grid with an X. If no response is seen mark with an O. It is important that the mouse should be still but not asleep when testing. Similarly the mice should not be grooming as this may influence the response (Callahan et al., 2008).

7. If the last hair produced a response, select the next lower weight hair. Conversely, if last hair failed to produce a response, select the next higher weight hair. Ensure that the application of the hair was correct before considering the response or no response. Leave approximately 1 min between each application. If may be time efficient to serially time each mouse, once you return to the first mouse a sufficient amount of time should have passed.

8. Continue applying increasing or decreasing weighted hairs and marking the results in the first column until a change in response occurs—e.g., XXXO or simply OX. 9. Once this perception threshold has been crossed, continue applying increasing (no response) or decreasing (response) weighted hairs, and record the next five applications.

Threshold calculation 10. Use the reference table (Table 1) to look up the corresponding κ value for each response series. Note that the κ value is inverted if the series begins with X (or multiple Xs). Table 1 von Frey Kappa Value Reference Tablea

K for test series whose first part is:

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O

OO

OOO

OOOO

XOOOO

−0.547

−0.547

−0.547

−0.547

OXXXX

XOOOX

−1.250

−1.247

−1.246

−1.246

OXXXO

XOOXO

0.372

0.380

0.381

0.381

OXXOX

XOOXX

−0.169

−0.144

0.142

−0.142

OXXOO

XOXOO

0.022

0.039

0.040

0.040

OXOXX

XOXOX

−0.5

−0.458

−0.453

−0.453

OXOXO

XOXXO

1.169

1.237

1.247

1.248

OXOOX

XOXXX

0.611

0.732

0.756

0.758

OXOOO

XXOOO

−0.296

−0.266

−0.263

−0.263

OOXXX

XXOOX

−0.831

−0.763

−0.753

−0.752

OOXXO

XXOXO

0.831

0.935

0.952

0.954

OOXOX

XXOXX

0.296

0.463

0.500

0.504

OOXOO

XXXOO

0.500

0.648

0.678

0.681

OOOXX

XXXOX

−0.043

0.187

0.244

0.252

OOOXO

XXXXO

1.603

1.917

2.000

2.014

OOOOX

XXXXX

0.893

1.329

1.465

1.496

OOOOO

X

XX

XXX

XXXX

−K for test series whose first part is: a The number of responses before the response threshold is crossed determines the column. The following

pattern of responses determines the row. Note that the κ value is inverted if the response series prior to threshold crossing with X. Current Protocols in Mouse Biology

11. Use the formula below to calculate the 50% threshold (grams).

50% threshold = (10[χ+κδ] )/10,000) χ = log of the final von Frey hair used κ = tabular value δ = log of mean difference between stimuli (typically 0.224 for most von Frey sets). Repeated Measures Von Frey Test This alternative to the up-down method for the von Frey test (Basic Protocol 6) is used to assess light touch. In contrast to the up-down method, the repeated measures test enables testing of both sub- and supra-threshold stimuli.

SUPPORT PROTOCOL 2

For materials, see Basic Protocol 6. For habituation procedures, see Basic Protocol 6.

Testing 1. Apply the lightest von Frey hair, perpendicular to the paw (Fig. 3), until it buckles slightly, and hold for 3 sec. 2. Repeat application of each filament an additional four times at 60 second intervals. 3. Record number of responses and repeat steps 1 and 2 using the next hair in order of force. The upper cut-off should be 6 g. von Frey hairs above this weight will simply lift the hindpaw without buckling.

Threshold calculation 4. Determine the threshold as the von Frey hair that elicited a withdrawal response in 40% (two out of five) or more of applications. Automatic Von Frey Test Digital von Frey instruments are also available. The mice are habituated in the same way, but the automatic von Frey instrument applies the hair perpendicular to the paw when the button is pressed, and returns a threshold value. The speed of acceleration and maximum force applied can be altered. This protocol measures the absolute threshold and not a calculated 50% threshold, as with the manual von Frey hairs.

ALTERNATE PROTOCOL 3

Materials Mice of optimal age (6 to 8 weeks) Test compartments with a mesh floor (e.g., Mesh Stand; IITC Life Science, cat. no. 410; http://www.iitcinc.com) Automatic von Frey apparatus (e.g., Electronic von Frey IITC Life Science, cat. no. 2390; http://www.iitcinc.com/) Additional reagents and equipment for habituation of mice to von Frey apparatus (Basic Protocol 6) Habituation 1. Habituate mice as described in Basic Protocol 6. Testing 2. Using the mirror, position the automatic von Frey probe directly underneath the stimulus area of the hindpaw. It is important that the mouse should be still but not asleep when testing. Similarly, the mice should not be grooming as this may influence the response (Callahan et al., 2008).

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3. Extend the probe at the chosen acceleration rate by pressing start button. Ensure that the probe is hitting the paw at the correct point (Fig. 3) and not the wire mesh. The probe will automatically retract and display the threshold value, following a withdrawal response.

4. Record a maximum of six responses. Leave ∼1 min between each application of the probe to the same mouse.

Threshold calculation 5. Average the threshold values obtained for each mouse. BASIC PROTOCOL 7

Randall-Selitto Test: Noxious Mechanical Pressure This protocol outlines the steps to determine the noxious pain threshold of mouse using a modified version of the Randall-Selitto test that applies pressure to the tail via a 3mm2 blunt probe (Takesue et al., 1969). Randall and Selitto originally described the application of uniformly increasing pressure to the rat paw as a measure of inflammation, but measuring the tail in the mouse is more reliable (Randall and Selitto, 1957).

Materials Mice of optimal age (6 to 8 weeks) Mouse restrainer (consisting of base plate, body tube, and head cover) Randall-Selitto apparatus (e.g., Analgesy-Meter; Ugo Basile, cat. no. 37215) Habituation 1. Clean the mouse restrainer. 2. Place a single subject mouse into a restrainer (Fig. 1). It is recommended to use a restrainer with a removable base plate since it facilitates getting the mouse into the restrainer. Remove the base plate and cover the tube of the restrainer with tissue to make an ideal hiding place for a mouse. Next, place the test subject mouse next to the tube opening and gently tug on its tail to trigger escape behavior, encouraging the mouse to enter the restrainer tube. If the mouse repeatedly places its head within tube only to withdraw again, this may indicate that the bore is too narrow and a larger restrainer is required.

3. Adjust head cover for body length of mouse. Adjust the head cover so that the mouse’s movements are minimized without causing unnecessary distress.

4. Check tail for evidence of injury (e.g., from fighting). If tail is significantly injured, consider using another mouse or testing the injured mouse at a later stage.

5. Habituate the mouse for 5 to 10 min, until breathing is normal and mouse is not agitated. A cover can be placed over the restrainer to help calm the animal, but this must be removed before the test so that the animal can be seen fully.

Testing 6. Place tail onto the pedestal of the Randall-Selitto apparatus and rest the blunt cone on top of the tail. Behavioral Measures of Pain Thresholds

Choose a point on the tail approximately one quarter of the way down from the base of the tail, so that the animal is able to withdraw the tail easily.

7. Apply pressure to the foot pedal to increase the weight exerted onto the tail.

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8. Observe the mouse closely. Release the foot pedal and lift the blunt cone at the first sign of struggling, vocalization, or withdrawal of the tail. The animal may not always be physically able to move the tail, so other pain behaviors should always be considered when deciding an endpoint.

9. Record the number reached on the scale and multiply by the weights to obtain the final force exerted on the tail. If the number is very close to the beginning or the end of the scale, consider adding or subtracting a weight to minimize errors from user reaction time or “maxing out.”

10. Repeat a maximum of three times in one session with at least 60 sec between stimuli. Be careful not to choose the exact same point on the tail each time.

Digital Paw Pressure Test A hand-held digital paw pressure analgesia instrument is also available. The protocol is the same as above (Basic Protocol 7), but the force is automatically calculated and there is no need to add or subtract additional weights.

ALTERNATE PROTOCOL 4

Materials Mice of optimal age (6 to 8 weeks) Mouse restrainer Digital Paw Pressure Analgesia Instrument (IITC Life Science, cat. no. 2500) Habituation 1. Place mouse into restrainer so that the head of the mouse is snug against the nose cone and the tail is extending out of the end plate. Choose the right size of restrainer—the mouse should be unable to turn around in the restrainer but not be uncomfortable. Time spent in the restrainer should be kept to <30 min, to minimize restraint-induced stress.

2. Habituate the mouse for 5 to 10 min until breathing is normal and mouse is not agitated. A cover can be placed over the restrainer to help calm the animal, but this must be removed before the test so the animal can be seen fully.

Testing 3. Open the pressure applicator and place the probe underneath the tail. Choose a point on the tail approximately one quarter of the way down from the base of the tail, so that the animal is able to withdraw the tail easily.

4. Close the pressure applicator of the digital paw pressure test instrument. Be careful to keep the pressure applicator horizontal, to maintain a consistent force.

5. Observe the mouse closely and release the pressure applicator at the first sign of struggling, vocalization, or withdrawal of the tail. The animal may not always be physically able to move the tail so other pain behaviors should always be considered when deciding an endpoint. A cut-off limit of 500 g should also be used to prevent tissue damage. The electronic unit will capture and store the peak force applied.

6. Repeat a maximum of three times in each session with at least 60 sec between stimuli. Be careful not to choose the exact same point on the tail each time.

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INFLAMMATORY PAIN BASIC PROTOCOL 8

Mechanical and/or Thermal Hyperalgesia in the Hindpaw This protocol describes the method for inducing inflammation in the hindpaw of a mouse. A number of agents can be used to induce inflammation of varying durations and severity, some of which are listed in Table 2. The method for injection remains the same for all of these agents. See Table 2 for specific information about the inflammatory agent and the subsequent behavioral measures and experiment time course, as these are not the same for all inflammatory agents.

Materials Mice of optimal age (6 to 8 weeks) Hamilton syringe and 29-G needle, or 0.5-ml disposable insulin syringe Inflammatory agent (see Table 2 for concentrations and dosage) Additional reagent and equipment for determining mechanical and thermal pain thresholds (see protocols above) Hindpaw injection 1. Restrain mouse gently by covering with tissue so the mouse is cupped under your hand securely. You should be able to access the paw and hold it firmly with the plantar surface facing upwards.

2. Insert the needle into the center of the paw at a shallow angle and inject the required dose of the inflammatory agent. This is an intraplantar injection—some agents may require intradermal injections, which are in the same location on the paw, but by inserting the needle almost parallel to the paw it is possible to inject just under the surface of the skin. The needle does not need to be inserted deeply, and the mouse should not bleed during or after injection.

Table 2 Examples of Inflammatory Agents and Suggested Doses

Inflammatory agent Concentration

Dose

Time course of inflammation

Type of injection

Type of hyperalgesia

Complete Freund’s adjuvant

100%

20 μl

More than 2 weeks

Intraplantar

Mechanical and thermal

2%

20 μl

30 min to up to 6 days

Intraplantar

Mechanical and thermal

100 ng

2.5 μl

Intradermal

Mechanical and thermal

Zymosan

0.2 mg/ml

20 μl

30 min to up to 6 days

Intraplantar

Mechanical and thermal

Formalin

0.5%-5%

20 μl

Acute phase: 0-10 min Inflammatory phase: 10-60 min

Intraplantar

Spontaneous

Bradykinin

1-10 nM

10-20 μl

0-60 min

Intraplantar

Spontaneous

Carrageenan PGE2

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Testing 3. After injection, record the mechanical and/or thermal thresholds of the inflamed paw using Hargreaves and/or von Frey protocols described in Basic Protocols 4 and/or 6 at the desired time points If desired, both paws can be injected; if not, the uninflamed paw can be used as an ipsilateral control. After the measurements are taken, the mice should be culled.

Spontaneous Inflammatory Pain Inflammation can also cause spontaneous pain in the form of flinching/licking biting of the affected area, usually the hindpaw. See Strategic Planning for more descriptions of pain behaviors.

ALTERNATE PROTOCOL 5

Materials Mice of optimal age (6 to 8 weeks) Inflammatory agent (see Table 2 for concentrations and dosage) Clear, lidded Perspex box, ∼15 cm × 15 cm Mirror Stopwatches Hamilton syringe and 29-G needle or 0.5-ml disposable insulin syringe Habituation 1. Habituate mouse to the Perspex box for 30 to 60 min or until exploratory behavior has ceased. It is useful to have two boxes available so that one mouse can be habituating while the first mouse is being observed.

Hindpaw injection 2. Remove the mouse from the box for injection. Restrain mouse gently by covering it with tissue so the mouse is cupped under your hand securely. You should be able to access the paw and hold it firmly with the plantar surface facing upwards.

3. Insert the needle into the center of the paw at a shallow angle and inject the required dose. This is an intraplantar injection—some agents may require intradermal injections, which are in the same location on the paw, but by inserting the needle almost parallel to the paw, it is possible to inject just under the surface of the skin. The needle does not need to be inserted deeply and the mouse should not bleed during or after injection.

Observation 4. After injection place the mouse into the Perspex box and use the stopwatch to time the amount of time spent conducting pain behaviors for the desired amount of time. A time course of the pain can be graphed if the pain behaviors are put into 5-min bins. The mirror should be used to ensure all views of the mouse can be seen and no behaviors are missed.

VISCERAL PAIN This protocol describes the steps required to produce viscera-specific behaviors in mice; this model causes both spontaneous pain and hyperalgesia. Common chemical irritants

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used are mustard oil and capsaicin, but this method would also be suitable to test more specific agonists/antagonists if required. BASIC PROTOCOL 9

Intracolonic Administration of Irritants Materials Mice of optimal age (6 to 8 weeks) Anesthetic (e.g., isoflurane) Isoflurane Oxygen Irritant (mustard oil or capsaicin) Catheter (0.6 mm diameter, ∼4 cm long; e.g., Harvard Apparatus, cat. no. 732851) 1-ml syringe Test compartment with mesh floor (e.g., IITC Life Science, cat. no. 410; http://www.iitcinc.com/) Habituation 1. Habituate mouse to the enclosure for 30 to 60 min or until exploratory behavior has ceased. It is useful to have two enclosures available so one mouse can be habituating while the first mouse is being observed.

Intracolonic administration 2. Remove the mouse from the enclosure, and, under brief anesthesia (using ∼3% isoflurane/oxygen mix), insert the catheter approximately 2.5 cm into the rectum. This ensures that the catheter has reached the colon; it can be useful to mark the catheter before insertion to make sure sufficient depth is achieved. Vaseline can also aid insertion and prevent topical contact of the perineal area with the irritant chemicals.

3. Administer 50 to 100 μl of irritant such as mustard oil (0.25 to 1%) or capsaicin (0.03% to 0.1%).

Observation 4. Replace mouse into box and observe behaviors (licking, stretching, and contractions of abdomen and squashing abdomen against the floor) for 20 min. The latency for appearance of first behavior can be recorded along with the numbers and types of behaviors. After the measurements have been taken, the mouse should be culled. SUPPORT PROTOCOL 3

Von Frey Hair Mechanical hypersensitivity after intracolonic administration of irritants can also be tested. The following protocol describes a method to test mechanical sensitivity of the abdomen using von Frey hairs. Referred hyperalgesia on other parts of the body such as the hindpaws may also be present in this model and can be tested using the protocol described in Basic Protocol 6.

Additional Materials (also see Basic Protocol 9) Calibrated set of von Frey hairs (e.g., Stoelting Touch Test Sensory Evaluator)

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Before intracolonic administration 1. Apply a range of von Frey hairs 10 times each in ascending order of force and note the number and intensity of responses. Each hair should be applied for 1 to 2 sec with an inter-stimulus interval of 5 to 10 sec. Do not stimulate the same point twice in succession, and avoid the external genitalia region. Current Protocols in Mouse Biology

2. Observe withdrawal response. A positive withdrawal response is either a sharp retraction of the abdomen, immediate licking or scratching of site of application of the von Frey hair, or jumping.

20 min after intracolonic administration 3. Repeat application of von Frey hairs as before. NEUROPATHIC PAIN: HYPERALGESIC ALLODYNIA MEDIATED BY PERIPHERAL NERVE DAMAGE Chronic Constriction Injury (CCI): Loose Ligation of the Sciatic Nerve Originally developed by Bennett and Xie in rats, CCI has also been shown to sensitize the affected limb to mechanical and thermal (both hot and cold) stimuli (Bennett and Xie, 1988; Bennett, 1994). The CCI model seems to involve a large immune-mediated component. The model involves tying sutures around the sciatic nerve (Fig. 4).

Figure 4

CCI and Seltzer diagram.

BASIC PROTOCOL 10

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Materials Mice of optimal age (6 to 8 weeks) Isoflurane Oxygen Iodine antiseptic solution Apparatus for administering isoflurane anesthesia, including muzzle for mouse Heat mat Surgical swabs Two size-5 forceps Size-15 scalpel 3-0 non-absorbable suture Suture needles and needle holders 3-0 absorbable suture Wound clips Wound clip remover Anesthetize mouse 1. Place mouse in induction chamber. Fill chamber with ∼3% isoflurane/oxygen mix. Exact isoflurane concentration will depend on exact equipment being used. It is recommended that you consult the manual and optimize the isoflurane/oxygen mix beforehand.

2. Once mouse is unconscious, transfer to anesthetic muzzle with∼1% isoflurane/oxygen mix. Place the mouse on a heat mat to maintain body temperature during surgery. Again, isoflurane concentration will depend on exact equipment being used. It is recommended that you consult the manual and optimize the isoflurane/oxygen mix beforehand.

3. Check the mouse’s withdrawal reflex by pinching the hindpaw with forceps. If mouse withdraws paw, wait 30 sec and check again. If after three attempts, the mouse still withdraws, adjust isoflurane concentration. If the isoflurane oxygen mix is too high this can be lethal. So any adjustments should be made gradually.

4. Once withdrawal reflex is no longer seen, proceed with surgery. However, anesthetic depth should be monitored throughout surgery. This can be achieved through monitoring breathing rate and depth.

Surgery 5. Clean the shaved skin over the dorsal side designated for surgery with an iodine antiseptic solution (or equivalent). This area should extend from the midline to the knee, with a width of at least 15 mm.

6. Make a 0.15-mm incision in the skin along the length of the femur. 7. Break through the biceps femoris using blunt dissection to expose the sciatic nerve. The sciatic nerve lies just beneath the biceps femoris. Using two sharp size-5 forceps to expose and gently separate the muscle avoids the risk of nicking the sciatic nerve. IMPORTANT NOTE: For sham surgery, proceed directly to step 12. DO NOT perform steps 8 to 11.

Behavioral Measures of Pain Thresholds

8. Carefully separate a 6-mm section of the sciatic nerve of from the surrounding tissue using blunt dissection. 9. Gently slide a closed forceps under the sciatic nerve and lift enough to pass the suture needle under the nerve.

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10. Pull the non-absorbable suture through, leaving 10 mm, protecting the sciatic nerve by lifting with forceps to avoid friction damage. 11. Loosely tie the non-absorbable suture around the nerve, trimming any excess sutures. Repeat previous step three times, leaving 1 mm between each ligature. Bennett and Xie (1988) state that ligatures should be tied such that the diameter of the nerve should just barely be constricted when viewed with 40× magnification.

Wound closure 12. Close the biceps femoris with a 3-0 absorbable suture, if necessary. 13. Bring together the two sides of the incision, making sure both sides are aligned. Close with three surgical wound clips, or the minimum number of clips possible. Avoid putting the wound clips too close to the edge of the incision as these may rip out. Concomitantly, avoid putting the clips too ‘deep’ and bunching up the skin, as this may cause irritation and distress to the mice.

Recovery 14. Remove the anesthetic muzzle and transfer to a recovery cage. It is important that mice be allowed to recover in isolation before being returned to their original littermates in their original cage.

15. Monitor mice until they regain consciousness. It is recommended that a second person monitor the recovery of the previous mouse while the first person continues performing surgery with the remaining mice.

Post-operative care 24 hr after surgery 16. Check that none of the wound clips have fallen out and that the incision has not reopened. Apply additional wound clips where necessary. 17. Inspect the mice for any signs of infection, lameness, or other unexpected adverse side effects. Treat the mice if possible; otherwise it may be necessary to cull mice with severe adverse side effects.

7 days after surgery 18. Remove any remaining wound clips and inspect for signs of infection. Treat the mice if possible. Pain tests 19. Allow mice 48 hr to recover before undergoing any behavioral measures of nociception. The time taken for allodynia and hyperalgesia-like responses to develop can vary. Therefore, it may be useful to test the mice on the 3rd, 5th, and 7th day following surgery. Following this, less frequent measures are necessary, i.e., once or twice per week. The effects of surgery can be detected up to 60 days (or more) following surgery.

Seltzer: Partial Ligation of the Sciatic Nerve Originally developed by Seltzer using rats, partial ligation of the sciatic nerve has also been shown to sensitize the affected limb to mechanical and thermal (both hot and cold) stimuli (Seltzer et al., 1990). Of the listed models included in this unit, it is practically the most simple to perform. The model involves tying a suture through half the sciatic nerve (Fig. 4).

ALTERNATE PROTOCOL 6

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Additional Materials (also see Basic Protocol 10) Dissection microscope Anesthesia 1. Anesthetize mouse as described in steps 1 to 4 of Basic Protocol 10. Surgery 2. Clean the shaved skin over the dorsal side designated for surgery with an iodine antiseptic solution (or equivalent). This area should extend from midline to the knee, with a width of at least 15 mm.

3. Make a15-mm incision in the skin along the length of the femur. 4. Break through the biceps femoris using blunt dissection to expose the sciatic nerve. The sciatic nerve lies just beneath the biceps femoris. Using two sharp size-5 forceps to expose gently separate the muscle avoids the risk of nicking the sciatic nerve. IMPORTANT NOTE: For sham surgery, proceed directly to step 9. DO NOT perform steps 5 to 8.

5. Carefully separate an ∼3 mm section of the sciatic nerve from surrounding tissue using blunt dissection. 6. Gently slide a closed forceps under the sciatic nerve and lift enough to pass the suture needle under the nerve. 7. Use a dissection microscope to insert the needle one-third to one-half way through the thickness of the sciatic nerve. 8. Loosely tie the suture around the nerve, trimming any excess suture. Be careful not to pull the suture too tight, thereby cutting the nerve.

Wound closure 9. Close wounds with wound clips as described in steps 12 to 13 of Basic Protocol 10. Recovery 10. Allow animals to recover as described in steps 14 to 15 of Basic Protocol 10. Post-operative care 11. Care for mice post-operatively as in steps 16 to 18 of Basic Protocol 10. Pain tests 12. Allow mice 48 hr to recover before undergoing any behavioral measures of nociception. The time taken for allodynia- and hyperalgesia-like responses to develop can vary. Therefore, it may be useful to test the mice on the 3rd, 5th and 7th day following surgery. Following this, less frequent measures are necessary—i.e., once or twice per week. The effects of surgery can be detected up to 60 days (or more) following surgery. ALTERNATE PROTOCOL 7

Behavioral Measures of Pain Thresholds

Spinal Nerve Ligation (SNL) For almost 20 years, the Kim and Chung spinal nerve ligation model has been widely used in both rats and mice. SNL has been shown to sensitize the affected limb to mechanical and thermal (both hot and cold) stimuli (Kim and Chung, 1992). However, its widespread use has led to a number of slight variations, such as tight ligation of L5 or L5 and L6, as well as axotomy of one or both of L5 and L6. These differences are likely to account for some phenotypic difference and should be taken into account when comparing results to published findings. The protocol below specifically describes L5 axotomy (Fig. 5).

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Figure 5

SNL diagram.

Additional Materials (also see Basic Protocol 10) Surgical swabs Retractor, glass hook: surgical hooks are commercially available; however these has proven awkward to use and can result in damage to the other sciatic nerve roots—using a Bunsen burner to melt and draw out glass Pasteur pipets into fine hooks has proven a more useful tool (Fig. 6) Dissection microscope Size-7 forceps Size-11 scalpel Micro-scissors Anesthetic 1. Anesthetize mouse as described in steps 1 to 4 of Basic Protocol 10. Behavioral Measures of Pain Thresholds

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Figure 6

Glass hook for surgery. Scale bar = 1 mm.

Surgery 2. Clean the shaved skin over the dorsal side designated for surgery with an iodine antiseptic solution (or equivalent) and place in the prone position. This area should extend along the midline from second lumbar vertebrae (L2) and second sacral vertebra (S1), with a width of at least 15 mm.

3. Make a 15-mm longitudinal incision in the skin along the spinal column, from L3 to S1. 4. Use the forefinger and thumb to gently locate the iliac crests. Starting ∼2 mm below the iliac crests make a 7-mm incision in the paraspinal muscle ∼2 mm from the center of the spinal column, ∼3 mm deep. Be careful not to extend the incision past L4, as this can increase the risk of the incision rapidly filling with blood, thereby obstructing the view of the spinal nerves. If the incision causes a bleed, insert a swab into the incision and gently apply pressure in the direction of the bleed source (it may be necessary to hold this in place for up to 60 sec). When the incision is in the correct location, you should be able to feel slight ‘bumps’ as the blade runs along the transverse processes. IMPORTANT NOTE: Sham surgery: move directly to step 12; DO NOT perform steps 5 to 11.

5. Carefully insert the retractor into the incision and gently expand the opening. Behavioral Measures of Pain Thresholds

Be careful not to insert the retractor between the iliac crest and the spinal column, since opening the retractor in this case may fracture the hip.

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6. Using a dissection microscope, gently feel for the L5 transverse processes using closed forceps. Once located, use two size-5 forceps to separate the muscle to reveal the bone. 7. Once the L5 transverse process is exposed, use a size-11 scalpel to gently score the processes as close to the base as possible. Then, using the original cut as a guide, repeatedly score the bone while gently applying pressure until the bone becomes detached. It may be necessary to remove bone splinters, using size-7 forceps, from the stump of the L5 transverse process.

8. Firmly grip the ligand attached to the tip of the L5 transverse process, and cut with micro-scissors, allowing the transverse process to be removed. The L5 branch of the sciatic nerve should be immediately below the recently removed L5 transverse process.

9. Use the glass hook of the retractor to separate the nerve from the surrounding tissue. 10. Slide the glass hook under the L5 nerve and gently lift away from the surrounding tissue. Use the micro-scissors to cut the nerve. 11. Grip the distal section of the cut nerve with the size-5 forceps and cut off a 1- to 2-mm section.

Wound closure 12. Close the paraspinal muscle with two 3-0 absorbable sutures. 13. Bring together the two sides of the incision, making sure both sides are aligned. Close with four surgical wound clips (or the minimum number of possible).

Recovery 14. Allow mice to recover as described in steps 14 to 15 of Basic Protocol 10. Post-operative care 24 hr after surgery 15. Check that none of the wound clips have fallen out and that the incision has not reopened. Apply additional wound clips where necessary. 16. Inspect the mice for any signs of infection, lameness, or other unexpected adverse side effects. Treat the mice if possible; otherwise, it may be necessary to cull mice with severe adverse side effects. Dragging of the operated side is indicative of damage to the L4 spinal nerve, since it innervates a number of the hind-limb muscles. These mice should be removed since the nociceptive measures require paw withdrawal as a cutoff point.

7 days after surgery 17. Remove any remaining wound clips and inspect for signs of infection. Treat the mice if possible. Pain tests 18. Allow mice 48 hr to recover before undergoing any behavioral measures of nociception. The time taken for allodynia and hyperalgesia-like responses to develop can vary. Therefore it may be useful to test the mice on the 3rd, 5th, and 7th day following surgery. Following this, less frequent measures are necessary, i.e., once or twice per week. The effects of surgery can be detected up to 60 days (or more) following surgery.

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COMMENTARY Background Information Thermal thresholds Heat The Hargreaves test is a recent addition to the arsenal of thermal nociception measures. The test paradigm has some advantages over other commonly used thermal nociception measures, namely the tail flick (D’Amour and Smith, 1941) and the hot plate (Macdonald et al., 1946). The main strengths of the test paradigm are: (1) the mice are unrestrained, minimizing stress-related analgesia; (2) a clear (in some cases automated) and quantifiable outcome measure; (3) repeated testing within a single session is not associated with the induction of hyperalgesia; (4) laterality, enabling within-subject controls. However, there seems to be some disparity between the thermal stimuli in the various tests described above. Chapman et al. (1985) among others suggest that the tail-flick represents a spinal reflex, while the hot plate involves supraspinal processing. Therefore, experimental conditions that affect one and not the other can be considered spinal or supraspinal, respectively. However, it seems a number of other factors may also contribute to differences between the two tests. Cold Abrahamsen et al. (2008) demonstrate that noxious heat and cold are very separate pain modalities. Ablating peripheral neurons expressing SCN10A leads to loss of nociceptive behavior to cold (0◦ C), while nociceptive behavior to heat remains intact (both at 50◦ and 55◦ C). Single-fiber recordings further highlight that the sensory neuron sodium channel Nav1.8 is essential for nociception in the cold and therefore cold pain (Zimmermann et al., 2007). Therefore it is critical to use separate assays for both noxious heat and cold to fully characterize thermal nociception. The temperature border for noxious cold is less clear; however, it can generally be considered that <0◦ C is noxious. By this definition, the acetone test does not induce cold pain when applied to a na¨ıve mouse, since it only causes a decrease of ∼5◦ C. However, it is listed, since it can detect cold allodynia associated with some neuropathic pain models. Mechanical thresholds Behavioral Measures of Pain Thresholds

von Frey Touch thresholds in humans and animals have been determined using von Frey hairs

since the 1890s. The calibrated monofilaments are most commonly made of nylon and apply a consistent, accurate force when used correctly. The up-down method for obtaining the 50% threshold using von Frey hairs was described by Chaplan et al. (1994). This method makes use of the statistical formula described by Dixon (1980) to determine the LD50 . The advantage of the up-down method is reduced applications of von Frey hairs while maintaining the same level of accuracy. Randall-Selitto The Randall-Selitto test was developed by L.O. Randall and J.J. Selitto in 1957 as a way of measuring efficacy of analgesics on inflamed tissue (Randall and Selitto, 1957). The test was originally applied to a rat’s paw but has been modified for use on mice using the tail. It is possible to use the Randall-Selitto test on a mouse paw, but the level of restraint required for this increases stress levels and makes the pain behaviors difficult to observe. Instead, mechanical sensitivity on the mouse paw should be measured using the von Frey method Inflammatory models Complete Freund’s adjuvant (CFA) CFA has been used as a reliable model of persistent inflammatory hyperalgesia since the 1980s (Larson et al., 1986). The thermal and mechanical hyperalgesia observed is linked to TNF-α and cytokine release in the periphery (Cunha et al., 1992), develops approximately 24 hr after injection, and remains for up to 2 weeks. Carrageenan The injection of carrageenan into the hindpaw was originally developed as a model of inflammation for the screening of antiinflammatory drugs (Crunkhorn and Meacock, 1971). The edema develops over 6 hr and, as with CFA, is linked to activation of cytokine cascades (Cunha et al., 1992). ProstaglandinE2 (PGE2) PGE2 is synthesized from arachidonic acid by COX-2 and PGE synthase enzymes, a pathway which is the major site of action for NSAIDs. Intraplantar administration of PGE2 causes a short-lasting mechanical hyperalgesia by acting directly on primary afferent nociceptors causing sensitization (Taiwo et al., 1987; Southall and Vasko, 2000).

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Formalin The formalin test allows assessment of continuous pain, lasting ∼1 hr in the absence of evoked stimuli. The behaviors occur in two distinct phases. The first phase starts immediately after injection and lasts 5 to 10 min, and is due to direct chemical stimulation of C fibers acting through TRPA1 (McNamara et al., 2007). The second phase starts around 15 to 20 min after injection and lasts for 20 to 40 min. This phase involves central changes within the spinal cord and inflammatory molecules (Coderre et al., 1990). The concentration of formalin injected can be altered to study the two phases—a low concentration (0.5% to 1%) is sufficient to study the early phase, whereas concentrations of over 1% are required to produce the second phase of pain behaviors (Rosland et al., 1990). The concentration should be kept as low as possible to avoid unnecessary suffering. Bradykinin (BK) Bradykinin is a major inflammatory mediator that produces severe spontaneous pain and mechanical and thermal hyperalgesia (Couture et al., 2001). Hyperalgesia is caused by sensitization of sensory ion channels such as TRPV1 and TRPA1 (Chuang et al., 2001;Wang et al., 2008). Zymosan Zymosan is a glucan that forms the cell wall of Saccharomyces cerevisiae, which produces thermal and mechanical hyperalgesia along with edema and spontaneous pain (Meller and Gebhart, 1997). Visceral pain model This intracolonic stimulation model was developed by Laird et al. (2001) to provide a way to obtain information on visceral pain, hyperalgesia, and colon inflammation from the same animal (Laird et al., 2001). The extent of spontaneous pain behaviors is concentration dependent, with 1% mustard oil and 0.1% capsaicin giving the peak behavioral responses. However, high concentrations can also produce ‘freezing’ in-between the pain behaviors. Mechanical hyperalgesia may only be seen at the higher concentrations of irritant. Neuropathic pain model Although the mechanisms underlying neuropathic pain are currently not fully understood, a number of factors can lead to nerve injury and, in turn, neuropathic pain—for example, diabetes, herpes zoster, nerve compression, channelopathies, and autoimmune dis-

ease. Hence, there are a large variety of neuropathic pain models that address these different etiologies. Similarly, neuropathic pain can also result from central sensitization mechanisms. Information regarding central pain and the relevant animals models can be found in Bennett (1994). However, this unit focuses on three commonly used models of surgically induced peripheral nerve damage.

Critical Parameters Operator influence As with many behavioral measures, the operator’s scoring criteria can vary and result in variation with respect to the actual outcome score. Therefore, the same operator, when conducting multiple repetitions on the same test group (e.g., before and after treatment), can counteract this. It seems that an important factor in operator influence is animal-handling technique (Chesler et al., 2002), which possibly exerts its effect by inducing varying degrees of stress. On the other hand, it seems that operator age, sex, and experience level play an insignificant role. Test environment The influence of the test environment on behavioral outcome measures has previously been shown (Chesler et al., 2002). These factors include cage density, humidity, lighting, etc., as well as non-physical factors such as time of day. Careful monitoring of these factors in order to keep them consistent may minimize their influence. However, if the test subject mice fail to settle after the habituation period, a number of measures can be taken: a. Reducing light levels to a minimum and ensuring that it is consistent for all subject mice. b. Covering transparent test compartments so that operator is not in the direct line of sight. c. Checking background ultrasound levels, as this may cause distress to mice but without being audible to test operator. d. Playing low-level white noise may mask operator and/or external noises that may alarm the mice. e. Using unscented cleaning products and minimizing exposure to novel or threatening smells (e.g., operator wearing a new perfume or animal scents, such as rat). f. Checking room temperature and environment: either too hot or cold can alter behavioral response or habituation. Similarly, high levels of humidity can also be detrimental. g. Carrying out tests at same time of day, since activity levels can vary during a working

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Table 3 Troubleshooting Summary

Problem

Possible cause

Solution

One mouse will not settle

Various

Remove mouse from experiment and continue with the remaining eleven

All mice will not settle

Multiple possible causes, e.g., ultrasonic The test environment may be noise, high intensity light, smells, time of day unsuitable. Find alternative location.

All mice reach cut-off point Apparatus faulty or incorrectly calibrated

Refer to apparatus manual

Table 4 Approximate Anticipated Results for Acute Pain Tests

Mouse strain

129sv

BALB/c

C57BL/6

DBA/2

von Frey hairs

0.8 g

0.8 g

1-2 g

1g

Automatic von Frey

9-10 g

8-9 g

9-10 g

8-9 g

100-120 g

150-200 g

20-25 sec

30-35 sec

15-20 sec

25-30 sec

15-20 sec

20-25 sec

5-10 sec

10-15 sec

10 sec

15 sec

10 sec

10 sec

Randall-Sellitto ◦

Hot plate (53 C) ◦

Cold plate (15 C) Hargreaves

day. Also, mice are nocturnal, and therefore it may be necessary to house them in a room with an inverted dark-light cycle if specifically investigating behavior during the active phase. Alternatively, see Mogil et al. (2001) and R´acz and Zimmer (2006) for further discussion. Strain-specific variation As with humans, mice display highly variable nociceptive responses to identical stimuli, injuries, or pathologies. Mogil et al. (1999) systematically surveyed the relative sensitivity of 11 inbred mouse strains using 12 different behavioral measures of nociception and reported that performance in all of the strains varied greatly. This highlights the importance of using appropriate littermate controls, as well as establishing the baseline response for any novel mouse strain. When investigating a new strain or behavioral paradigm, it is advisable to establish the baseline response for your particular setup, even if the strain or behavioral paradigm is commonly used. This will ensure that your apparatus is calibrated, that your test environment is suitable, and that your test operator is confident with the test paradigm.

Behavioral Measures of Pain Thresholds

Inter-stimulus interval Slugg et al. (2000) demonstrate that C fibers undergo significant fatigue in response when the inter-stimulus interval between the paired stimuli was <150 sec (Slugg et al., 2000). On the other hand, A fibers did not demonstrate a significant fatigue until the inter-stimulus

interval was <30 sec in monkeys. This can be used as guideline for determining interstimulus interval for the above tests. Sample size When comparing two groups, it recommended to start with at least six animals per group. However, it may be necessary to increase this number.

Troubleshooting A list of common issues and solutions is provided in Table 3. If inflammation fails to develop, first check that the agents have been stored and made up correctly. CFA needs to be shaken vigorously before administration to emulsify. Also, practice injection technique to ensure the full dose of inflammatory agent is being administered.

Anticipated Results Test results can be influenced by the operator and environment (Chesler et al., 2002) as well as mouse strain (Mogil et al., 1999). Therefore, it is recommended to perform a literature search for similar experimental conditions. Example results for wild-type C57BL/6, 129sv and BALB/c mouse strains are listed in Table 4. Mechanical hyperalgesia At the peak of inflammation, the 50% threshold measured by von Frey hairs will often be reduced to absolute minimum (a

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response given at the lowest weight hair used). The threshold measured by an automatic von Frey should drop by more than 50%. The thresholds will slowly improve back to baseline depending on the duration of inflammation. Thermal hyperalgesia At the peak of inflammation, latency should drop by more than 50% and slowly improve back to baseline depending on the duration of inflammation. Spontaneous pain Spontaneous pain behaviors have a large amount of subjectivity, which can make it difficult to predict absolute numbers. Consistency is more important, and in the case of the formalin test, the plotted data should show a biphasic response (>1% formalin concentrations).

Time Considerations Habituation Habituation of the animals for 30 min to 2 hr as stated in the individual protocols is essential. The following estimates of time considerations exclude this habituation time, so this should be added on accordingly. Mechanical and thermal threshold determination The majority of commercial enclosures for the Hargreaves’ test, von Frey test, and acetone test allow habituation and testing of up to 12 animals in one session. For each of these tests, a minimum of 1.0 to 1.5 hr should be allotted for 12 animals. The hot plate, cold plate, tail-flick, and Randall-Selitto tests need to be carried out individually, and will take around 10 min per mouse per temperature, including handling time. Inflammatory and visceral pain models With practice, the injection or intracolonic administration of the chemical or irritant should take no more than 5 min per mouse. Surgery It is preferable to have two people for the surgical procedures, one to anesthetize and perform the nerve ligation and one for wound repair and recovery. With two people, an entire day should be allotted to perform surgery on 12 to 18 animals. The time needed for each surgery will depend greatly on the level of expertise.

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Dixon, W.J. 1980. Efficient analysis of experimental observations. Annu. Rev. Pharmacol. Toxicol. 20:441-462. Eddy, N.B. and Leimbach, D. 1953. Synthetic analgesics. II. Dithienylbutenyl- and dithienylbutylamines. J. Pharmacol. Exp. Ther. 107:385-393.

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Gray, J.A . 1978. The neuropsychology of anxiety. Br. J. Psychol. 69:417-434.

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Hargreaves, K., Dubner, R., Brown, F., Flores, C., and Joris, J. 1988. A new and sensitive method for measuring thermal nociception in cutaneous hyperalgesia. Pain 32:77-88.

Randall, L.O. and Selitto, J.J. 1957. A method for measurement of analgesic activity on inflamed tissue. Arch. Int. Pharmacodyn. Ther. 111:409419.

Kim, S.H. and Chung, J.M. 1992. An experimental model for peripheral neuropathy produced by segmental spinal nerve ligation in the rat. Pain 50:355-363.

Rosland, J.H., Tjlsen, A., M¨uhle, B.R., and Hole, K. 1990. The formalin test in mice: Effect of formalin concentration. Pain 42:235242.

Laird, J.M.A., Martinez-Caro, L., Garcia-Nicas, E., and Cervero, F. 2001. A new model of visceral pain and referred hyperalgesia in the mouse. Pain 92:335-342.

Seltzer, Z., Dubner, R., and Shir, Y. 1990. A novel behavioral model of neuropathic pain disorders produced in rats by partial sciatic nerve injury. Pain 43:205-218.

Larson, A.A., Brown, D.R., el-Atrash, S., and Walser, M.M. 1986. Pain threshold changes in adjuvant-induced inflammation: A possible model of chronic pain in the mouse. Pharmacol. Biochem. Behav. 24:49-53.

Slugg, R.M., Meyer, R.A., and Campbell, J.N. 2000. Response of cutaneous A- and C-fiber nociceptors in the monkey to controlled-force stimuli. J. Neurophysiol. 83:2179-2191.

Lee, D.E., Kim, S.J., and Zhuo, M. 1999. Comparison of behavioral responses to noxious cold and heat in mice. Brain Res. 845:117-121. Macdonald, A.D., Woolfe, G., Bergel, F., Morrison, A.L., and Rinderknecht, H. 1946. Analgesic action of pethidine derivatives and related compounds. Br. J. Pharmacol. Chemother. 1:4-14. McNamara, C.R., Mandel-Brehm, J., Bautista, D.M., Siemens, J., Deranian, K.L., Zhao, M., Hayward, N.J., Chong, J.A., Julius, D., Moran, M.M., and Fanger, C.M. 2007. TRPA1 mediates formalin-induced pain. Proc. Natl. Acad. Sci. U.S.A. 104:13525-13530. Meller, S.T. and Gebhart, G.F. 1997. Intraplantar zymosan as a reliable, quantifiable model of thermal and mechanical hyperalgesia in the rat. Eur. J. Pain 1:43-52. Mogil, J.S., Wilson, S.G., Bon, K., Lee, S.E., Chung, K., Raber, P., Pieper, J.O., Hain, H.S., Belknap, J.K., Hubert, L., Elmer, G.I., Chung, J.M., and Devor, M . 1999. Heritability of nociception I: responses of 11 inbred mouse strains on 12 measures of nociception. Pain 80:67-82. Mogil, J., Wilson, S., and Wan, Y. 2001. Assessing nociception in murine subjects. In Methods in Pain Research (L. Kruger, ed.) pp. 11-40. CRC Press, Boca Raton, Fla.

Southall, M.D. and Vasko, M.R. 2000. Prostaglandin E(2)-mediated sensitization of rat sensory neurons is not altered by nerve growth factor. Neurosci. Lett. 287:33-36. Taiwo, Y.O., Goetzl, E.J., and Levine, J.D. 1987. Hyperalgesia onset latency suggests a hierarchy of action. Brain Res. 423:333-337. Takesue, E.I., Schaefer, W., and Jukniewicz, E. 1969. Modification of the Randall-Selitto analgesic apparatus. J. Pharm. Pharmacol. 21:788789. Vetulani, J., Castellano, C., Las´on, W., and Oliverio, A. 1988. The difference in the tail-flick but not hot-plate response latency between C57BL/6 and DBA/2J mice. Pol. J. Pharmacol. Pharm. 40:381-385. Wang, S., Dai, Y., Fukuoka, T., Yamanaka, H., Kobayashi, K., Obata, K., Cui, X., Tominaga, M., and Noguchi, K. 2008. Phospholipase C and protein kinase A mediate bradykinin sensitization of TRPA1: A molecular mechanism of inflammatory pain. Brain 131:12411251. Zimmermann, K., Leffler, A., Babes, A., Cendan, C.M., Carr, R.W., Kobayashi, J., Nau, C., Wood, J.N., and Reeh, P.W. 2007. Sensory neuron sodium channel Nav1.8 is essential for pain at low temperatures. Nature 447:855-858.

Behavioral Measures of Pain Thresholds

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Characterization of Whole Body Cholesterol Fluxes in the Mouse Gemma Brufau1 and Albert K. Groen1,2 1

Department of Pediatrics, Center for Liver, Digestive, and Metabolic Diseases, University Medical Center Groningen, University of Groningen, Groningen, The Netherlands 2 Department of Laboratory Medicine, Center for Liver, Digestive, and Metabolic Diseases, University Medical Center Groningen, University of Groningen, Groningen, The Netherlands

ABSTRACT Atherosclerosis is characterized by excessive cholesterol accumulation in the vessel wall. Current therapies mainly aim at decreasing influx through lowering plasma LDLcholesterol levels. The challenge is to develop therapeutic interventions to increase efflux of excess cholesterol from the vessel wall. The pathway that mediates this efflux from vessel wall to final excretion in the feces is called reverse cholesterol transport. Recently, it has become apparent that the intestine plays an important regulatory role in this pathway. This article describes in detail a variety of experimental approaches to measure cholesterol fluxes in the hepatobiliary system as well as in the intestinal pathway. Curr. C 2011 by John Wiley & Sons, Inc. Protoc. Mouse Biol. 1:413-427  Keywords: mouse r fecal cholesterol excretion r biliary excretion r transintestinal cholesterol r excretion

INTRODUCTION Cardiovascular disease remains a major cause of mortality and morbidity in developed societies. Atherosclerosis is the hallmark of cardiovascular disease and is mainly caused by accumulation of cholesterol in the vessel wall. A typical western-type diet accelerates the onset of atherosclerosis, which progresses through life (Keys et al., 1981). Cholesterol accumulation is a continuous process inducing a positive whole-body cholesterol balance—i.e., intake and de novo synthesis of cholesterol are higher than excretion. The etiology underlying this disturbed cholesterol balance is still largely unknown. Since it is difficult to study the distribution of body cholesterol fluxes in humans, animal models are employed to obtain a better understanding of the systems involved in regulation of body cholesterol excretion. For the calculation of cholesterol excretion via the intestine, several fluxes must be considered. The input fluxes of cholesterol into the body are dietary cholesterol intake and fractional cholesterol absorption. The output fluxes are the daily biliary cholesterol secretion, flux of cholesterol excreted by direct transport from the blood to intestine pathway, and appearance of neutral sterols (cholesterol and its metabolites) in the feces. The calculation of these different fluxes will allow investigation of pharmacological/dietary treatments that aim to enhance the body cholesterol efflux and to decrease the input of cholesterol in the body. In the following sections, the different methodologies available to measure these fluxes are described: Basic Protocol 1 describes the measurement of dietary cholesterol intake and neutral sterol excretion; Basic Protocol 2 describes the measurement of biliary cholesterol excretion; Basic Protocol 3 describes the direct measurement of fractional cholesterol absorption by observation of lymphatic cholesterol transport; and, finally,

Current Protocols in Mouse Biology 1: 413-427, December 2011 Published online December 2011 in Wiley Online Library (wileyonlinelibrary.com). DOI: 10.1002/9780470942390.mo110118 C 2011 John Wiley & Sons, Inc. Copyright 

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Basic Protocol 4 describes how to estimate the transintestinal cholesterol excretion using the sterol balance method. NOTE: All protocols using living animals must first be reviewed and approved by an Institutional Animal Care and Use Committee (IACUC) or must conform to governmental regulations regarding the care and use of laboratory animals. BASIC PROTOCOL 1

MEASUREMENT OF DIETARY CHOLESTEROL INTAKE AND NEUTRAL STEROL EXCRETION The correct calculation of cholesterol intake depends on the correct measurement of food intake (FI), which directly reflects the amount of cholesterol ingested, and the correct measurement of fecal excretion (FE), which directly reflects the amount of sterol excreted per period of time. Therefore, these two parameters should be evaluated, always using the same housing conditions (e.g., type of bedding, presence of enrichment in the cages).

Materials Appropriate mouse strain Mouse diet Control feces (sample that has been repeatedly analyzed and has given consistent results) Standards: 5α-cholestane, coprostanol, epi-coprostanol, cholesterol, dihydro-cholesterol, coprostan-3-one and 3-keto-cholesterol (Sigma) Absolute ethanol 1 M sodium hydroxide (NaOH; Sigma) Methanol (Merck) Petroleum ether, boiling point range 60◦ to 80◦ C (Merck) Pyridine (Thermo Scientific) N,O-Bis(trimethylsilyl)trifluoroacetamide (BSTFA; Sigma) Trimethylchlorosilane (TMCS; Thermo Scientific) Nitrogen source Hexane (Merck) Single-housing cages for mice Scale with 0.1 g accuracy (to weigh food and mice) Container to accommodate mice during weighing Separate mortar and pestle for grinding the feces and the food Analytical balance with 0.1 mg accuracy 10-ml glass screw-top vials 80◦ and 40◦ C heat blocks GC vials Gas chromatographic (GC) system (van der Veen et al., 2009) Additional reagents and equipment for injection of mice (Donovan and Brown, 2006a) Initial sample collection and preparation 1. At a given hour of the day, change mice to clean cages and weigh the amount of food given. Define in advance the time when the cages will be changed, since future animal handling must be done at the same hour.

Whole Body Cholesterol Fluxes in the Mouse

2. Three days later, at the same time of the day, Accurately weigh the remaining food, including any chunks of pellets in the cage, using an analytical balance. At the same time, collect the bedding plus the feces, then separate out all of the pieces of feces. It is very important to collect all pieces of feces in order to have a good estimation of the fecal output).

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Table 1 Calibration Curve to Measure Neutral Sterols and Cholesterol in Feces and Diet, Respectively

5αEpiCoprostanol Cholesterol cholestane coprostanol

Dihydrocholesterol

Coprostan3-one

3-ketocholesterol

nmol

nmol

nmol

nmol

nmol

nmol

nmol

Standard 1

50

0

0

0

0

0

0

Standard 2

50

10

10

10

10

10

10

Standard 3

50

25

25

25

25

25

25

Standard 4

50

50

50

50

50

50

50

Standard 5

50

100

100

100

100

100

100

3. Accurately weigh the feces using an analytical balance and keep them in an open glass container at 21◦ C to dry. Weigh the feces daily until the weight does not vary. At this time point, we consider that the feces to be dry. Alternatively, the feces can be lyophilized.

4. Grind the feces and the food separately in order to obtain a powder. Feces and food can be ground once they are dry at 21◦ C. The final powders can be kept at 21◦ C in a closed glass container.

Preparation of samples for analysis 5. Prepare samples for measurement of the content of cholesterol (diet) or neutral sterols (feces) by gas-chromatography (GC) (van der Veen et al., 2009). Weigh around 50 mg of feces or diet into a 10-ml glass screw-top tube containing 50 nmol of 5α-cholestane (internal standard) dissolved in absolute ethanol (0.5 μmol/ml concentration;100 μl volume). Record the exact weight of the feces for future calculations. Include one control feces, which will be analyzed every time that this protocol will be performed.

6. Add 2 ml of a 1:3 (v/v) mixture of 1 M NaOH and methanol to all the samples and keep the samples closed at 80◦ C for 2 hr. 7. Cool down the samples and subsequently add 3 ml petroleum ether. 8. Mix every sample for at least 30 sec, and centrifuge 10 min at 1764 × g, 21◦ C. 9. Pipet the upper layer in a new glass tube and repeat the same extraction procedure two more times. 10. Collect the three extraction aliquots and evaporate them at 40◦ C under nitrogen flow. In parallel, prepare a calibration curve according to the scheme in Table 1. 11. Once all the samples and standards are dry, add 100 μl of a solution of pyridine:BSTFA:TMCS (5:5:0.1; v/v/v) and cap the tubes. Keep the tubes closed at 21◦ C for at least 1 hr (this procedure can also be performed overnight). 12. Evaporate the reagent at 21◦ C under the flow of nitrogen. Immediately after the tubes are dry, add 1 ml of hexane. Mix and pour the content into a GC vial. These samples are ready to be injected into the GC as previously described (van der Veen et al., 2009)

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Calculations and analysis of data 13. Calculate the dietary cholesterol (DC) in g/mouse/day according to the formula:

DC( g mouse day ) =

w0 - wnd •Cdiet nd

Equation 1

where w0 is the weight (g) of the amount of food given, wnd is the weight (g) of the food remaining in the cage at the end of the feeding period, nd is the number of days over which FI is calculated (normally it is 3 days), and Cdiet is the concentration of cholesterol in the diet (μmol/g). 14. Calculate the neutral sterol secretion (NSS) in g/mouse/day according to the formula:

NSS (g mouse day) =

fend • NSfeces nd

Equation 2

where fend is the weight (g) of feces produced in the period, nd is the number of days over which FI is calculated (normally 3 days), and NSfeces is the concentration of neutral sterols in the feces (μmol/g). BASIC PROTOCOL 2

MEASUREMENT OF BILIARY CHOLESTEROL EXCRETION The reliability of the determination of biliary cholesterol secretion depends on several factors: (i) the ability of the technician to cannulate the gall bladder; (ii) the time of day chosen to perform the bile cannulations; (iii) the duration of bile collection; and (iv) the body temperature of the mice during bile collection. In this protocol, the most optimal conditions are described, as well as the limitations of the measurement.

Materials Appropriate mouse strain 0.315 mg/ml fentanyl citrate 10 mg/ml fluanisone 5 mg/ml diazepam Phosphate-buffered saline (PBS) 70% (v/v) ethanol Standards: 5α-cholestane, coprostanol, epi-coprostanol, cholesterol, dihydro-cholesterol, coprostan-3-one, and 3-keto-cholesterol (Sigma) Control bile (sample that has been repeatedly analyzed and has given consistent results) Chloroform (Merck) Methanol (Merck) Nitrogen source Pyridine (Thermo Scientific) N,O-Bis(trimethylsilyl)trifluoroacetamide (BSTFA; Sigma) Trimethylchlorosilane (TMCS; Thermo Scientific) Heptane Whole Body Cholesterol Fluxes in the Mouse

Single-housing cages for mice 0.5-ml microcentrifuge tubes Analytical balance with 0.1 mg accuracy

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Scale with 0.1 g accuracy (to weigh food and mice) Animal clippers Surgical instruments including Backhaus towel clamps Zoom stereomicroscope Cotton swabs 6–0 silk suture 20-G needle PE-10 cannula Incubator to keep the animals at a constant temperature of 37◦ C 10-ml glass screw-top tubes 50◦ C heat block Glass syringe GC vials Gas chromatographic (GC) system (van der Veen et al., 2009) Additional reagents and equipment for injection of mice (Donovan and Brown, 2006a) Cannulate gall bladder and collect bile 1. Accurately weigh (using an analytical balance) and label the microcentrifuge tubes before the bile is collected. 2. At a given hour of the day, cannulate the mouse’s gallbladder as follows: weigh the animals and anesthetize them by an intraperitoneal injection (Donovan and Brown, 2006a) of 10 μl of fentanyl citrate (0.315 mg/ml):fluanisone (10 mg/ml):diazepam (5 mg/ml):PBS at a ratio of 0.5:0.5:1:8 (v/v/v/v). 3. Shave the abdominal region, clean it with ethanol 70% (v/v), and perform a laparotomy through an upper-midline incision. Keep both sides of the incision separated by using two Backhaus towel clamps. The rest of the surgical procedure must be performed with magnification provided by a zoom stereomicroscope.

4. With a wet cotton swab, move the liver to one side in order to localize the bile duct and ligate it with a 6-0 silk suture. Make sure to avoid any damage to the liver, since liver injury will impair the bile flow. The contents of the gallbladder will flow out into the abdominal cavity, but due to its very small volume, it has no deleterious effects on tissues.

5. Next, pull the liver down with the wet cotton swab, in order to have a clear view to the gallbladder. With the help of a forceps, hold up the gallbladder and with the other hand puncture the gallbladder with a 20-G needle. 6. Insert a PE-10 cannula into the gallbladder and secure it with silk 6-0 suture to the gallbladder. Check whether there is flow, and tape the cannula in order to avoid any undesirable movement. Let the bile flow for the first 5 min. After this time, collect bile for a determined period of time. Timing is important for this procedure. Since biliary secretion rates vary throughout the day, it is crucial that bile be collected at the same time every day. This technique must be performed by an experienced animal technician. It is very important to ligate the biliary duct close to the intestine to avoid any leakage of bile into the intestine.

7. Collect bile in preweighed 0.5-ml microcentrifuge tubes for a chosen period while keeping the animals in an incubator at 37◦ C.

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Table 2 Calibration Curve to Measure Neutral Sterols and Cholesterol in Feces and Diet, Respectively

5αEpiCoprostanol Cholesterol cholestane coprostanol

Dihydrocholesterol

Coprostan3-one

3-ketocholesterol

nmol

nmol

nmol

nmol

nmol

nmol

nmol

Standard 1

5

0

0

0

0

0

0

Standard 2

5

1

1

1

1

1

1

Standard 3

5

2.5

2.5

2.5

2.5

2.5

2.5

Standard 4

5

5

5

5

5

5

5

Standard 5

5

10

10

10

10

10

10

Changes in body temperature will influence the bile flow. Therefore, it is imperative to keep the animals at a constant temperature of 37◦ C during bile collection. The duration of bile collection has ramifications for the total volume of bile collected, as bile flow decreases over time due to exhaustion of the endogenous bile salt pool. In general, bile collection times range between 10 and 30 min. An average bile flow ranges between 6.0 to 8.0 μl/min/100 g body weight.

Measure cholesterol content 8. Accurately weigh (using an analytical balance) the microcentrifuge tubes with the bile, and measure the concentration of cholesterol as previously described (Bligh and Dyer, 1959), with some modifications. Pipet 15 μl of bile into a 10-ml glass screw-top tube containing 5 nmol of 5α-cholestane (internal standard) dissolved in absolute ethanol (0.5 μmol/ml concentration;100 μl volume) and 1.2 ml distilled water. Include control bile, which will be analyzed every time that this protocol will be performed. 9. Add 4.5 ml chloroform:methanol (1:2, v/v), cap the tube with a plug, and mix by vortexing for 20 to 30 sec. 10. Add 1.5 ml of chloroform and mix by vortexing for 20 sec. 11. Add 1.6 ml of distilled water and mix by vortexing for 20 sec. 12. Centrifuge 10 min at 441 × g, 21◦ C. 13. Transfer the chloroform layer (bottom layer) with a glass syringe into a new 10-ml glass screw-top tube. 14. Evaporate the chloroform at 50◦ C under nitrogen flow. In parallel, prepare a calibration curve according to the scheme in Table 2. 15. Once all the samples and standards are dry, add 100 μl of a solution of pyridine:BSTFA:TMCS (5:5:0.1; v/v/v) and cap the tubes. Keep the tubes closed at 21◦ C for at least 1 hr (this procedure can also be performed overnight). 16. Evaporate the reagent at 21◦ C under the flow of nitrogen. Immediately after the tubes are dry, add 100 μl of heptane. 17. Mix and pour the contents into a GC vial. These samples are ready to be injected into the GC as previously described (van der Veen et al., 2009) Whole Body Cholesterol Fluxes in the Mouse

Calculate cholesterol secretion 18. Calculate the biliary cholesterol secretion (BCS; μmol/mouse/day) according to the following formula:

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BCS =

wbile – w0 • Cbile• 1.44 t Equation 3

where wbile is the weight of the tube with the bile collected, w0 is the weight (mg) of the empty tube, t is the time when the bile was collected (min), and Cbile is the concentration of cholesterol in the bile (nmol/μl).

DIRECT MEASUREMENT OF FRACTIONAL CHOLESTEROL ABSORPTION BY DETERMINATION OF LYMPHATIC CHOLESTEROL TRANSPORT

BASIC PROTOCOL 3

One of the important parameters to take into account in order to estimate the source of cholesterol found in the feces is the percentage of cholesterol absorbed in the intestine, or the fractional absorption (Fa). Several methods (direct and indirect) have been used to measure the fractional absorption (Wang and Carey, 2003): lymphatic cholesterol transport (Basic Protocol 3), the fecal dual-isotope ratio method (Alternate Protocol 1), and plasma dual-isotope ratio method (Alternate Protocol 2). The lymphatic cholesterol transport method described here is based on administration by gavage of radioactive or stable isotope labeled cholesterol. Subsequently, appearance of this compound in the lymph is monitored. Labeled palmitic acid is administered to control the uptake of lipid and should reach an absorption of about 90% (see Fig. 1).

Materials Appropriate mouse strain Sodium pentobarbital Solution of 2.5 μCi of [14 C]cholesterol, 5 μCi [3 H]palmitic acid, and 0.5% (w/v) taurocholate (NEN Life Science Products) dissolved in 100 μl medium-chain triglyceride oil (Mead Johnson) Solution of 0.5% taurocholate in medium-chain triglyceride oil (Mead Johnson) Scintillation fluid 0.5-ml heparinized microcentrifuge tubes 3-ml syringe Zoom stereomicroscope PE-10 polyethylene catheter 6–0 silk suture Adhesive tape (13 mm × 30 m) Infusion pump (Harvard Apparatus, cat. no. 70-2209) Incubator to keep the animals at a constant temperature of 37◦ C Scintillation counter capable of measuring 14 C and 3 H in same measurement Additional reagents and equipment for injection of mice (Donovan and Brown, 2006a) Animal surgery 1. At a given hour of the day, perform the lymph cannulations according to the standard protocol described (users may also reference Wang and Carey, 2003). Fast the mice overnight allowing water ad libitum. Weigh the animals and anesthetize them by an intraperitoneal injection (Donovan and Brown, 2006a) of pentobarbital (35 mg/kg). 2. Perform laparotomy under sterile conditions through an upper-midline incision. For better visualization of the mesenteric lymphatic duct, arch the animal dorsally over a 3-ml syringe.

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Cumulative radioactivity (%)

100 90 80 70 60 50 40 30 20 10 0

cholesterol palmitic acid

0

1

2

3

4 5 6 7 8 9 10 11 12 Infusion time (hr)

Figure 1 Simulated data for lymphatic transport of cholesterol. Approximately 35% of the administered [14 C] cholesterol and 93% [3 H]palmitic acid is recovered in the lymph.

3. With magnification provided by a zoom stereomicroscope, insert a PE-10 polyethylene catheter into the mesenteric lymphatic duct. 4. Externalize the catheter through the right abdominal wall and connect it with a heparinized microtube for collecting lymph. Immediately, insert a PE-10 catheter into the duodenum 5 mm distal to the pylorus, and secure it with a 6–0 silk suture and adhesive tape. 5. Next, externalize the duodenal catheter through the left abdominal wall and connect it to an infusion pump. During surgery and lymph collection, the mouse’s body temperature must be maintained at 37◦ C. Anesthesia may be maintained by administering intraperitoneal injections of pentobarbital at a dose of 17 mg/kg every 2 hr.

Tracer infusion 6. Administer 100 μl of medium-chain triglyceride oil containing 2.5 μCi of [14 C]cholesterol, 5 μCi [3 H]palmitic acid, and 0.5% taurocholate through the duodenal catheter. 7. Administer a solution of medium-chain triglyceride mixed with 0.5% taurocholate at a constant flow of 300 μl/hr during the whole experiment through the duodenal catheter.

Sample collection and analysis 8. Collect lymph hourly for up to 12 hr (there will be 12 tubes containing 1 hr worth of lymph per animal). 9. Vigorously mix aliquots of 50 μl lymph with 10 ml of scintillation fluid (ratio 1:200, v/v) for 10 min. Count the presence of [14 C]cholesterol and [3 H]palmitic in the lymph in a scintillation counter. 10. Calculate the percentage of cholesterol absorbed from the cumulative radioactivity in the lymph in the steady state (Fig. 1). ALTERNATE PROTOCOL 1 Whole Body Cholesterol Fluxes in the Mouse

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INDIRECT MEASUREMENT OF FRACTIONAL CHOLESTEROL TRANSPORT VIA FECAL DUAL-ISOTOPE RATIO METHOD Cholesterol absorption is calculated by studying the ratio between [14 C]cholesterol and [3 H]sitostanol in the mixture administered compared to that in the feces, assuming that 100% of the dose of sitostanol administered by gavage appears in the feces. Therefore, the fraction of cholesterol that it is not absorbed is calculated from the ratio of cholesterol/sitostanol that appears in feces. Current Protocols in Mouse Biology

The techniques to measure fractional cholesterol absorption described in Alternate Protocols 1 and 2 use radiolabeled isotopes. However, the radiolabel can be replaced by stable isotopes in order to reduce the risk of radioactive contamination. Only trained personal may perform the experiment using radiolabeled isotopes due to the risk of contamination and the potential harmful risk of these compounds. The use of one isotope or the other will depend on whether the laboratory is equipped with a gas-chromatograph-mass spectrometer to determine stable isotope enrichment.

Additional Materials (also see Basic Protocols 1, 2, and 3) Appropriate mouse strain Solution of 1 μCi of [14 C]cholesterol and 2 μCi [3 H]sitostanol (NEN Life Science Products) dissolved in 150 μl medium-chain triglyceride oil (Mead Johnson) Gavage needle 1. Administer 150 μl of medium chain triglyceride oil containing 1 μCi [14 C]cholesterol and 2 μCi [3 H]sitostanol by gavage. 2. Collect feces over a period of 4 days, allowing the animals to have food ad libitum. 3. Extract the radiolabel from 50 mg of feces as previously described (van der Veen et al. 2009). Add 10 ml of scintillation fluid to the dried extract. Mix vigorously for 10 min and count. Count the presence of [14 C]cholesterol and [3 H]cholesterol in the feces in a scintillation counter that allows the measurement of two different isotopes in one measurement. 4. Calculate the percentage of cholesterol absorbed according to the following formula:

Fa =

[

14

C]

[

3

H ] gavage – [ 14 C]

[

14

C]

[

3

[

3

H ] feces

H ] gavage

• 100

Equation 4

where Fa is the fractional absorption of cholesterol (%), [14 C] are the dpm of the [14 C]cholesterol and [3 H] refers to dpm of the [3 H]sitostanol.

INDIRECT MEASUREMENT OF FRACTIONAL CHOLESTEROL TRANSPORT BY THE PLASMA DUAL-ISOTOPE RATIO METHOD

ALTERNATE PROTOCOL 2

Similar to the fecal dual-isotope ratio method (Alternate Protocol 1), cholesterol absorption is calculated by studying in plasma the ratio between [14 C]cholesterol administered by gavage and [3 H]cholesterol administered intravenously 72 hrs after administration. The fraction of cholesterol absorbed is estimated by calculating the ratio of [3 H]cholesterol/[14 C]cholesterol, considering that 100% of the dose of intravenously administered [14 C]cholesterol appears in blood.

Additional Materials (also see Basic Protocols 1, 2, and 3) Appropriate mouse strain Solution of 2.5 μCi of [3 H]cholesterol (NEN Life Science Products) dissolved in Intralipid (20%, w/v) (Pharmacia) Solution of 1 μCi [14 H]cholesterol (NEN Life Science Products, Boston, MA) dissolved in 150 μL medium-chain triglyceride oil (Mead Johnson) Heparinized microcentrifuge tubes Additional reagents and equipment for obtaining blood from the mouse by cardiac puncture (Donovan and Brown, 2006b)

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1. Administer intravenously (Donovan and Brown, 2006a) 100 μl of Intralipid containing 2.5 μCi of [3 H]cholesterol and orally 150 μl of medium-chain triglyceride oil containing 1 μCi [14 C]cholesterol, under anesthesia. 2. Return the mice to their cages and give them food and water ad libitum. 3. After 72 hrs, bleed the animals from the heart (around 400 μl blood; Donovan and Brown, 2006b) into a microcentrifuge tube containing heparin as anticoagulant. Obtain plasma by centrifuging the blood 10 min at 956 × g, 4◦ C. 4. Vigorously mix aliquots of 100 μl plasma with 10 ml of scintillation fluid (ratio 1:100, v/v) for 10 min. Count the presence of [14 C] cholesterol and [3 H] cholesterol in the plasma and feces in a scintillation counter that allows the measurement of two different isotopes in one measurement. 5. Calculate the percentage of cholesterol absorbed according to the following formula:

Fa =

percentage of oral dose [14 C ] cholestrol per ml plasma percentage of iv dose [ 3 H ] cholestrol per ml plasma

•100

Equation 5

where Fa is the fractional absorption of cholesterol. BASIC PROTOCOL 4

ESTIMATION OF TRANSINTESTINAL CHOLESTEROL EXCRETION BY THE STEROL BALANCE METHOD Two different methods can be used to calculate the rate of cholesterol excreted directly through the intestine. A simple method is to determine cholesterol balance via the sterol balance method, which gives only an approximate result but does not require very sophisticated equipment. The second method gives a quantitative estimate of the various pathways of cholesterol excretion by modeling cholesterol fluxes using stable isotopes (Alternate Protocol 3), but it requires a gas chromatograph equipped with a mass spectrometer. The sterol balance method takes into account the different fluxes of cholesterol that enter and leave the intestine. Cholesterol enters the intestine via the diet, the bile, or the nonbiliary cholesterol pathway (e.g., transintestinal cholesterol excretion, intestinal shedding, cholesterol synthesized in the intestine and excreted directly to the intestine, or cholesterol synthesized in the liver and excreted directly in the intestine with the bile, without appearing in the blood compartment). Non-(re)-absorbed cholesterol leaves the body directly with the feces. The rate of cholesterol excreted via the non-biliary cholesterol pathway is calculated as follows:

NBC =

NSS – (DC + BCS ) (1 – Fa ) Equation 6

Whole Body Cholesterol Fluxes in the Mouse

where NBC is the rate of nonbiliary cholesterol excretion (μmol/mouse/day), NSS is the rate of neutral sterols excreted with the feces (μmol/mouse/day; Equ. 2), Fa is the fractional absorption of cholesterol (%; Basic Protocol 3), DC is the rate of cholesterol consumed in the diet (μmol/mouse/day; Equ. 1), and BCS is the rate of cholesterol excreted with the bile (μmol/mouse/day; Equ. 3).

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MEASUREMENT OF TRANSINTESTINAL CHOLESTEROL EXCRETION BY MODELING OF CHOLESTEROL FLUXES DETERMINED VIA STABLE ISOTOPE ENRICHMENT ESTIMATION

ALTERNATE PROTOCOL 3

The measurement of transintestinal cholesterol excretion by modeling the cholesterol fluxes includes the use of stable isotopes to follow the tracers given in the different compartments. This protocol requires the determination of the different cholesterol fluxes in the body, such as cholesterol intake, biliary cholesterol secretion, fractional cholesterol absorption, and neutral sterol excretion. The main advantage of this protocol is that it allows calculation of the different cholesterol fluxes in the mouse body. However, it requires knowledge on computational modeling of biological fluxes as well as the assistance of an experienced technician in gas chromatography-mass spectrometry.

Additional Materials (also see Basic Protocols 1, 2, and 3) D7 -cholesterol (purity >90%) (Cambridge Isotope Laboratories Inc.) Absolute ethanol (Merk, Darmstadt, Germany) D5 -cholesterol (purity >90%) (Medical Isotopes Inc.) Intralipid solution (20%; v/v) (Pharmacia) Medium chain triglyceride oil (Mead Johnson) Small sharp scissors Filter paper (Munktell & Filtrak, http://www.munktell.com) Sample preparation of the intravenous solution 1. Dissolve 5 mg of D7 -cholesterol in 1 ml of absolute ethanol (warm to 40◦ C if necessary). Evaporate the ethanol under a nitrogen stream and redissolve the residue in absolute ethanol to obtain a final solution of 1 mg D7 -cholesterol in 40 μl absolute ethanol (warm to 40◦ C). This solution is only stable for 24 hr at 21◦ C (do not store it at 4◦ C because this induces precipitation of cholesterol). The first solubilization of the cholesterol in ethanol will change its crystal form and will facilitate its solubilization in lipid solutions.

2. One day before the experiment, add 150 μl of Intralipid per 0.3 mg of D7 -cholesterol (directly to the ethanolic solution). Use a new bag of Intralipid every time solutions are prepared.

Sample preparation of the oral solution 3. Dissolve 5 mg of D5 -cholesterol in 1 ml of absolute ethanol (warm to 40◦ C) and evaporate it to dryness. Add 200 μl of medium-chain triglyceride per 0.6 mg of D5 -cholesterol. Make sure it is completely dissolved before administration. Isotope administration and sample collection 4. In order to obtain a baseline blood sample, cut a small piece of the tail (∼0.5 mm) using small sharp scissors. Take a small blood sample (∼5 μl) from the tail on a filter paper. 5. Just after the blood sampling, anesthetize the mice with isofluorane 0.8% and administer a solution of D7 -cholesterol intravenously (0.3 mg/mouse solved in 150 μl of Intralipid) and D5 -cholesterol orally (0.6 mg/mouse solved in 200 μl of mediumchain triglyceride oil). 6. Collect blood samples (∼5 μl) on filter paper as described in step 4 at the following time points: 3 hr, 6 hr, 12 hr, 24 hr, 48 hr, 72 hr, 96 hr, 120 hr, 144 hr, 168 hr, 192 hr, 216 hr, and 240 hr. The blood samples on the filter paper should dry at 21◦ C. These samples can be kept at 21◦ C until further analysis.

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7. During the last 3 days of the experiment, measure dietary cholesterol intake and neutral sterols excretion as previously described (Basic Protocol 1). 8. After the last blood sample, determine biliary cholesterol excretion, as described in Basic Protocol 2. 9. Measure the stable isotope enrichment of cholesterol in the blood spots, in the bile, and in the feces as previously described (van der Veen et al., 2009). 10. Determine the concentration of neutral sterols in the feces and the concentration of cholesterol in the bile as previously described as described in Basic Protocol 1 and Basic Protocol 2 (van der Veen et al., 2009).

Calculation of cholesterol excretion rates 11. Calculate the different rates of cholesterol excretions as follows: Fractional absorption (Fa) is calculated taking into the account the appearance of the label administered orally and the label administered intravenously according to the following formula (for further explanation see Alternate Protocol 2, plasma dual isotope-ratio method):

Fa =

AUCoral Div • • 100 AUCiv Doral Equation 7

where Fa is the fraction of cholesterol that it is (re)absorbed in the intestine (%), AUCoral is the area under the curve of the label administered orally, AUCiv is the area under the curve of the label administered intravenously, Div is the dose of label administered intravenously (μmol), and Doral is the dose of label administered orally (μmol). Cholesterol excreted in feces originating from the diet (D[NS]) is calculated by multiplying the flux of dietary cholesterol by the fraction of cholesterol that it is not absorbed.

D [NS ] = DC • (1 – Fa) Equation 8

where D[NS] is the flux of cholesterol appearing in feces that originates from the feces, DC is the amount of cholesterol coming from the diet (μmol/mouse/day; Equ. 1), and Fa is the fraction of cholesterol that it is (re)absorbed in the intestine (Equ. 7). Cholesterol excreted in feces originating from the bile (B[NS]) is calculated by multiplying the flux of cholesterol excreted via the bile by the fraction of cholesterol that it is not absorbed: B [NS] = BCS • (1 - Fa ) Equation 9

Whole Body Cholesterol Fluxes in the Mouse

where B[NS] is the flux of cholesterol appearing in feces that originates from biliary excretion, BCS is the rate of cholesterol excreted via the biliary route (μmol/mouse/day; Equ. 3) and Fa is the fraction of cholesterol that it is (re)absorbed in the intestine (Equ. 7).

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Cholesterol excreted in feces originating from transintestinal cholesterol excretion (TICE) is calculated by taking into account the enrichment of D7 -cholesterol in the different compartments (blood, feces and bile), according to the following formula:

TICE =

[(

IVchol feces 168 hr- 240 hr chol blood

IV

•

( )] NSS 1 - Fa

hr – 240 hr )• BCS] – [(IVchol bile • IV 168 chol blood

Equation 10

where TICE is the amount of cholesterol appearing in feces that originates from transintestinal cholesterol excretion (μmol/mouse/day), IVchol feces is the enrichment of D7 -cholesterol appearing in the feces (%), IVchol blood is the average of the enrichment of D7 -cholesterol appearing in the blood spots from the time point 168 hr and 240 hr (%), NSS is the neutral sterol secretion (μmol/mouse/day; Equ. 2), Fa is the fractional absorption (%; Equ. 7), IVchol bile is the enrichment of D7 -cholesterol appearing in the bile (%), and BCS is the biliary cholesterol secretion (μmol/mouse/day; Equ. 3). Cholesterol excreted in feces originating from other sources (intestinal shedding, cholesterol synthesized either in the intestine or in the liver and excreted directly into the feces). This fraction of cholesterol that appears in the feces is calculated as the remaining fraction that does not account for any of the other sources of fecal sterols.

REST = NSS –

(( D [NS] ) +(B [NS] ) + TICE

Equation 11

where REST is the fraction of cholesterol, TICE is the flux of cholesterol appearing in feces that originates from intestinal shedding or newly synthesized cholesterol that does not appear in the blood compartment (μmol/mouse/day), NSS is the neutral sterol secretion (μmol/mouse/day; Equ. 2), D[NS] is the cholesterol appearing in feces coming from the diet (μmol/mouse/day; Equ. 8), B[NS] is the cholesterol appearing in feces coming from the bile (μmol/mouse/day; Equ. 9), and TICE is the flux of cholesterol appearing in feces that originates from transintestinal cholesterol excretion (μmol/mouse/day; Equ. 10).

COMMENTARY Background Information In clinical cardiovascular practice, pharmacological therapies aim at decreasing LDL cholesterol levels. Until recently, little attention has been directed to assessment of fluxes instead of concentrations. However, times are changing and there is an increasing interest to better understand the cholesterol fluxes in the body, and to develop new approaches to increase the flux of cholesterol from the vessel wall towards the feces. Recent studies in animal models (van der Velde et al., 2008; van der Veen et al., 2009) demonstrated the intestine to be a key organ in regulating cholesterol excretion to the feces. This article reviews several experimental

approaches to measure cholesterol fluxes in the hepatobiliary system and in the intestinal pathway. One of the fluxes to consider is the measurement of the cholesterol intake with the diet by assessing the amount of food consumed per day and determining the amount of cholesterol present in the diet (Basic Protocol 1). A second important flux to measure is the rate of cholesterol (and its metabolites) excreted with the feces (Basic Protocol 1) by collecting feces over a period of time and, later on, determining the concentration of cholesterol in this matrix. The biliary cholesterol excretion is a third flux that needs to be determined since it accounts for at least 1/3 of the total excretion of

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cholesterol from the body (Basic Protocol 2). By collecting hepatic bile production, the daily biliary cholesterol excretion can be estimated. In normal wild-type mice, it is estimated that 40% to 60% of the cholesterol in the intestine is absorbed. However, this fraction may change depending on the mouse strain or the dietary/pharmacological treatment. Therefore, the calculation of the fractional absorption is warranted to study the rate of cholesterol excreted from the intestine (Basic Protocol 3). Finally, a last flux recently discovered, is the excretion of cholesterol directly through the intestine (transintestinal cholesterol excretion) (Basic Protocol 4). It has been estimated that in mice, half of the cholesterol excreted in the feces originates from this flux. Therefore, the measurement of this flux is crucial for a good estimation of the source of the cholesterol loss in the intestine.

Critical Parameters and Troubleshooting Measurement of dietary cholesterol intake and neutral sterol excretion (Basic Protocol 1) The measurement of dietary cholesterol intake and neutral sterol excretion will depend on the level of stress of the mice during the experiments as well as the accuracy of the researcher in performing the measurements. A reliable estimation of the food intake and fecal production is warranted for the success of these measurements. Therefore, it is important to make sure that the animals have a constant food intake by reducing the level of stress (e.g., avoiding extreme noise and excess handling). In wild-type mice fed chow diet, a normal food intake ranges between 16 and 20 g/day/100 g body weight. The amount of food intake is reduced to 8 to 12 g/day/100 g body when the mice are fed semi-synthetic diet. In case the animals are under a lot of stress, a reduction in food intake will be observed. Besides, the accurate recovery of the feces from the bedding will guarantee the success of the experiment, since the value of cholesterol excretion depends on the total fecal excretion. Wild-type mice fed chow diet excrete between 3.2 to 4.0 g/d/100 g body weight. Furthermore, changes in diet may influence the fecal excretion. Whole Body Cholesterol Fluxes in the Mouse

Measurement of biliary cholesterol (Basic Protocol 2) The most important factor in this protocol is the good training of the animal technician

in order to obtain a reliable hepatic bile flow. The bile flow in wild-type mice fed chow diet ranges between 6.0 and 8.0 μl/min/100 g body weight. In addition, several other factors may influence the outcome of this measurement. (1) The chosen time of the day to perform the experiment must be constant during the whole experiment, since biliary secretion rates have a strong circadian rhythm. It is advised to perform the bile cannulations within a margin of 3 hr. A larger range of time would result in significant variation between the first animal that has been cannulated and the last one. (2) The bile duct has to be properly closed to ensure there is no leakage of bile into the intestine. (3) The mice must be kept at a constant temperature of 37◦ C to avoid oscillations in the bile flow. (4) The most common procedure is to collect bile for a period of 20 to 30 min, since longer periods of time will deplete the bile acid pool resulting in oscillations in the bile flow. (5) The gas chromatograph must be validated in order to avoid variations in the measurement. The inclusion of control samples will determine the accuracy of the measurement. Measurement of fractional cholesterol (Basic Protocol 3 and Alternate Protocols 1-2) There are basically three different protocols to measure fractional cholesterol absorption. The decision to choose one or the other will depend on the skills of the animal technician and the equipment available for the analytical determinations. The most direct measurement of cholesterol absorption is based on determining the appearance of cholesterol in the lymph. However, it is not uncommon to observe variations in the lymph flow. Therefore, several controls must be included to ensure the reliability of this measurement. It is advised to measure the lymph flow every hour to ensure that this is constant over the whole experiment. The most indirect method consists on measuring the appearance the label sitostanol and cholesterol into the feces. Assuming that sitostanol is not absorbed, the ratio between cholesterol and sitostanol will give an estimate of the cholesterol that it is absorbed. This method has one major disadvantage: it assumes that 0% sitostanol is absorbed, although it is known that there is a small fraction of sitostanol that may be absorbed (Ostlund, 2002). These differences will introduce variations in the measurement. A third method to measure fractional absorption is the measurement of the appearance

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of two different cholesterol labels in the blood. The main advantage of this method is that is very easy to perform and it gives accurate results, in spite of being an indirect method. Estimation of transintestinal cholesterol excretion (Basic Protocol 4 and Alternate Protocol 3) In the literature, two different methods have been developed to estimate the in vivo flux of transintestinal cholesterol excretion. The first is based on performing a cholesterol balance by subtracting the fluxes that enter the body from the fluxes that exit the body. The main advantage of this measurement is that it is easy to perform and it does not require major equipment. However, this method gives only an approximate result. The second method requires the administration of stable isotopes. Therefore, one of the main disadvantages of this method is that it requires experience in modeling physiological fluxes as well as in gas-chromatography mass spectrometry. However, this method allows for a very accurate determination of the fluxes in the body, not only in the intestine, but also in the liver.

Anticipated Results Measurement of the different fluxes The values given as a reference in this article refer to male C57Bl/6J mice (12 weeks old) fed chow diet ad libitum. Major differences in every flux may be observed in case of changes in mouse strain, diet, gender or age. A control group (from which references values have been obtained in previous experiments) should always be included in order to compare results with previous experiments. In addition, some of the techniques require very sophisticated surgical procedures. Therefore, it is warranted that these procedures are performed by a technician with enough experience to detect problems during the surgery. The success of the surgical procedure will guarantee the reliability of the results.

Time Considerations In principle, all the flux measurements described in this protocol can be performed on a single mouse in a 2 weeks time period. Of course, additional time is required to perform the rather time consuming mass spectrometry determinations in the analytical laboratory; the time required will depend on the facilities available in every facility. Clearly careful planning of the experiment is imperative in order to obtain as much information as possible, but always ensuring the reliability of the results. For example, biliary cholesterol excretion cannot be calculated in the same mouse where fractional absorption is calculated by using the lymphatic measurement (direct approach).

Literature Cited Bligh, E.G. and Dyer, W.J. 1959. A rapid method of total lipid extraction and purification. Can. J Biochem. Physiol. 37:911–917. Donovan, J. and Brown, P. 2006a. Parenteral injections. Curr. Protoc. Immunol. 73:1.6.1-1.6.10. Donovan, J. and Brown, P. 2006b. Blood collection. Curr. Protoc. Immunol. 73:1.7.1-1.7.9. Keys, A., Aravanis, C., Van Buchem, F.S.P., Blackburn, H., Buzina, R., Djordjevic, B.S., Dontas, A.S., Fidanza, F., Karvonen, M.J., Kimura, N., Menotti, A., Nedeljkovic, S., Puddu, V., Punsar, S., and Taylor, H.L. 1981. The diet and allcauses death rate in the Seven Countries Study. Lancet 2:58–61. Ostlund, R.E. Jr. 2002. Phytosterols in human nutrition. Ann. Rev. Nutr. 22:533-549. van der Veen, J.N., van Dijk, T.H., Vrins, C.L., van Meer, H., Havinga, R., Bijsterveld, K., Tietge, U.J., Groen, A.K., and Kuipers, F. 2009. Activation of the liver X receptor stimulates trans-intestinal excretion of plasma cholesterol. J. Biol. Chem. 284:19211-19219. van der Velde, A.E., Vrins, C.L., van den Oever, K., Seemann, I., Oude Elferink, R.P., van Eck, M., Kuipers, F., and Groen, A.K. 2008. Regulation of direct transintestinal cholesterol excretion in mice. Am. J. Physiol. Gastrointest. Liver Physiol. 295:G203-G208. Wang, D.Q.H. and Carey, M.C. 2003. Measurement of intestinal cholesterol absorption by plasma and fecal dual-isotope ratio, mass balance, and lymph fistula methods in the mouse. J. Lipid Res. 44:1042–1059.

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Quantal Analysis of Endplate Potentials in Mouse Flexor Digitorum Brevis Muscle Richard R. Ribchester1 1

Euan MacDonald Centre for Motor Neurone Disease Research, University of Edinburgh, George Square, Edinburgh, Scotland, United Kingdom

ABSTRACT The isolated flexor digitorum brevis (FDB) muscle from mice is extremely well suited to rapid acquisition of data and analysis of neurotransmitter release and action at neuromuscular junctions, because the muscle and its tibial nerve supply are simple to dissect and its constituent muscle fibers are short (<1 mm) and isopotential along their length. Methods are described here for dissection of FDB, stimulation of the tibial nerve, microelectrode recording from individual muscle fibers, and quantal analysis of endplate potentials (EPPs) and miniature endplate potentials (MEPPs). Curr. Protoc. Mouse Biol. C 2011 by John Wiley & Sons, Inc. 1:429-444  Keywords: neuromuscular junction r endplate potential r intracellular recording r electrophysiology r quantal analysis

Neuromuscular junctions (NMJs) in skeletal muscles of vertebrates are highly accessible experimental objects, favorable for discovery and analysis of fundamental principles of synaptic structure and function (Katz, 1996). In the 1950s, intracellular recordings with glass microelectrodes and electron microscopy laid the foundation for understanding the mechanism of transmission by exocytosis of neurotransmitter molecules and their action on specific receptors (Katz, 1969). Such analysis was subsequently shown to explain synaptic transmission at all chemical synapses, in every part of the nervous system, in every species examined experimentally. More recently, the advent of transgenic technology and various advanced forms of microscopy, as well as the blossoming of optical techniques for making physiological measurements, have broadened the horizons for research into the principles governing neuromuscular connectivity (“connectomics”) and the regulation of synaptic survival and strength (Livet et al., 2007; Lu et al., 2009; Nagwaney et al., 2009; Ribchester, 2009; Wong et al., 2009; Ruiz et al., 2011). However, quantal analysis of neuromuscular transmission still forms the basis of many studies into the physiology of neuromuscular development, maintenance, repair, aging, and disease (Gillingwater et al., 2002; Ribchester et al., 2004; Slater et al., 2006; Slater, 2008; Zitman et al., 2008). Standard preparations that permit data to be obtained on spontaneous and evoked neurotransmitter release and action efficiently are therefore both desirable and useful. The mechanism of neuromuscular transmission was originally established from studies of the frog sartorius muscle, but there are many disadvantages to this preparation, stemming mostly from the thickness of the muscle and the time-consuming nature of positioning the recording microelectrode in the vicinity of an NMJ. The cutaneous pectoris muscle is a better alternative in this species because this muscle is very thin and the location of the motor nerve terminals can more readily be gauged using dark-field illumination and a standard dissecting microscope or a compound microscope (Blioch et al., 1968; Betz and Bewick, 1993). Standard neuromuscular preparations for electrophysiological analysis of neuromuscular transmission in mice include the fast-twitch extensor digitorum longus

Current Protocols in Mouse Biology 1: 429-444, December 2011 Published online December 2011 in Wiley Online Library (wileyonlinelibrary.com). DOI: 10.1002/9780470942390.mo110127 C 2011 John Wiley & Sons, Inc. Copyright 

BASIC PROTOCOL

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muscle, the slow-twitch soleus muscle, and the hemidiaphragm (Harris and Ribchester, 1979a,b). While these preparations have the principal merit of being well characterized, they are not the best for making routine electrophysiological recording or analysis of synaptic (endplate) potentials. The principal disadvantages are their thickness—making precise visualization of the NMJs problematical and the risks of hypoxia greater—and the time taken for careful placement of a recording microelectrode tip at the NMJ, which is required for high-fidelity recording of spontaneous and evoked postsynaptic responses in cases where the muscle fibers are relatively long. Thin preparations that facilitate visualization of murine NMJ under a dissecting microscope include, for example, levator auris longus, triangularis sterni, or transversus abdominis (McArdle et al., 1981; Angaut-Petit et al., 1987; Gillingwater et al., 2002). Good dissection is relatively difficult, but paramount for setting up these preparations, and this is relatively time consuming. These preparations also require careful placement of the electrode tip in the near vicinity of the NMJ, or else the recordings are attenuated by the “cable properties” of the muscle fibers (particularly the time constant and length constant) with excessive leakage of synaptic current occurring if the microelectrode is located more than ∼200 μm distant from the current source at the endplate (Auerbach and Betz, 1971; McLachlan and Martin, 1981). These requirements are especially disadvantageous if data throughput is of high priority. The isolated flexor digitorum brevis (FDB) of rodents is extremely well suited to rapid acquisition of data and quantal analysis neurotransmitter release and action at neuromuscular junctions. The principal advantages of this preparation are as follows: a. The muscle fibers of mouse or rat FDB are short: ∼300 to 800 μm in length and about 30 μm in diameter. This property renders the fibers “isopotential” along their length: that is, current generated from a source located at any point along the fiber (including its NMJ) can be faithfully recorded by a microelectrode located anywhere else in the fiber (Bekoff and Betz, 1977a,b). Thus, it is unnecessary to deliberately place a recording microelectrode near an NMJ in order to obtain high-fidelity recording of either spontaneous MEPPs or evoked EPPs: the fibers can be impaled at random spots and maximal, unattenuated, endplate potentials with a rapid time course will be observed (Ribchester et al., 1995, 2004; Gillingwater et al., 2002). b. The muscle is located superficially on the plantar side of the foot and it is therefore quite easy, after just a few attempts, for a student or other junior researcher who has reasonable manual dexterity to dissect an undamaged FDB muscle with its tibial/medial-plantar nerve supply intact. c. The muscle fibers have a pennate (feathered) organization. This feature is very ‘forgiving’ during the training period when dissection technique is being learned, especially if self-taught or with minimal instruction or supervision. Damage to part of the muscle during dissection, for example, still leaves many intact fibers in other parts of the muscle, from which good resting membrane potentials (around −70 mV) and MEPPs, at least, may still be observed and recorded.

Quantal Analysis of Endplate Potentials in Mouse FDB Muscle

Although imperfect for whole mounts, the FDB muscle, particularly in mice, is also suitable for subsequent morphological analysis of NMJ using conventional or confocal microscopy, for instance after staining endplates with fluorescent conjugates of α-bungarotoxin (Gillingwater et al., 2002). FDB muscles can also be quite readily dissected or dissociated into single fibers using collagenase, then either studied acutely using a variety of physiological methods, or after culture in vitro (Bekoff and Betz, 1977a,b; Gillespie and Ribchester, 1988; Lupa and Caldwell, 1991; Yeung et al., 2002; DiFranco et al., 2011; Nocella et al., 2011).

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NOTE: All protocols using live animals must first be reviewed and approved by an Institutional Animal Care and Use Committee (IACUC) or must conform to governmental regulations regarding the care and use of laboratory animals.

Materials Physiological saline solution (see recipe) Mouse (any laboratory strain of any age/gender; e.g., C57B16) μ-conotoxin GIIIB (Bachem) 4 M potassium acetate Dissection tools (iris scissors, spring scissors, fine forceps) Sylgard-lined Petri dish (see recipe) Fine minutien pins (Fine Science Tools) Suction electrode (see recipe) Micromanipulators (e.g., Leica or Sutter Instruments) Recording chamber: the author’s laboratory uses a Perspex chamber made in-house, but a Sylgard-lined Petri dish (see recipe) will suffice 1-mm glass capillary tubing containing and internal welded glass filament (1.5 mm O.D., 0.84 mm I.D. standard wall borosilicate glass with filament; World Precision Instruments, cat. no. 1B150F-6) Electrode puller: Among the most popular electrode pullers with electrophysiologists is the Brown-Fleming puller manufactured by Sutter Instruments Syringe with tip drawn out after warming the plastic in a Bunsen burner flame Microelectrode amplifier (Axoclamp 2B, Axon Instruments) Silver-silver chloride wire or pellet (0.8 mm diameter × 20 mm Ag/AgCl electrode; World Precision Instruments, cat. no. EP08) Mains interference filter (Digitimer Humbug; optional) Low-pass (<2 kHz) filters (Neurolog, Digitimer, UK) Pulse train generator (Digitimer D4030 Programmer) Isolated stimulator (Digitimer DS2) Dissecting microscope Light source for dissecting microscope (dark-field condenser or flexible fiber-optic light) Recording display device (oscilloscope, or computer running WinWCP) WinWCP software program (Strathclyde Electrophysiology Software; (http://spider.science.strath.ac.uk/sipbs/showPage.php?page=software ses) Data acquisition unit (CED micro 1401, Cambridge Electronic Design, http://www.ced.co.uk/) Additional reagents and equipment for sacrifice of the mouse (Donovan and Brown, 2006) Perform dissection 1. Make up one liter of physiological saline solution. 2. Sacrifice a mouse using an authorized, legal method approved by the institution where the research is to be conducted. Use large scissors to remove one or both hind limbs near the hip. Stunning, immediately followed by cervical dislocation (Donovan and Brown, 2006) is suitable and swift.

3. Use iris scissors to make a skin incision along the anterior of the amputated limb and dorsum of the foot. Grip the skin firmly and strip it in one continuous motion from the leg and foot.

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4. Using fine minutien pins, pin the isolated, skin-stripped limb to the base of a Petri dish lined with Sylgard, and containing enough physiological saline to keep the exposed tissue submerged during the dissection. The plantar surface of the foot and medial side of the calf should be pinned uppermost.

5. Expose the tibial nerve by separating the superficial calf musculature and carefully cutting through connective tissue fascia and fat using miniature spring scissors. Cut the tibial nerve close to the knee and then dissect it along its length into the foot, taking special care where the nerve traverses the heel, where it can easily be damaged. Cut the lateral plantar branch of the tibial nerve and continue the dissection of the medial plantar nerve into the foot, passing alongside the FDB muscle. A side branch of the medial plantar nerve enters the muscle midway along its length. Although it may appear aesthetically pleasing to clean away all the connective tissue adhering to the nerve and the medial side of the FDB muscle, there is no real need to do so; attempting this can result in stretching or other form of damage to the nerve, sufficient to block nerve conduction.

6. Cut the proximal tendon of the FDB at the heel and dissect connective tissue from both the lateral and medial sides. Reflect the nerve-muscle preparation towards the toes and cut the three distal tendons, close to their insertions in the underlying calcaneus. This involves, in passing, cutting through the belly of superficial lumbrical muscles originating on the distal tendons of FDB.

7. Secure the isolated FDB muscle with attached tibial/medial-plantar nerve to the Sylgard-lined dish with fine minutien pins inserted through the proximal tendon and all three distal tendons. Clean away connective tissue and superficial nerve branches running over the distal third of the muscle that are cutaneous branches of the medial plantar nerve. The preparation should now resemble the one shown in Figure 1A.

Stimulate and record EPPs 8. Connect a suction electrode (mounted on a coarse micromanipulator) to the nerve. Aspirate the tibial nerve into the pipet tip and apply a brief (0.2 msec) maximal (nominally 10 V) stimulating pulse to the nerve. The muscle should twitch. If the intention is then to record action potentials, the next step can be skipped. However, if the aim is to perform a quantal analysis the muscle action potential and the twitch must first be blocked.

9. Expel the nerve from the suction pipet. Incubate the muscle in a small volume of physiological saline containing 2 μM μ-conotoxin GIIIB for 20 to 30 min. This should be sufficient to block the NaV 1.5 channels in the muscle fibers while leaving presynaptic action potentials and evoked release of transmitter unaffected.

10. Transfer the isolated nerve-muscle preparation to a recording chamber. If using gasequilibrated, bicarbonate-buffered physiological saline, ensure that the chamber is perfused with this solution at a rate of about 1 ml/min in order to maintain the pH. If the experiments are to be conducted at ambient temperature (up to 25◦ C), as assumed here, no perfusion should be necessary when using thoroughly oxygenated or aerated HEPES-buffered physiological saline. Quantal Analysis of Endplate Potentials in Mouse FDB Muscle

11. Pull glass microelectrodes from 1-mm glass capillary containing an internal welded glass filament. Back-fill the electrodes (take care not to touch or break the tips) with 4 M potassium acetate using a syringe whose tip has previously been drawn out after warming the plastic in a Bunsen flame.

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A

tibial nerve

B rise time (1–2 msec)

amplitude (1–40 mV) half-decay time (2–3 msec)

stimulus artifact

latency (1–2 msec)

Figure 1 Isolated tibial nerve/FDB muscle preparation and intracellular recordings of EPPs. (A) Typical appearance of a satisfactory working dissection of the FDB muscle and its attached tibial nerve supply. The tendon of origin at the heel (top) and the three distal tendons are pinned to a Sylgard-lined dish with minutien pins. (B) Superimposed digital sweeps during nerve stimulation at about 1/sec from an approximate location in the muscle indicated in A. Dotted lines and arrows show properties of the EPP that are routinely measured, for instance, using WinWCP software. The recording was made after blocking muscle fiber action potentials using μ-conotoxin GIIIB.

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The dimensions and shape of the electrode tips should confer electrical resistances (or impedances) of about 30 M, which can be checked using a built-in current source in most microelectrode amplifier systems.

12. Connect the filled electrode to the high-impedance headstage of a suitable amplifier, for example an Axoclamp 2B, mounted on a micromanipulator. Micromanipulate its tip into the solution bathing the isolated FDB. Ground the solution in the recording chamber with a silver-silver chloride wire or pellet. Adjust the DC offset on the amplifier to read zero millivolts and check the electrode resistance. Connect other shielding wires from manipulators, chamber holder, microscope stand, and other equipment to a common earthing point, to reduce or abolish ‘ground loops.’ A ‘Humbug’ (Digitimer) mains interference filter is useful and usually effective for eliminating any residual 50-Hz noise (60 Hz in the U.S.A.) if attempts to minimize earth loops by grounding to a common point are only partially successful. Low-pass (< 2 kHz) filters (for example, Neurolog, Digitimer) are also important for optimizing the fidelity of recordings with minimal attenuation of the biological signals, while removing other types of high-frequency noise.

13. Aspirate the tibial nerve into the suction pipet/electrode connected to the isolated stimulator. The timing of the stimulus pulses should be triggered with an appropriate device: for example, a Digitimer 4030 Programmer connected to a DS2 stimulator (Digitimer). Set rectangular pulse duration to 0.2 msec and stimulating voltage to between 1 and 10 V. Stimulation frequency should initially be 0.5 to 1 Hz.

14. Use a dissecting microscope and micromanipulator to bring the microelectrode tip into the vicinity of a muscle fiber. Monitor the voltage at the microelectrode tip simultaneously using one channel with direct coupling (DC) at low gain (20 mV/cm on a digital oscilloscope) and another channel in parallel with alternating coupling (AC) at higher gain (1 mV/cm or higher). 15. Use the micromanipulator to lower the tip of the electrode to just above the surface of the preparation while illuminating the preparation through a dark-field condenser or an appropriately-angled fiber-optic light source to help visualize the approximate location of the electrode tip. Gently tap the micromanipulator while carefully manipulating the microelectrode tip onto the surface of a muscle fiber—if the electrode tip is sharp enough, whiplash-penetration of a fiber will register a clean step to around −70 mV on the DC recording. Note this resting membrane potential. Spontaneous MEPPs should be visible on the AC recording, provided the resting potential is more negative than about −45 mV. These will occur unpredictably at a mean frequency between about 0.2-2.0/sec.

16. Trigger the recording display device (oscilloscope or computer running WinWCP; see step 17) and apply a stimulus to the nerve (0.2 m pulse duration; nominally 1- to 5-V pulse amplitude), delaying the stimulus pulse for about 5 msec.

Quantal Analysis of Endplate Potentials in Mouse FDB Muscle

An action potential may be observed if the dissection is good and the resting potential is more negative than about −60 mV. The muscle may contract and dislodge the microelectrode, causing subsequent loss of resting potential. If μ-conotoxin has been administered correctly, however, the recording should be stable and response should be a large (>20 mV) EPP. The muscle should not contract. Repetitive stimulation at 1/second should evoke EPPs resembling those shown superimposed in Figure 1B. These repeated responses may be quite uniform in amplitude if the quantal content is high, but fluctuate more in amplitude from stimulus to stimulus if the quantal content is relatively low.

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Perform quantal analysis 17. Capture trains of EPPs using a computer running a public-domain software package, free to academic users, called WinWCP (Strathclyde Electrophysiology Software; (http://spider.science.strath.ac.uk/sipbs/showPage.php?page=software ses). A suitable A/D interface is required, such as the CED micro 1401 (Cambridge Electronic Designs). Analysis of stored data can be performed offline for measurement mean amplitude, rise time, and time to half decay of the EPPs as their principal characteristics. Expect EPP rise times to be 1 to 2 msec; amplitudes to be in excess of 20 mV, and half-decay times on the order of 2 to 4 msec.

18. Carry out a quantal analysis using the ‘variance method’ built into WinWCP. This method is based on the Poisson equation. Quantal content, m, is given by the reciprocal of the square of the coefficient of variation of EPP amplitudes, after correction for nonlinear summation (see Commentary). Check EPPs for drift using the facility in WinWCP to graph EPP amplitude against stimulus number. Quantal analysis should only be applied to EPPs that fluctuate around a stationary mean. Using the last 10 to 30 EPPs in a train may be sufficient to meet this criterion. The software routines in WinWCP will calculate the quantal content of EPPs after correction for nonlinear summation according to an empirically derived formula based on voltage-clamp experiments (McLachlan and Martin, 1981; see Commentary under Equation 2). For relatively long muscle fibers such as those in classical preparations like EDL, soleus, diaphragm, or triangularis sterni, the value of 0.8 is about appropriate for f. For FDB muscle fibers, however, enter an "f" factor of 0.3 to 0.4. The mean amplitudes of MEPPs (either measured or estimated by eye) can be entered to compare the “direct” calculation of quantal content with the variance method. Quantal contents of around 50 are typical for mouse FDB muscle fibers. The variability in the EPP amplitude can be increased, with a concomitant reduction in mean amplitude, by exchanging the physiological saline for one in which Mg2+ concentration is increased to 3 to 4 mM and Ca2+ concentration is reduced to 1 mM. Under such conditions, the mean quantal content, m, may be reduced to about 3 or less, and either the direct method or the method of failures may be used to estimate m (see Commentary).

Analyze MEPPs If there is interest only in recording spontaneous transmitter release, then resting frequency of MEPPs can be increased if desired, by increasing the temperature of the bathing solution from ambient to 32◦ to 37◦ C, although be aware that as the temperature rises to within the physiological range, more effective perfusion with oxygenated saline may be required, to counteract hypoxia in the motor nerve terminals. Alternatively, e.g., at room temperature, MEPP frequency can be elevated by increasing the K+ ion concentration in the bathing medium to 10 to 15 mM. Also expect depolarization of the resting membrane potential of the muscle fibers to about −50 mV, with this level of elevated potassium ions (for an elegant simulator of the effects of changing the concentration of sodium, potassium and chloride ions on the resting potential, see http://www.nernstgoldman.physiology.arizona.edu/). Adding lanthanum chloride (2 μM) also elevates basal MEPP frequency (Curtis et al., 1986; Ribchester et al., 1998). 19. Appraise the stochastic nature of occurrences of MEPPs by counting the number of MEPPs in successive time intervals (sweeps) of 1- to 10-sec duration. The distribution of frequencies (0,1,2,....; .etc.) per sweep normally conforms to a Poisson distribution:

P(x)=

e− m m x x!

Equation 1

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where P(x) is the observed frequency of (x) MEPPs per sweep, and m is the mean frequency per sweep. The observed and predicted distributions may be compared statistically using a c2 test.

20. Derive and measure properties of spontaneous MEPPs, including their amplitude, rise time, decay time, mean frequency, and interval distribution, offline, from data files of continuous records using a commercial software program for PCs called Mini Analysis (Synaptosoft). The program incorporates data-conversion software for several industry-standard hardware interfaces and data file formats, including Axon Instruments and CED1401 series of interfaces.

REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps.

Physiological saline solution For 1 liter: 40 ml 3 M NaCl 5 ml 1 M KCl 10 ml 0.2 M CaCl2 5 ml 0.2 M MgCl2 2 ml 0.2 M NaH2 PO4 938 ml H2 O. Mix and add 2 g NaHCO3 and 1 g D-glucose, then bubble through tubing or an aspirator with a 95% O2 /5%CO2 gas mix for at least 10 min before use. As an alternative, omit NaH2 PO4 and NaHCO3 and instead add HEPES to a final concentration of 5 mM, and adjust pH to 7.2 to 7.4. Bubble with either air or 100% O2 .

Suction electrode This electrode can be manufactured from 2 mm capillary glass, narrowed in a Bunsen flame, cut with a serrated blade or diamond knife, and flame-polished to a tip diameter about 50% larger than the diameter of the tibial nerve at its tip. This pipet should then be fitted with an internal silver or platinum wire and a similar but longer, external wire coiled around the capillary glass to its tip. Connect the wires via appropriate plugs to an isolated stimulator and connect a small syringe, via silicone tubing, to the back of the suction electrode/pipet. Mount the whole assembly onto a coarse micromanipulator.

Sylgard-lined Petri dishes Using the Dow Corning Sylgard 184 silicone elastomer kit, mix 1 ml of silicone elastomer curing agent with 9 ml of silicone elastomer base and pour into a 60 × 15–mm plastic petri dish. Leave on a level surface to cure (may take 2 to 3 days to cure at room temperature). Alternatively place in a 37◦ C oven overnight.

COMMENTARY Background Information

Quantal Analysis of Endplate Potentials in Mouse FDB Muscle

Intracellular recording of synaptic potentials using glass microelectrodes connected to high-impedance amplifiers was introduced in the late 1940s and energetically applied to the analysis of chemical synaptic transmission at neuromuscular junctions since the early 1950s, initially by B. Katz and his

colleagues at University College London. Their electrophysiological analysis of synaptic transmission at neuromuscular junctions in frog skeletal muscle quickly established the “quantal hypothesis” of synaptic transmission. Subsequent morphological analysis of early transmission electron micrographs translated this concept to a “vesicle

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hypothesis” (Katz, 1969, 1996; Slater, 2008). According to this—now central—dogma in synaptic biology, the neurotransmitter in motor nerve terminals—acetylcholine—is synthesized from acetyl CoA and choline via the enzyme cholineacetyltransferase; stored in synaptic vesicles by activation of proton/acetylcholine co-transporters; released by Ca-dependent exocytosis following the depolarization of motor nerve terminals; acts on postsynaptic acetylcholine receptors that gate changes in motor endplate cationic permeability; and is inactivated and terminated in action by acetylcholinesterase enzymatic activity in the synaptic basal lamina. There are good accounts of the steps and molecular events that underpin neurotransmitter release and action at cellular and molecular levels in several neuroscience textbooks (e.g., Byrne and Roberts, 2009). In outline, transmitter release by exocytosis depends on depolarization and the inward flux of Ca2+ ions that it triggers. Cooperativity in the binding of Ca2+ to synaptotagmin and, possibly, other presynaptic proteins, enhances the probability that synaptic vesicles docked at active zones in motor nerve terminal membranes will undergo exocytosis (Liu et al., 2011). Each vesicle in a motor nerve terminal stores 5,000 to 10,000 molecules of the neurotransmitter acetylcholine. Thus, fusion of a single vesicle with the presynaptic terminal membrane, through voltage and Ca-dependent interaction of SNARE proteins, releases a discrete bolus (“quantum”) of transmitter molecules. These diffuse across the synaptic cleft and, within a few microseconds, a proportion of them bind to postsynaptic acetylcholine receptors. These ligand-gated ion channels transiently open, initiating a transient, net inward current. This current is the result of the net flux of cations, mainly influx of Na+ and efflux of K+ ions through the transmembrane pores formed by five subunits of ACh receptors (2 a, 1 b, 1 g/e and 1 d). The summed activation of the set of ACh receptors/channels bound by the ACh molecules released from one synaptic vesicle produces a transient inward current, called the miniature endplate current (MEPC). This generates a transient depolarization, ∼1 mV in amplitude, with a rise time of about 1 msec and a 50% decay time of about 2.5 msec, called the miniature end-plate potential (MEPP). MEPPs can be quite easily recorded using sharp-tipped (<1 μm diameter) glass microelectrodes filled with concentrated electrolyte (typically 3 M KCl or 4 M potassium acetate) connected via

a high-impedance headstage, through highpass and low-pass filters, amplified and displayed either on an oscilloscope screen or digitized through an A/D converter into a personal computer. Miniature end-plate potentials and quantal size MEPPs occur intermittently and unpredictably in resting muscle fibers (without nerve excitation) at a frequency averaging about 1/second (Fig. 2A). The intervals between MEPPs vary randomly and follow an exponential distribution, analogous to the random decay of radioactive nuclides (Figure 2B). They have a time course very similar to that of the EPP but are considerably smaller in amplitude, varying from just greater than the noise level to about 2 to 4 mV (Figure 2C,D). Occasionally, ‘giant’ MEPPs greater than 5 mV may be observed. Measurement of the mean amplitude of a series of spontaneous MEPPs (excluding ‘giants’) establishes the “quantal size” for that neuromuscular junction (Slater, 2008). The overall incidence of MEPPs can be increased by tonic depolarization of nerve terminals, most simply by elevating extracellular potassium ion concentration, although the disadvantage of this method is that the amplitude of MEPPs is reduced, because muscle fiber depolarization also reduces the inward “driving force” on the permeant ions that carry positive ionic current. A judicious combination of modestly elevated K+ (10 to 15 mM) plus a low concentration of lanthanum ions (La3+ , 2 μM) can produce an enhancement of MEPP frequency without substantially compromising their amplitude (Ribchester et al., 1998). Evoked end-plate potentials and quantal content Nerve excitation, as indicated above, transiently elevates intracellular Ca2+ and, within about a millisecond, this promotes fusion of many docked vesicles at active zones (Nagwaney et al., 2009). The result is exocytotic fusion of several tens of vesicles with the presynaptic membrane, releasing their contents into the synaptic cleft within microseconds. The resulting multi-quantal bolus of ACh activates a multiple of the receptors activated by the contents of a single vesicle. How many vesicles fuse as a result of excitation of the nerve terminal depends on the Ca2+ influx, mainly through voltage-gated P/Q type Ca-channels, opened by the presynaptic action potential (Urbano et al., 2002).

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A

B

40 30 20 10

3 mV 0 0

2000

30 sec

C

4000

6000

8000

10000

Inter-event interval (msec)

D

25 20 15 10 5

1 mV

0 0 5 msec

F

10

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Fraction

E

5 Amplitude (mV)

0.2

0.1

1 mV 0.0 0.0 0.4 0.8 1.2 1.6 2.0 2.4 2.8 10 msec

Amplitude (mV)

Figure 2 Typical quantal analysis of EPPs and MEPPs. (A) Slow scan of resting membrane potential in an FDB muscle fiber showing random, spontaneous MEPPs occurring with unpredictable intervals between events. (B) Histogram of inter-MEPP intervals, by frequency, from the data shown in A. (C) average of about 30 MEPPs from A, on a fast time base, showing principle characteristics and double-exponential curve fit to the decay time. (D) MEPP amplitude frequency histogram of the individual records. (E) Quantized fluctuations, including failures, recorded in an FDB muscle fiber from a preparation bathed in physiological saline with reduced Ca2+ and increased Mg2+ . (F) EPP amplitude histogram from 30 successive EPPs elicited at 0.5 Hz from a preparation bathed in physiological saline with reduced Ca2+ and increased Mg2 (different fiber from E). There were two ‘failures’ in this series, indicating a mean quantal content of 2.71. The variance method indicated a mean quantal content of 3.11 and the direct method based on a mean MEPP amplitude of 0.4 mV gave 2.84 as the mean quantal content.

Quantal Analysis of Endplate Potentials in Mouse FDB Muscle

The number of vesicles undergoing nerveevoked exocytosis is called the “quantal content” of the evoked endplate current (EPC) or corresponding endplate potential (EPP). Normally, an EPP is substantially larger in amplitude than that required to trigger an action potential in the muscle fiber. Thus, in order to observe the EPP, unadulterated by the regenerative effects of voltage-activated Na or K channels, it is necessary to suppress

the postsynaptic, muscle-fiber action potential. In mammalian muscle this can most effectively be achieved using the marine snail toxin, μ-conotoxin GIIIB at concentrations of 1 to 2 μM (Braga et al., 1992; Wood and Slater, 1997; Gillingwater et al., 2002). At this concentration, the toxin binds reversibly to the NaV 1.5 isoform of voltage-gated Na channel normally present in skeletal muscle fiber membranes, while leaving the presynaptic action

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potential unscathed, since this arises from Na-channels that are much less sensitive to μ-conotoxin GIIIB. A classical, but inferior alternative method is to partially block the postsynaptic response with curariform drugs such as d-tubocurarine. However, this method also reduces the amplitude of the MEPPs, rendering accurate measurements of quantal size virtually impossible. A third method is the simple expedient of cutting the ends of the muscle fibers, leaving the endplates intact (Auerbach and Betz, 1971; Harris and Ribchester, 1979a,b). This causes a large depolarization of the muscle fibers (down to about −30 mV or less), inactivating voltage-gated Na channels and thus blocking muscle fiber action potentials. However, because of the reduced voltage driving force, the MEPPs are also normally obscured by the baseline noise in the recordings, which again makes accurate measurement of quantal size unfeasible. Nonlinear summation of EPPs Unfortunately, it is not possible to ascertain the number of vesicles released by a nerve stimulus, under normal physiological conditions, simply by dividing the amplitude of the evoked EPP by the mean amplitude of the MEPP (the so-called ‘direct method’). This is because quantal components sum nonlinearly—that is, the depolarization (i.e., the EPP) caused by instantaneous action of the contents of, say, 50 vesicles on ACh receptors is much less than 50 times the depolarization caused by a single vesicle, each of which alone would produce a depolarization equivalent to a MEPP. To illustrate this, suppose the average amplitude of a series of MEPPs recorded at an NMJ is 1 mV. An EPP generated by two vesicles would produce an EPP of about 2 mV. But an EPP with a quantal content of 50 will not depolarize the endplate by 50 mV; it is more likely to be about 30 to 40 mV (depending on several other properties including the resting potential and other intrinsic biophysical properties of the muscle fiber). Nonlinear summation arises principally because the effect of ACh is to gate the nonselective cationic permeability of the ion channel that is integral to the receptor. Thus, in the theoretical limit, an infinite amount of ACh (or an infinitely high quantal content) could only produce depolarization to the net equilibrium potential (“reversal potential,” or “transmitter null potential”) for the permeant ions. This theoretical limiting membrane potential is about −5 mV (inside negative). Equations have been derived that compensate for nonlin-

ear summation before estimating quantal content, but none is entirely satisfactory, and the best way to avoid this problem altogether is to voltage-clamp the endplate, normally using a two-electrode method, and to measure miniature and evoked synaptic currents, which sum linearly (McLachlan and Martin, 1981; Wood and Slater, 1997; Thyagarajan et al., 2010). However, this technique is more difficult than single-microelectrode recording, limiting the number of recordings in any one session, and, thus, the yield of data is correspondingly smaller. McLachlan and Martin’s formula for correcting an observed mean EPP amplitude, v, for nonlinear summation is:

v′ =

v 1 − fv / ( Er − Em) Equation 2

where Er is the ACh null potential, Em is the resting membrane potential, and f is the ‘fudge’ factor necessary to align the endplate current with the EPP as a function of transmitter released. One way to avoid the problem of nonlinear summation altogether is to reduce the amount of transmitter released by a presynaptic action potential (i.e., quantal content) to small multiples of the quantal size. This is most simply achieved by reducing the extracellular Ca2+ concentration below its normal physiological level of 2 mM (e.g., to 1 mM) or increasing the extracellular Mg2+ from its normal level of 1 mM (e.g., to 4 mM), or both. Under these conditions, the probabilistic nature of exocytosis can be observed from the marked fluctuation of EPP amplitudes—from no response (‘failure’) to quantal multiples up to 10 times the mean MEPP amplitude. As originally calculated by Katz and his colleagues, the distribution of EPP amplitudes (and therefore, quantal contents) should resemble a binomial distribution:

P ( x) =

n! (n− x) pxq x ! (n − x) ! Equation 3

where the quantities n, p, and q (= 1 – p) represent number of quanta (or release sites) available for release and their individual release probabilities. Both quantities represent unknowns that cannot be measured independently. However, in the limiting case where probability of observing individual components of the EPP is low and these components occur independently of one another, then

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the binomial distribution reduces to a Poisson distribution:

P ( x) =

e− m m x x!

Equation 4

where the number of quantal components of any given EPP, x, can take any integer value or zero, and m is the mean quantal content. This equation can be exploited in various ways in practice to estimate quantal content. For instance, the method of failures is based on a straightforward count of the number of times there is no response (‘failure’) during in a series of test stimuli. No amplitude measurements on either EPPs or MEPPs are necessary because, according to the Poisson equation, when x = 0:

P (0) = exp (− m ) Equation 5

and, since P(0) is simply the ratio of failures to tests, taking natural logarithms of both sides of the equation and re-arranging for m yields:

m = Ln ( Tests/Failures) Equation 6

This is usually in very good agreement with estimates made directly by dividing the mean EPP amplitude by the mean MEPP amplitude. So, for instance, if the mean MEPP amplitude is 1 mV and the mean EPP amplitude 3.5 mV, then 3.5 would be the mean quantal content. From the above formula, this mean quantal content would predict that there should be about three ‘failures’ in a train of responses to 100 stimuli. Put the other way, if there were three failures in response to 100 stimuli, then, according to the above formula, the mean quantal content would be 3.51. Another method based takes advantage of the equivalence of mean and variance in the Poisson distribution and hence the variance method for calculating mean quantal content from measurements of the amplitudes of a train of EPPs:

⎛v ⎞ m=⎜ ⎟ σ ⎝ v⎠

2

Equation 7 Quantal Analysis of Endplate Potentials in Mouse FDB Muscle

where is the mean EPP amplitude and sv the standard deviation. Like the method of failures, one benefit of the variance method is that it is unnecessary to

have accurate (or any) measurements of MEPP amplitude. The above methods for calculating quantal content are somewhat oversimplified, although adequate for most comparative purposes: for instance, the formulae given do not take account of high frequency stochastic fluctuations (membrane or electrode ‘noise’) in the recordings. Care should also be taken when applying this method since the assumptions underlying the applicability of a Poisson distribution may not be valid under normal physiological conditions of transmitter release. For a more advanced discussion, see for example Byrne and Roberts (2009) or monographs and review articles on these issues (Hubbard et al., 1969; Christensen and Martin, 1970; Johnson and Wernig, 1971; Bennett and Robinson, 1990; Cooper et al., 1995; Clements and Silver, 2000; Silver, 2003). Neuromuscular ‘safety factor’ and synaptic homeostasis Estimates of MEPP frequency, amplitude (quantal size), and EPP amplitude (and quantal content) are useful for estimating the ‘safety factor’ for neuromuscular transmission: that is, the excess of neurotransmitter released (or quantal content) over that required to depolarize the muscle fiber sufficiently for action potential generation and muscle contraction. Normally, at least in rodent muscle, the safety factor is about 3 to 5. In conditions where either neurotransmitter is reduced (such as the Lambert-Eaton Myasthenic Syndrome; or mild botulism) or the sensitivity to neurotransmitter is compromised (as in classical myasthenia gravis; or ‘curarization’ during surgical anesthesia), then the safety factor for neuromuscular transmission may be reduced by a significant margin, producing muscle weakness or flaccid paralysis (Wood and Slater, 1997; Slater, 2008). Under some chronic conditions of this nature, poorly understood ‘homeostatic’ mechanisms are activated to restore the safety factor, either by up-regulating post-synaptic sensitivity (quantal size) or by up-regulating quantal content (Plomp et al., 1994, 1995; Plomp and Molenaar, 1996). Conversely, but only demonstrated convincingly so far by genetic manipulations in Drosophila, up-regulation of one facet of synaptic transmission (e.g., quantal size) in some instances causes a compensatory down-regulation of the complementary quantity (i.e., quantal content), maintaining the overall amplitude of the evoked voltage response, that is, the EPP

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amplitude (Frank et al., 2006; Bergquist et al., 2010). Another important factor influencing the magnitude of endplate depolarization is the ‘input resistance’ of the muscle fiber, which is determined by its dimensions, myoplasmic electrical impedance, and passive membrane ionic permeability. Other things being equal, relatively large diameter muscle fibers have a lower input resistance than relatively small diameter muscle fibers. Assuming that the number of molecules of ACh in one presynaptic vesicle is largely independent of the target muscle fiber size (or input resistance), then the endplate current for a given quantal content will be the same on small- or large-diameter muscle fibers (but see Wilkinson et al., 1992). However, by Ohm’s law, the depolarizations (i.e., MEPPs) will depend on the input resistance (V = IRin ), so the MEPPs recorded from large-diameter muscle fibers will be smaller in mean amplitude than those recorded from a more slender muscle fiber that has a higher input resistance. Again, there is evidence that homeostatic mechanisms bring about comparability in the mean size of the evoked response. This is at least partly achieved by proportional variation in the area of synaptic contact; motor terminals on large diameter muscle fibers occupy a proportionally larger area of the muscle fiber surface at the endplate than those on smaller diameter fibers. Since the quantal content of EPPs is determined partly by the density and number of active zones, then the larger the area of synaptic contact, the greater the quantal content of the EPPs. Large endplates are found on relatively largediameter fibers, and so these generally have a higher quantal content than neuromuscular junctions of smaller area typically found on small-diameter fibers. This relationship therefore compensates for the inverse effect of fiber diameter on quantal size and serves to maintain the average amplitude of the EPP, and therefore the safety factor for neuromuscular transmission, independently of muscle fiber size (Harris and Ribchester, 1979a,b; Ribchester et al., 2004; Slater, 2008). The packing density and length of the postsynaptic folds at NMJ also affects the safety factor for neuromuscular transmission by altering the current density in the synaptic cleft (Martin, 1994; Wood and Slater, 1997). A high packing density of ACh receptors at the crests of the junctional folds and the voltage-gated Na-channels that normally occupy the crypts of the folds thus bring about a higher safety

factor for transmission for a given area of synaptic contact and muscle fiber input resistance. This enables, for instance, the characteristically small NMJs of some human muscle fibers (which have a low quantal content, but a high junctional fold density) to nonetheless sustain an adequate safety factor for neuromuscular transmission, with no overt signs of muscle weakness (Slater et al., 1992, 2006). Termination of ACh action The molecular effects of ACh on their receptors are either intercepted or terminated by the action of the hydrolytic enzyme acetylcholinesterase (AChE), embedded in the synaptic basal lamina (Massoulie and Millard, 2009). The affinity and potency of this enzyme ensures that only about 50% of the molecules released into the synaptic cleft cross the synaptic basal lamina to binding sites on their postsynaptic receptors. Unbound ACh molecules that detach from receptors are swiftly hydrolyzed as well. In the presence of inhibitors of AChE, termination of transmitter efficacy is accomplished by a slower process of diffusion of ACh from the synaptic cleft. The main discernible indicator of this is the prolonged time course of repolarization during the EPP (Magleby and Terrar, 1975).

Troubleshooting 1. Problem: The skin does not strip easily from the foot at the outset of dissection. This is common in mice older than about 4 months; or in younger mice if the skin is tugged too brusquely. The skin can be dissected away surgically if the stripping method does not work. 2. Problem: The isolated FDB muscle does not contract in response to nerve stimulation. This can arise if the nerve is stretched or damaged, blocking nerve conduction; or if there has been an error in the making-up of the physiological saline; or if the stimulator is not working (check the batteries and make sure that it is switched on); or the triggering device has not been set up correctly; or the wires are not connected correctly to the stimulator; or there is insufficient fluid contacting the internal and/or external wires of the suction electrode; or the stimulating voltage is insufficient in magnitude or duration. 3. Problem: The microelectrode signal cannot be adjusted to zero before fiber penetration. Check that the bath ground and the microelectrode tip are in the bathing medium, and that the amplifier is turned on. Sometimes this

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error can result from an electrode resistance that is too high (>200 M), for example, if the electrolyte in the electrode is wrong or the pipet tip is too narrow. Test this by breaking the electrode tip and submerging the broken end into the bathing medium. It should now be easy to ‘zero’ the trace. If not, there may be a fault in the headstage or amplifier. 4. Problem: The noise level is too great. If the noise is of very high frequency (a thick baseline of 500 μV peak-to-peak or greater) this can be due to poor contacts between the bath (earth/ground) electrode or the connection to the microelectrode, or an excessively high electrode resistance (>200 M). High-amplitude noise (>1 mV peak to peak) at mains frequency (50 Hz or 60 Hz) is normally due to imperfect grounding. Ensure all metal objects in the vicinity of the recording electrode are connected to a common earth/grounding point. If the problem persists, use a Humbug filter to remove the residual noise, although it is better to remove the source of the ground loops directly. 5. Problem: The resting membrane potentials are less negative than expected. This arises if the muscle has been damaged during dissection, if the microelectrodes are too blunt (check the electrode resistance, it should be greater than 10 M), or if the potassium concentration in the bathing medium is higher than it should be. 6. Problem: The resting membrane potential is initially quite high (about −65 mV) but then drifts steadily to less negative values. This may occur when the electrode is too blunt to produce a clean penetration of the target muscle fiber, or significant amounts of chloride ion are leaking from the micropipet tip into the muscle cytoplasm. Check the tip resistance of the microelectrode (it should be about 10 M or higher). Change the microelectrode if the problem persists with successive fiber impalements, or change the electrolyte used to fill the electrodes from 3 M KCl to 4 M potassium acetate. 7. Problem: There are no MEPPs. This can arise if the temperature is low (MEPP frequency is temperature sensitive), or if the resting membrane potentials are very low (less negative than −40 mV), or if there is an acetylcholine receptor blocker, such as d-tubocurarine or α-bungarotoxin present, or if there was some pathological feature of the mouse from which the dissected muscle was obtained.

8. Problem: There are no evoked responses to nerve stimulation, or the evoked responses are much smaller than expected. This may arise if the electrode has not in fact penetrated the muscle fiber (bending of the electrode tip can produce a spurious voltage deflection that resembles membrane penetration), or if the Ca2+ /Mg2+ ratio in the bathing fluid is too low, or if the nerve is not responding to stimulation (see point 2, above), or if there was some pathological feature of the mouse from which the dissected muscle was obtained. 9. Problem: Quantal analysis gives a large disparity between failures, variance, and direct methods for estimating quantal content. This arises if the analysis of quantal content does not take account of the requirement for a stationary mean value around which the random quantal fluctuations vary (check the graph of amplitude against record number), or an incorrect entry has been made for MEPP amplitude, or the value entered for the nonlinear summation ‘fudge’ factor (f) is incorrect or inappropriate, or if the membrane noise levels exceed the quantal variations in the evoked response, or if uniquantal (MEPP) responses are buried in noise, leading to overestimates of the number of ‘failures’; it may also be the result of some pathological feature of the mouse from which the dissected muscle was obtained.

Anticipated Results A photomicrograph of an isolated FDB muscle and representative recordings of EPPs and MEPPs are shown in Figures 1 and 2. Figure 1 shows an isolated FDB muscle and typical records of several EPPs superimposed, produced by repetitive stimulation after blocking muscle action potentials with μ-conotoxin GIIIB. Figure 2 shows typical spontaneous MEPPs and evoked responses with reduced extracellular Ca2+ ions and elevated Mg2+ ions, indicating typical quantized fluctuations and ‘failures.’

Time Considerations 1. Making solutions may take about 30 min. 2. Dissection time by a skilled operator, from killing of the mouse to mounting an FDB nerve-muscle preparation in a recording chamber, can be as little as 15 to 30 min. 3. Making microelectrodes and recording from thirty muscle fibers sufficient for a quantal analysis may take 1 to 4 hr. 4. Offline analysis of quantal size and quantal content from 30 fibers recorded in one muscle may take 1 to 4 hr.

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Acknowledgments The University of Edinburgh is a charitable body, registered in Scotland, with registration number SC005336.

Literature Cited Angaut-Petit, D., Molgo, J., Connold, A.L., and Faille, L. 1987. The levator auris longus muscle of the mouse: A convenient preparation for studies of short- and long-term presynaptic effects of drugs or toxins. Neurosci. Lett. 82:83-88. Auerbach, A. and Betz, W. 1971. Does curare affect transmitter release? J Physiol. 213:691-705. Bekoff, A. and Betz, W.J. 1977a. Properties of isolated adult rat muscle fibres maintained in tissue culture. J. Physiol. 271:537-547. Bekoff, A. and Betz, W.J. 1977b. Physiological properties of dissociated muscle fibres obtained from innervated and denervated adult rat muscle. J. Physiol. 271:25-40. Bennett, M.R. and Robinson, J. 1990. Probabilistic secretion of quanta from nerve terminals at synaptic sites on muscle cells: Non-uniformity, autoinhibition and the binomial hypothesis. Proc. R Soc. Lond. B Biol. Sci. 239:329-358. Bergquist, S., Dickman, D.K., and Davis, G.W. 2010. A hierarchy of cell intrinsic and target-derived homeostatic signaling. Neuron 66:220-234. Betz, W.J. and Bewick, G.S. 1993. Optical monitoring of transmitter release and synaptic vesicle recycling at the frog neuromuscular junction. J. Physiol. 460:287-309. Blioch, Z.L., Glagoleva, I.M., Liberman, E.A., and Nenashev, V.A. 1968. A study of the mechanism of quantal transmitter release at a chemical synapse. J. Physiol. 199:11-35. Braga, M.F., Anderson, A.J., Harvey, A.L., and Rowan, E.G., 1992. Apparent block of K+ currents in mouse motor nerve terminals by tetrodotoxin, mu-conotoxin and reduced external sodium. Br. J. Pharmacol. 106:91-94. Byrne, J.H. and Roberts, J.L. 2009. From Molecules to Networks, 2nd ed. Elsevier, Amsterdam. Christensen, B.N. and Martin, A.R. 1970. Estimates of probability of transmitter release at the mammalian neuromuscular junction. J. Physiol. 210:933-945. Clements, J.D. and Silver, R.A. 2000. Unveiling synaptic plasticity: A new graphical and analytical approach. Trends Neurosci. 23:105-113. Cooper, R.L., Stewart, B.A., Wojtowicz, J.M., Wang, S., and Atwood, H.L. 1995. Quantal measurement and analysis methods compared for crayfish and Drosophila neuromuscular junctions, and rat hippocampus. J. Neurosci. Methods 61:67-78. Curtis, M.J., Quastel, D.M., and Saint, D.A. 1986. Lanthanum as a surrogate for calcium in transmitter release at mouse motor nerve terminals. J. Physiol. 373:243-260.

DiFranco, M., Tran, P., Quinonez, M., and Vergara, J.L. 2011. Functional expression of transgenic 1sDHPR channels in adult mammalian skeletal muscle fibres. J. Physiol. 589:1421-1442. Donovan, J. and Brown, P. 2006. Euthanasia. Curr. Protoc. Immunol. 73:1.8.1-1.8.4. Frank, C.A., Kennedy, M.J., Goold, C.P., Marek, K.W., and Davis, G.W. 2006. Mechanisms underlying the rapid induction and sustained expression of synaptic homeostasis. Neuron 52:663-677. Gillespie, J.I. and Ribchester, R.R. 1988. Optical measurements of intracellular pH in intact isolated muscle fibres and muscle growth cones in culture. Q. J. Exp. Physiol. 73:995-1000. Gillingwater, T.H., Thomson, D., Mack, T.G., Soffin, E.M., Mattison, R.J., Coleman, M.P., and Ribchester, R.R. 2002. Age-dependent synapse withdrawal at axotomised neuromuscular junctions in Wld(s) mutant and Ube4b/Nmnat transgenic mice. J. Physiol. 543:739-755. Harris, J.B. and Ribchester, R.R. 1979a. Pharmacological aspects of neuromuscular transmission in the isolated diaphragm of the dystrophic (Rej 129) mouse. Br. J. Pharmacol. 65:411-421. Harris, J.B. and Ribchester, R.R. 1979b. The relationship between end-plate size and transmitter release in normal and dystrophic muscles of the mouse. J. Physiol. 296:245-265. Hubbard, J.I., Llin´as, R., and Quastel, D.M.J. 1969. Electrophysiological analysis of synaptic transmission. Monogr. Physiol. Soc. no. 19. Johnson, E.W. and Wernig, A. 1971. The binomial nature of transmitter release at the crayfish neuromuscular junction. J. Physiol. 218:757-767. Katz, B. 1969. The Release of Neural Transmitter Substances. Liverpool University Press, Liverpool, U.K. Katz, B. 1996. Neural transmitter release: From quantal secretion to exocytosis and beyond. The Fenn Lecture. J. Neurocytol. 25:677-686. Liu, Y., Sugiura, Y., and Lin, W. 2011. The role of Synaptobrevin1/VAMP1 in Ca2+ -triggered neurotransmitter release at the mouse neuromuscular junction. J. Physiol. 589:1603-1618. Livet, J., Weissman, T.A., Kang, H., Draft, R.W., Lu, J., Bennis, R.A., Sanes, J.R., and Lichtman, J.W. 2007. Transgenic strategies for combinatorial expression of fluorescent proteins in the nervous system. Nature 450:56-62. Lu, J., Tapia, J.C., White, O.L., and Lichtman, J.W. 2009. The interscutularis muscle connectome. PLoS Biol. 7:e32. Lupa, M.T. and Caldwell, J.H. 1991. Effect of agrin on the distribution of acetylcholine receptors and sodium channels on adult skeletal muscle fibers in culture. J. Cell Biol. 115:765-778. Magleby, K.L. and Terrar, D.A. 1975. Factors affecting the time course of decay of end-plate currents: A possible cooperative action of acetylcholine on receptors at the frog neuromuscular junction. J. Physiol. 244:467-495.

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Martin, A.R. 1994. Amplification of neuromuscular transmission by postjunctional folds. Proc. Biol. Sci. 258:321-326. Massoulie, J. and Millard, C.B. 2009. Cholinesterases and the basal lamina at vertebrate neuromuscular junctions. Curr. Opin. Pharmacol. 9:316-325. McArdle, J.J., Angaut-Petit, D., Mallart, A., Bournaud, R., Faille, L., and Brigant, J.L. 1981. Advantages of the triangularis sterni muscle of the mouse for investigations of synaptic phenomena. J. Neurosci. Methods 4:109-115. McLachlan, E.M. and Martin, A.R. 1981. Nonlinear summation of end-plate potentials in the frog and mouse. J. Physiol. 311:307-324. Nagwaney, S., Harlow, M.L., Jung, J.H., Szule, J.A., Ress, D., Xu, J., Marshall, R.M., and McMahan, U.J. 2009. Macromolecular connections of active zone material to docked synaptic vesicles and presynaptic membrane at neuromuscular junctions of mouse. J. Comp. Neurol. 513:457-468. Nocella, M., Colombini, B., Benelli, G., Cecchi, G., Bagni, M.A., and Bruton, J.D. 2011. Force decline during fatigue is due to both a decrease in the force per individual cross-bridge and the number of cross-bridges. J. Physiol. 589:33713381.

Ruiz, R., Cano, R., Casanas, J.J., Gaffield, M.A., Betz, W.J., and Tabares, L. 2011. Active zones and the readily releasable pool of synaptic vesicles at the neuromuscular junction of the mouse. J. Neurosci. 31:2000-2008. Silver, R.A. 2003. Estimation of nonuniform quantal parameters with multiple-probability fluctuation analysis: Theory, application and limitations. J. Neurosci. Methods. 130:127-141. Slater, C.R. 2008. Reliability of neuromuscular transmission and how it is maintained. Handb. Clin. Neurol. 91:27-101. Slater, C.R., Lyons, P.R., Walls, T.J., Fawcett, P.R., and Young, C. 1992. Structure and function of neuromuscular junctions in the vastus lateralis of man: A motor point biopsy study of two groups of patients. Brain 115:451-478. Slater, C.R., Fawcett, P.R., Walls, T.J., Lyons, P.R., Bailey, S.J., Beeson, D., Young, C., and Gardner-Medwin, D. 2006. Pre- and postsynaptic abnormalities associated with impaired neuromuscular transmission in a group of patients with ‘limb-girdle myasthenia’. Brain 129:2061-2076.

Plomp, J.J. and Molenaar, P.C. 1996. Involvement of protein kinases in the upregulation of acetylcholine release at endplates of alphabungarotoxin-treated rats. J. Physiol. 493:175186.

Thyagarajan, B., Potian, J.G., Garcia, C.C., Hognason, K., Capkova, K., Moe, S.T., Jacobson, A.R., Janda, K.D., and McArdle, J.J. 2010. Effects of hydroxamate metalloendoprotease inhibitors on botulinum neurotoxin A poisoned mouse neuromuscular junctions. Neuropharmacology 58:1189-1198.

Plomp, J.J., van Kempen, G.T., and Molenaar, P.C. 1994. The upregulation of acetylcholine release at endplates of alpha-bungarotoxin-treated rats: Its dependency on calcium. J. Physiol. 478:125136.

Urbano, F.J., Rosato-Siri, M.D., and Uchitel, O.D. 2002. Calcium channels involved in neurotransmitter release at adult, neonatal and P/Q-type deficient neuromuscular junctions (Review). Mol. Membr. Biol. 19:293-300.

Plomp, J.J., Van Kempen, G.T., De Baets, M.B., Graus, Y.M., Kuks, J.B., and Molenaar, P.C. 1995. Acetylcholine release in myasthenia gravis: Regulation at single end-plate level. Ann. Neurol. 37:627-636.

Wilkinson, R.S., Lunin, S.D., and Stevermer, J.J. 1992. Regulation of single quantal efficacy at the snake neuromuscular junction. J. Physiol. 448:413-436.

Ribchester, R.R. 2009. Mammalian neuromuscular junctions: modern tools to monitor synaptic form and function. Curr. Opin. Pharmacol. 9:297-305. Ribchester, R.R., Tsao, J.W., Barry, J.A., AsgariJirhandeh, N., Perry, V.H., and Brown, M.C. 1995. Persistence of neuromuscular junctions after axotomy in mice with slow Wallerian degeneration (C57BL/WldS). Eur. J. Neurosci. 7:1641-1650. Ribchester, R.R., Thomson, D., Haddow, L.J., and Ushkaryov, Y.A. 1998. Enhancement of spontaneous transmitter release at neonatal mouse neuromuscular junctions by the glial cell linederived neurotrophic factor (GDNF). J. Physiol. 512:635-641. Quantal Analysis of Endplate Potentials in Mouse FDB Muscle

Huntington’s disease mutation. Eur. J. Neurosci. 20:3092-3114.

Ribchester, R.R., Thomson, D., Wood, N.I., Hinks, T., Gillingwater, T.H., Wishart, T.M., Court, F.A., and Morton, A.J. 2004. Progressive abnormalities in skeletal muscle and neuromuscular junctions of transgenic mice expressing the

Wong, F., Fan, L., Wells, S., Hartley, R., Mackenzie, F.E., Oyebode, O., Brown, R., Thomson, D., Coleman, M.P., Blanco, G., and Ribchester, R.R. 2009. Axonal and neuromuscular synaptic phenotypes in Wld(S), SOD1(G93A) and ostes mutant mice identified by fiber-optic confocal microendoscopy. Mol. Cell Neurosci. 42:296-307. Wood, S.J. and Slater, C.R. 1997. The contribution of postsynaptic folds to the safety factor for neuromuscular transmission in rat fast- and slow-twitch muscles. J. Physiol. 500:165-176. Yeung, E.W., Balnave, C.D., Ballard, H.J., Bourreau, J.P., and Allen, D.G. 2002. Development of T-tubular vacuoles in eccentrically damaged mouse muscle fibres. J. Physiol. 540:581592. Zitman, F.M., Todorov, B., Jacobs, B.C., Verschuuren, J.J., Furukawa, K., Furukawa, K., Willison, H.J., and Plomp, J.J. 2008. Neuromuscular synaptic function in mice lacking major subsets of gangliosides. Neuroscience 156:885897.

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Exploration of the Visual System: Part 1: Dissection of the Mouse Eye for RNA, Protein, and Histological Analyses Pascal Escher1,2 and Daniel F. Schorderet1,2,3 1

IRO-Institute for Research in Ophthalmology, Sion, Switzerland Department of Ophthalmology, University of Lausanne, Lausanne, Switzerland 3 EPFL-Ecole Polytechnique F´ed´erale, Lausanne, Switzerland 2

ABSTRACT Due to the power of genetics, the mouse has become a widely used animal model in vision research. However, its eyeball has an axial length of only about 2 mm. The present protocol describes how to easily dissect the small rodent eye post mortem. This allows collecting different tissues of the eye, i.e., cornea, lens, iris, retina, optic nerve, retinal pigment epithelium (RPE), and sclera. We further describe in detail how to process these eye samples in order to obtain high-quality RNA for RNA expression profiling studies. Depending on the eye tissue to be analyzed, we present appropriate lysis buffers to prepare total protein lysates for immunoblot and immuno-precipitation analyses. Fixation, inclusion, embedding, and cryosectioning of the globe for routine histological analyses (HE staining, DAPI staining, immunohistochemistry, in situ hybridization) is further presented. These basic protocols should allow novice investigators to obtain eye tissue C 2011 by samples rapidly for their experiments. Curr. Protoc. Mouse Biol. 1:445-462  John Wiley & Sons, Inc. Keywords: ophthalmology r visual sciences r retina r cornea r lens r iris r retinal pigment epithelium (RPE)

INTRODUCTION Despite a small eye size and reduced visual function, vision research in mice offers several advantages: (1) the mouse genome is available and annotated in detail (http://www.ensembl.org/Mus musculus/Info/Index); (2) by gene targeting and gene trapping every single protein-coding gene is being mutated (http://www.knockoutmouse.org); (3) standard inbred and recombinant inbred strains are being characterized (http://www.jax.org/phenome); (4) tissues are available for studies from embryonic timepoints on; (5) evolution of disease is more rapid in mice because of their 2- to 3-year life span; (6) diet and environment can be precisely controlled; and (7) space requirements for housing are limited (Smith et al., 2002). The rising interest in mice for visual studies prompted us to describe in the present unit basic protocols for the dissection of the mouse eye. Specifically, isolation of cornea, iris, lens, retina, RPE, and the optic nerve are described (Basic Protocols 1 and 2 and Support Protocol 1). Total RNA and proteins prepared from these eye tissues are suitable for downstream applications, such as RT-PCR, quantitative PCR, and immunoblotting (Basic Protocols 3 and 4 and Alternate Protocol). Additionally, fixation, inclusion, embedding, and cryosectioning of the mouse eyeball are presented (Basic Protocol 5). Cryosections are suitable for hematoxylin-eosin staining (Basic Protocol 6), DAPI staining (Basic Protocol 7), immunohistochemical analyses (Basic Protocol 8), and in situ hybridizations (Support Protocol 2).

Current Protocols in Mouse Biology 1: 445-462, December 2011 Published online December 2011 in Wiley Online Library (wileyonlinelibrary.com). DOI: 10.1002/9780470942390.mo110129 C 2011 John Wiley & Sons, Inc. Copyright 

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NOTE: To ensure successful experiments, dissection and processing of tissues should be performed under clean, sterile conditions, particularly if RNA analysis will be performed. NOTE: All protocols using live animals must first be reviewed and approved by an Institutional Animal Care and Use Committee (IACUC) and must follow officially approved procedures for care and use of laboratory animals. BASIC PROTOCOL 1

ENUCLEATION OF THE MOUSE EYE This protocol provides information necessary to enucleate the eye of a mouse properly. The steps are written under the assumption that the user will immediately proceed to dissection and downstream processing; dissection is detailed in Basic Protocol 2 and Basic Protocols 3 to 8 provide instructions for further sample preparation and experimental procedures.

Materials Mice at the specific age needed for the study 70% (v/v) ethanol IACUC-approved animal housing facility IACUC-approved animal cages and bedding IACUC-approved food and water Graefe forceps, 1-mm tip, straight (Fine Science Tools) Extra fine Bonn scissors, 8.5 cm, straight (Fine Science Tools) Additional reagents and equipment for euthanizing the mice (Donovan and Brown, 2006)

Dissection of the Mouse Eye for RNA, Protein, and Histological Analyses

Figure 1 Enucleation (Basic Protocol 1, step 4). Pull the eyeball gently off the orbit, by pressing with the fingers around the orbit and pulling them apart.

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Figure 2 Enucleation (Basic Protocol 1, step 5). Enucleate the eye by gently holding the eyeball with forceps and cutting the optic nerve with scissors at a distance of ∼2 mm from the eyeball.

1. House the mice in a 12 hr:12 hr light-dark cycle with unlimited access to food and water. To limit light-induced retinal degeneration, light conditions should be adapted for lightsensitive strains, e.g., albino mice.

2. Euthanize the mice by cervical dislocation (see Donovan and Brown, 2006). Regulations for euthanasia may vary and carbon dioxide (CO2 ) inhalation enforced.

3. Soak the head in 70% ethanol. This sterilization step decreases the number of hairs interfering with subsequent dissection.

4. Pull the eyeball gently off the orbit, by pressing with the fingers around the orbit and pulling them apart (Fig. 1). A nasal and temporal incision may be necessary to open the eyelids and to push the eyeball off the orbit.

5. Enucleate the eye by gently holding the eyeball with forceps and cutting the optic nerve with a scissor at a distance of about 2 mm from the eyeball (Fig. 2). Avoid crushing the optic nerve directly behind the eyeball, as this may cause myelin of the nerve to enter the eye (myelin artifact).

DISSECTION OF THE MOUSE EYE This protocol provides information necessary to properly dissect the eye of a mouse. Depending on the downstream applications, dissected tissues need to be immediately homogenized according to Basic Protocols 3, 4, and 5.

BASIC PROTOCOL 2 Dissection of the Mouse Eye for RNA, Protein, and Histological Analyses

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Materials SYLGARD 184-filled 60- or 100-mm cell culture plates (see Support Protocol 1) Eyeball (see Basic Protocol 1) Phosphate-buffered saline (PBS; see recipe) Cold-light source, e.g., KL200 (Schott/Zeiss) Stereomicroscope, e.g., Leica 16MZF, Leica M80, Zeiss Stemi 2000 Graefe forceps, 1-mm tip, straight (Fine Science Tools) 0.2 mm Minutien pins, length 10 mm (Fine Science Tools) Dumont #3c forceps (Fine Science Tools) Vannas spring scissors 2- to 4-mm blades (Fine Science Tools) Extra fine Graefe forceps, 0.5-mm tip, straight (Fine Science Tools)

Figure 3 Dissection of the eye (Basic Protocol 2, step 2). Hold the eyeball with forceps on the cell culture dish and immobilize with Minutien pins, pinning through the sclera and extraocular tissues close to the optic nerve (asterisk).

rectus tendons

retina cornea

optic nerve lens

sclera

iris

choroid RPE

Dissection of the Mouse Eye for RNA, Protein, and Histological Analyses

ora serrata

Figure 4 Schematic representation of the mouse eye. The size of the eyeball is ∼2 mm. The major components of the eye are indicated. Please note the reduced volume of the vitreous in the mouse eye. Abbreviations: RPE: retinal pigment epithelium.

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1. Place a SYLGARD 184-filled 60-mm or 100-mm cell culture plate under a stereomicroscope equipped with an external cold-light source Support Protocol 1 provides instructions for preparation of SYLGARD 184-filled plates.

2. Hold the eyeball (from the enucleated eye, see Basic Protocol 1) with forceps, e.g., Graefe forceps, on the cell culture dish and immobilize with pins (Fig. 3). Using forceps, e.g., Dumont #3c forceps, plant two to three pins into the sclera, close to the optic nerve and to remaining extraocular muscle tissue.

3. Cover the eyeball with 1× PBS. 4. Section the eyeball with spring scissors just posterior to the tendons of the rectus muscles, visible as a whitish strip on the sclera. See Figure 4 for a schematic representation of the mouse eye. This section will divide the eyeball into an anterior and posterior part, along an equatorial plane that corresponds inside the eyeball to the ora serrata/pars plana.

5. Remove the cornea and attached iris with forceps (Fig. 5). The iris detaches readily from the anterior segment and the cornea can be further cleaned by cutting and taking off foreign tissue with extra fine forceps and spring scissors.

Figure 5 Dissection of the eye (Basic Protocol 2, steps 5 to 7). After dissection of the mouse eye, several tissues can be isolated. The transparent lens (A) often has iris tissue attached to the equator plane (asterisk). The yellowish retina (B) too, may have attached iris tissue (asterisk). Dark pigmentation is further due to apical foldings of the RPE that are attached to the retina. The posterior optic cup comprises the melanized RPE and choriocapillaris, as well as the grayish sclera (C). The white optic nerve extends form the eyeball (asterisk). With respect to the anterior segment, the melanized iris remains attached to the transparent cornea (D).

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6. Recover the lens (Fig. 5). Parts of the iris often remain attached to the lens equator and have to be pulled out with extra fine forceps or cut with spring scissors.

7. Detach the retina from the RPE and recover it by cutting the optic nerve with spring scissors (Fig. 5). Iris tissue attached to the retina can be easily removed with spring scissors by spotting dark tissue on the yellowish retina.

8. Recover the optic nerve. 9. Clean the posterior eyecup comprised of RPE, choriocapillaris, and sclera by removing muscle and other foreign tissues with extra fine forceps and spring scissors (Fig. 5). 10. Proceed with RNA and protein preparation (see Basic Protocols 3 and 4) SUPPORT PROTOCOL 1

PREPARATION OF SYLGARD 184 SILICONE ELASTOMER SYLGARD 184 silicone elastomer is a convenient support for dissections and for storing dissection instruments. SYLGARD-filled containers should be made ahead of time to expedite experiment workflow.

Materials SYLGARD 184 silicone elastomer kit (Dow Corning) 60-mm and 100-mm cell culture dishes 50-ml conical polypropylene centrifuge tubes, with rim 1. Prepare the cell culture dishes and centrifuge tubes to be filled. The elastomer is not toxic and polymerization can be done on the workbench.

2. Mix the two components of the SYLGARD 184 silicone elastomer kit at a 10:1 ratio. 3. Pour the liquid elastomer into the cell culture dishes and centrifuge tubes. Cell culture dishes should be filled to a thickness of ∼4 mm. This allows easy pinning of tissues and leaves enough volume to cover the tissues during dissection with an appropriate liquid. Fill 50-ml centrifuge tubes with ∼7 to 10 ml elastomer (greater volumes can be poured to store smaller dissection tools).

4. Let polymerize at room temperature. Polymerization is completed within ∼10 min at room temperature. Polymerized SYLGARD 184 silicone elastomer can be stored for years at room temperature. BASIC PROTOCOL 3

RNA PREPARATION FROM EYE TISSUES This protocol describes preparation of total RNA from eye tissues that can be used for RT-PCR, quantitative PCR, and other RNA profiling studies. NOTE: For RNA work, always wear gloves and use plastics from packages that have not been previously opened. All aqueous solutions have to be treated overnight with DEPC (di-ethyl-pyrocarbonate) and then autoclaved.

Dissection of the Mouse Eye for RNA, Protein, and Histological Analyses

NOTE: As it has been previously, RPE in this section refers to the retinal pigment epithelium, and not to the commonly used Buffer RPE from the Qiagen RNA isolation kit.

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Materials Dissected eye (Basic Protocol 2) RNA isolation solution, e.g., TRI Reagent (Molecular Research Center) or TRIzol (Invitrogen) Chloroform RNase-free autoclaved water RNase-free 70% ethanol cDNA synthesis kit, e.g., First-strand cDNA synthesis kit for RT-PCR (AMV; Roche) RNase-free 1.5- and 2-ml microcentrifuge tubes Ventilated chemical hood 18-G Sterican needles (Braun) 1-ml syringes (Braun) FastPrep instrument (MP Biomedicals), optional Lysing Matrix D tubes (MP Biomedicals), optional Benchtop centrifuge, refrigerated 1. Place the cornea, iris, lens, optic nerve, and retina samples, dissected in Basic Protocol 2, in an RNase-free 2-ml microcentrifuge tube filled with 200 to 500 μl of TRIreagent (or similar RNA isolation solution). Work on ice under a ventilated chemical hood while using RNA isolation solution and chloroform.

2. Homogenize the tissues by passing the solution through 18-G Sterican needles placed on a 1-ml syringe. Homogenized samples can be stored in TRIReagent up to 1 year at −80◦ C. Alternatively, tissue homogenization can be done in a FastPrep instrument with greencapped Lysing Matrix D tubes (MP Biomedicals), using three 10-sec pulses with 20-sec intervals at 4◦ C, in a volume of 500 μl RNA isolation solution. RNA must then be extracted, precipitated, washed, resuspended, and quantitated according to the manufacturer’s instructions (proceed to step 7).

3. Place the cleaned posterior eyecup (Basic Protocol 2, step 9) comprising RPE, choriocapillaris and sclera, in a 2-ml microcentrifuge tube filled with 500 μl TRIreagent (or similar RNA isolation solution). 4. Dissociate the RPE and choriocapillaris from the sclera through homogenization with 18-G Sterican needles placed on a 1-ml syringe. 5. Separate RPE and choriocapillaris from the sclera by centrifuging 2 min at 1000 × g, 4◦ C. 6. Transfer the supernatant containing RPE and choriocapillaris into a new RNase-free 1.5-ml tube. RNA must then be extracted, precipitated, washed, resuspended, and quantitated according to the manufacturer’s instructions (proceed to step 7). This method yields total RNA of the RPE sufficiently pure for semi-quantitative analyses by RT-PCR. For quantitative PCR analysis, isolate pure RPE cells for total RNA preparation (see the Alternate Protocol).

7. Prepare total RNA from the different dissected eye tissues according to manufacturer’s instructions (e.g., TRI Reagent or TRIzol), with prolonged centrifugation times to increase RNA recovery. For total RNA preparation, chloroform, RNase-free autoclaved water, RNase-free 70% ethanol, and a refrigerated benchtop centrifuge must be prepared in advance. Current Protocols in Mouse Biology

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8. Generate cDNA from 100 to 500 ng of total RNA using a reverse transcriptase cDNA synthesis kit, such as First-strand cDNA synthesis kit for RT-PCR (AMV) or as required by a downstream application protocol. 9. Proceed with RNA analysis by PCR, quantitative PCR (e.g., on a Lightcycler 480II with Fast Start Universal SYBRGreen Master Mix, Roche), microarrays (e.g., Affymetrix), or other RNA expression profiling techniques. We routinely use the expression of the ribosomal protein L8 (RL8) as internal standard (Escher et al., 2005). ALTERNATE PROTOCOL

RNA PREPARATION FROM PURE RPE CELLS The protocol to isolate pure RPE cells for total RNA preparation is adapted from the one of the Streilein laboratory (Sugita and Streilein, 2003).

Materials Posterior eyecup (Basic Protocol 2, step 9) SYLGARD 184-filled 60-mm or 100-mm cell culture dish Vannas spring scissors 4 mm blades (Fine Science Tools) Trypsin Phosphate-buffered saline (PBS; see recipe) Cell culture medium RNA isolation solution, e.g., TRI Reagent (Molecular Research Center) or TRIzol (Invitrogen) Chloroform RNase-free autoclaved water RNase-free 70% ethanol 37◦ C, 5% CO2 incubator Extra fine Graefe forceps, 0.5-mm tip, straight (Fine Science Tools) RNase-free 2-ml microcentrifuge tubes RNase-free 200-μl pipet tips Benchtop centrifuge, refrigerated 1. Flatten the cleaned posterior eyecup (Basic Protocol 2, step 9) comprising RPE, choriocapillaris, and sclera, with three to four radial incisions. 2. Pin on a SYLGARD 184-filled 60 mm- or 100 mm-cell culture dish (see Support Protocol 1). 3. Cover the flattened eyecup with 0.2% trypsin made up in 1× PBS or serum-free mammalian cell culture medium. 4. Incubate for 30 to 60 min at 37◦ C in 5% CO2 atmosphere. 5. Gently peel off RPE cells with extra fine Graefe forceps. 6. Transfer pure RPE cells into a 2-ml microcentrifuge tube filled with 200 to 500 μl of TRIreagent or similar RNA isolation solution. 7. Homogenize by pipetting up and down using a 200-μl tip. 8. Proceed with total RNA preparation (see Basic Protocol 3, step 7) according to the manufacturer’s instructions (e.g., TRI Reagent or TRIzol). Dissection of the Mouse Eye for RNA, Protein, and Histological Analyses

For total RNA preparation, chloroform, RNase-free autoclaved water, RNase-free 70% ethanol, and a refrigerated benchtop centrifuge must be prepared in advance.

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PROTEIN PREPARATION FROM EYE TISSUES This protocol describes protein preparation from soft eye tissues, e.g., retina, RPE, iris, optic nerve and lens, as well as from cornea. This protein preparation is compatible with BCA protein quantification.

BASIC PROTOCOL 4

Materials Ice Tissue samples (see Basic Protocol 1 and 2 and the Alternate Protocol) RIPA buffer (see recipe) Tissue homogenization buffer (see recipe) 1.5-ml microcentrifuge tubes 18-G Sterican needles (Braun) 1-ml syringes (Braun) Plastic pestles adapted to 1.5-ml microcentrifuge tubes Microcentrifuge Standard equipment for immunoblotting Prepare for dissection 1. Prechill 1.5-ml microcentrifuge tubes on ice. 2. Dissect the eye according to Basic Protocols 1 and 2 and the Alternate Protocol.

Prepare protein from the cornea 3. Place the cornea into a prechilled microcentrifuge tube filled with 200 μl of ice-cold RIPA buffer. 4. Homogenize the tissues by passing the solution through 18-G Sterican needles placed on a 1-ml syringe.

Prepare protein from retina, iris, RPE, lens, and optic nerve 5. Place the retina, iris, RPE, lens, and optic nerve into a 1.5-ml prechilled microcentrifuge tube filled with 200 μl ice-cold tissue homogenization buffer. 6. Homogenize tissues with a plastic pestle. 7. Store all total protein lysates up to 3 years at −80◦ C until use.

Immunoblotting and Immuno-precipitation 8. Thaw the protein lysates. 9. Spin-down the thawed lysates for 2 min at maximal speed, room temperature, on a microcentrifuge. 10. Proceed with standard immunoblotting or immuno-precipitation protocols. For immunoblotting, we typically use 20 μg of protein extracts.

FIXATION, INCLUSION, AND CRYOSECTION OF THE MOUSE EYE This protocol describes how to prepare the enucleated mouse eye for cryosection. This protocol is adapted from the Yazulla laboratory (Eldred et al., 1983). NOTE: Cryosections obtained using this protocol are routinely used for in situ hybridizations. However, all aqueous solutions have to be treated overnight with DEPC (diethyl-pyrocarbonate) and then autoclaved. The previously described in situ hybridization protocol with DIG-labeled probes is well suitable for mouse eye analyses (Braissant and Wahli, 1998).

BASIC PROTOCOL 5

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Materials Eyeball (see Basic Protocol 1) Phosphate-buffered saline (PBS; see recipe), 1× 4% paraformaldehyde/1× PBS (see recipe) 30% sucrose/1× PBS (see recipe) Embedding medium, e.g., according to Yazulla (see recipe) Desiccating agent (e.g., drying pearls orange; Sigma) 1.5-ml microcentrifuge tubes Aluminum foil Cryostat, e.g., Leica CM1900 (Leica) SuperfrostPlus glass slides (Menzel) Heating block, optional Closed box for storing slides Perform fixation 1. Rinse the eyeball (see Basic Protocol 1) with 5 ml 1× PBS. 2. Transfer the eyeball into a 1.5-ml microcentrifuge tube filled with 500 μl of 4% paraformaldehyde/1× PBS. 3. Fix for a minimum of 2 hr, but up to overnight, at 4◦ C. For histological analyses, eyes are typically fixed overnight, but for immuno-histological analyses fixation does not exceed 6 hr.

Perform cryoprotection 4. Rinse the eyeball twice with 5 ml 1× PBS. 5. Transfer the eyeball into a 1.5-ml microcentrifuge tube filled with 500 μl of 30% sucrose/1× PBS. 6. Cryoprotect overnight at 4◦ C. In the beginning, the eyeball is floating on the surface, and then, sinking progressively to the bottom of the tube.

Perform embedding 7. Cut the caps of 1.5-ml microcentrifuge tubes to use as containers. Aluminum molds are commercially available.

8. Fill the caps with embedding medium at −20◦ C. Use the refrigerated chamber of the cryostat for these steps.

9. Place the eyeball into the embedding medium, orienting the eyeball so that a horizontal plane extending through the anterior segment and optic nerve is as parallel as possible with the bottom of the cap. Incisions and marks made before enucleation allow nasal-temporal and superior-inferior orientation of the eyeball.

10. Cover the oriented eyeball with embedding medium. 11. Freeze the embedded eyeball 5 min at −20◦ C. 12. Wrap in aluminum foil. Dissection of the Mouse Eye for RNA, Protein, and Histological Analyses

13. Store the embedded eyeball at −80◦ C until use. Embedded mouse eyes can be kept for years at −80◦ C, but freshly embedded eyes tend to be cut more easily.

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Cryosection the eyeballs 14. Remove the embedded eyeballs from the caps. 15. In the cryostat precooled to −23◦ C, cut the eyeballs at 10 to 12 μm and collect the cryosections on SuperfrostPlus glass slides. Up to 24 mouse eye sections can be put on a slide for histochemical analyses. For immunohistochemistry, four eye sections are typically put on a slide.

16. Dry the slides. Slides can be dried at 21◦ C overnight, or, alternatively, on a heating block at 50◦ C until the cryosections turn transparent. This takes a few minutes.

17. Store the slides at −20◦ C in a closed box containing a desiccating agent (e.g., drying pearls orange). Slides can be stored at 21◦ C for several weeks and up to 6 months at −20◦ C.

HEMATOXYLIN-EOSIN STAINING This protocol describes the classical hematoxylin-eosin staining. Hematoxylin is a basic dye that stains the nuclei in purple-blue. Eosin is an acidic dye staining cytoplasm in pink. The staining can be done directly on slides or in a 50-ml centrifuge tube. For staining series of slides at a time, use Coplin jars or similar staining stations.

BASIC PROTOCOL 6

Materials Slides with cryosections (see Basic Protocol 5) Phosphate-buffered saline (1×, PBS; see recipe) 4% paraformaldehyde in 1× PBS (see recipe) Hematoxylin solution, e.g., Accustain Hematoxylin solution, Gill No. 2 (Sigma) 70% ethanol/30% HCl 0.12 M solution (see recipe) Eosin solution, e.g., Accustain Eosin Y solution, alcoholic (Sigma) 50% glycerol/50% 1× PBS solution (see recipe) Nail polish Staining station, optional Coplin Jars or similar histology labware, optional Coverslips (Menzel) Light microscope 1. Equilibrate the slides with cryosections (see Basic Protocol 5) to room temperature.

Post-fix (optional) 2. Rinse the slides for 5 min with 1× PBS. When using Coplin jars or similar glassware, 100 ml solution should be prepared for each step. Alternatively, prepare 50 ml solution when using 50-ml centrifuge tubes. When processing only very few slides, milliliter amounts of solution can be directly pipetted onto the slides.

3. Fix the slides for 10 min at 4◦ C in 4% paraformaldehyde in 1× PBS.

Hematoxylin stain the slides 4. Hydrate the slides equilibrated to 21◦ C (see Basic Protocol 5) for 5 min in 1× PBS. 5. Stain for 10 min in hematoxylin staining solution. A blue-violet nuclear staining will start developing.

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6. Rinse for several seconds in a 70% ethanol/30 % HCl 0.12 M solution, until nuclear staining turns red.

Stain with eosin 7. Rinse the slides with tap water. 8. Stain the slides for 1 min in alcoholic eosin staining solution. 9. Rinse the slides with tap water. 10. Cover the sections with a drop of 50% glycerol/50% 1× PBS. 11. Cover the slide with coverslips and fix with nail polish.

Observe 12. Observe the slides using a light microscope. BASIC PROTOCOL 7

DAPI STAINING This protocol describes a fast and convenient fluorescent nuclear staining using DAPI (4 -6-diamidino-2-phenylindole dichloride). DAPI binds with high affinity to the minor groove of AT-rich DNA regions. Upon excitation by ultraviolet light (358 nm), a fluorescent signal at 461 nm is emitted, resulting in a blue nuclear staining.

Materials Slide with cryosections (see Basic Protocol 5) Phosphate-buffered saline (PBS) DAPI staining solution (see recipe) Cityfluor AF3 mounting medium (Cityfluor) Nail polish Coverslips Fluorescence microscope equipped with ultraviolet filter 1. Rehydrate the slides in 1× PBS. When using Coplin jars or similar glassware, 100 ml solution should be prepared for each step. Alternatively, prepare 50 ml solution when using 50-ml centrifuge tubes. When processing only very few slides, milliliter amounts of solution can be directly pipetted onto the slides.

2. Stain the slides in a 300 nM DAPI staining solution for 20 min at room temperature. 3. Wash the slides three times, each time for 5 min in 1× PBS. 4. Cover the sections with a drop of Cityfluor AF3. 5. Cover the slide with coverslips and fix with nail polish. 6. Image the slides using a fluorescence microscope equipped with ultraviolet filter. BASIC PROTOCOL 8

Dissection of the Mouse Eye for RNA, Protein, and Histological Analyses

IMMUNOHISTOCHEMISTRY This protocol describes basic immunohistochemistry methods of the retina, based on antibody detection or cell-selective lectin binding. Table 1 summarizes the commonly used markers of the different retinal cell types.

Materials Slides with cryosections (see Basic Protocol 5) Phosphate-buffered saline (PBS)

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Immunohistochemistry blocking solution (see recipe) Primary antibodies (commercially available or cited in literature) Secondary fluorescent antibodies (e.g., Molecular Probes, Invitrogen) Cityfluor AF3 mounting medium (Cityfluor) Nail polish Immunohistochemistry tray chamber with lid (e.g., StainTray Slide Staining System, IHC world) Coverslips Fluorescence microscope equipped with appropriate filters 1. Rehydrate the slides in 1× PBS. When using Coplin jars or similar glassware, 100 ml solution should be prepared for each step. Alternatively, prepare 50 ml solution when using 50-ml centrifuge tubes. When Table 1 Cell-Specific Markers of Mouse Retinal Cell Typesa

Antigens RPE

Retinol-binding protein receptor (Stra6) Retinal pigment epithelium 65 kD (Rpe65) Lecithin retinol acyltransferase (Lrat) Retinol dehydrogenase 5 (Rdh5)

Rod photoreceptors

Rhodopsin (Rho) Guanine nucleotide binding protein, alpha transducing 1 (Gnat1)

Cone photoreceptors

Guanine nucleotide binding protein, alpha transducing 2 (Gnat2)

Blue cone photoreceptors

S-opsin (opn1sw)

Green cone photoreceptors

M-opsin (opn1mw)

Rod bipolar cells

Metabotropic glutamate receptor 6 (mGluR6) Proteine kinase C, alpha subunit (PKCa)

Horizontal cells

Calbindin (Calb1)

Amacrine cells AII

Parvalbumin (Pvalb) Disabled 1 (Dab1)

Amacrine cells, dopaminergic

Tyrosine hydroxylase (Th)

Amacrine cells, cholinergic

Choline acetyltransferase (Chat)

Amacrine cells

Nitric oxide synthase (Nos)

Ganglion cells

Thymus cell antigen 1 (Thy1) Brn3a

ipGC

Melanopsin (opsin 4)

M¨uller glial cells

Glutamine synthetase (Glul) Vimentin

Lectins Photoreceptors

Wheat germ agglutinin

Cone photoreceptors

Peanut agglutinin

Blood vessels

Isolectin B4

a ipGC: intrinsically photosensitive ganglion cells. Rpe65 is also detected in cone photoreceptors. Protein kinase C alpha

subunit is predominantly expressed in rod bipolar cells. For a detailed description of immunological makers of the mouse retina, see also Haverkamp and W¨assle (2000).

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processing only very few slides, milliliter amounts of solution can be directly pipetted onto the slides.

2. Place the slides in a tray chamber. 3. Cover the slides with immunohistochemistry blocking solution for 1 hr at 21◦ C. 4. Dilute the primary antibody in blocking solution. Commercial antibodies are typically diluted 1:200.

5. Incubate the diluted primary antibody overnight at 4◦ C. Alternatively, incubate a minimum of 2 hr at 21◦ C.

6. Rinse the sections briefly twice with blocking solution. 7. Wash the sections with blocking solution for 5 min. 8. Dilute the secondary antibody in blocking solution. Secondary antibodies are typically diluted 1:1000.

9. Incubate the diluted secondary antibody for 1 hr at room temperature in the dark. 10. Rinse the sections briefly twice with 1× PBS. 11. Wash the sections three times, each time for 5 min in 1× PBS. 12. Cover the sections with a drop of Cityfluor AF3. 13. Cover the slide with coverslips and fix with nail polish. 14. Image the slides using a fluorescence microscope equipped with appropriate filters. SUPPORT PROTOCOL 2

MULTIPLE HISTOCHEMICAL STAININGS This protocol exemplifies a combination of fluorescent histochemical stainings described in Basic Protocols 7 and 8. First, a secondary antibody coupled to Alexa 594, i.e., emitting in the red, allows detection of a specific protein. Second, nuclei are stained with DAPI, i.e., emitting in the blue. Third, a fluorescein-coupled lectin, i.e., emitting in the green, allows detection of cone photoreceptors.

Materials Slides with cryosections (Basic Protocol 5) Phosphate-buffered saline (PBS), 1× Immunohistochemistry blocking solution (see recipe) Primary antibody Secondary antibody coupled to Alexa Fluor 594 (Invitrogen) DAPI staining solution (see recipe) Fluorescein-conjugated peanut agglutinin (FITC-PNA; Sigma) Cityfluor AF3 mounting medium (Cityfluor) Nail polish

Dissection of the Mouse Eye for RNA, Protein, and Histological Analyses

Immunohistochemistry tray chamber with lid (e.g., StainTray Slide Staining System, IHC world) Coplin jars or similar histology labware, optional 21◦ C incubator Coverslips Fluorescence microscope equipped with appropriate filters Perform immunohistochemistry 1. Rehydrate the slides in 1× PBS.

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2. Place the slides in a tray chamber. 3. Cover the slides with 1 ml immunohistochemistry blocking solution for 1 hr at 21◦ C. 4. Dilute the primary antibody in blocking solution. Commercial antibodies are typically diluted 1:200.

5. Incubate the sections with 0.2 ml of the diluted primary antibody overnight at 4◦ C. 6. Rinse the sections briefly twice with 0.2 ml blocking solution. 7. Wash the sections with 0.2 ml blocking solution for 5 min. 8. Dilute the secondary antibody coupled to Alexa Fluor 594 in blocking solution. Secondary antibodies are typically diluted 1:1000.

9. Incubate the sections with 0.2 ml of the diluted secondary antibody for 1 hr at 21◦ C in the dark. 10. Rinse the sections briefly twice with 0.2 ml 1× PBS. 11. Wash the sections three times, each time for 5 min with 0.2 ml 1× PBS.

Perform DAPI staining 12. Stain the slides with 0.2 ml of a 300 nM DAPI staining solution for 20 min at 21◦ C. For convenience, steps 12 to 15 can be performed in Coplin jars or similar glassware.

13. Wash the slides three times, each time for 5 min with 0.2 ml 1× PBS.

Perform lectin staining 14. Stain the slides with 0.2 ml of a 20 μg/ml fluorescein-conjugated peanut agglutinin/1× PBS solution for 75 min at 21◦ C in the dark. 15. Wash the slides three times, each time for 5 min with 0.2 ml 1× PBS.

Mount slides 16. Cover the sections with a drop of Cityfluor AF3. 17. Cover the slide with coverslips and fix with nail polish. 18. Image the slides using a fluorescence microscope equipped with appropriate filters.

REAGENTS AND SOLUTIONS Use autoclaved deionized water in all recipes and protocol steps.

DAPI staining solution Prepare the stock solution by dissolving DAPI (4 ,6-diamidino-2-phenylindole, dihydrochloride; Invitrogen) in deionized water to 100 μM Store aliquots protected from light up to several years at −20◦ C or for several months at 4◦ C Prepare the working dilution by diluting the stock solution 333 times to 300 nM in 1× PBS (see recipe) NOTE: Dilutions up to 1500 times (66 nM final DAPI concentration) still result in prominent nuclear staining.

Embedding medium (according to Yazulla) In a microwave (or waterbath), dissolve 3 g of gelatin in 20 ml deionized water Cool at 37◦ C continued

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Gently add 30 g bovine serum albumin (Fraction V; Sigma) in 50 ml of deionized water and let dissolve overnight Mix the two solutions Add deionized water up to 100 ml Stir gently to remove bubbles Store in aliquots up to 1 year at −20◦ C 70% ethanol/30% HCl 0.12 M Dilute 37% HCl (i.e., 12 M) 10 times, to 0.12 M Measure 70 ml of 70% Ethanol + 30% HCl (0.12 M) Add 30 ml of 0.12 M HCl Mix Store up to 1 month at 21◦ C 50% glycerol/50% 1× PBS Measure 50 ml glycerol, >99.5 % (Sigma) Add 50 ml of 1× PBS (see recipe) Mix thoroughly under agitation Store indefinitely at room temperature Immunohistochemistry blocking solution 1× PBS (see recipe) 2% goat serum 0.2% Triton X-100 Prepare blocking solution freshly Paraformaldehyde, 4%/1× PBS Weigh 4 g of paraformaldehyde Add 1× PBS (see recipe) up to 100 ml Add 10 drops of 1 M NaOH Dissolve under agitation at 60◦ C (about 10 min) Neutralize with 8 to 9 drops of 1 M HCl Adjust pH to 7-7.5 using 0.1 M HCl Dispense into 10-ml aliquots and store up to 1 year at −20◦ C CAUTION: Paraformaldehyde is toxic. Operate under a ventilated chemical hood wearing a protection mask.

Phosphate-buffered saline (PBS), 1× 154 mM NaCl 1 mM KH2 PO4 3 mM Na2 HPO4 heptahydrate Store up to 6 months at room temperature RIPA buffer

Dissection of the Mouse Eye for RNA, Protein, and Histological Analyses

50 mM Tris·Cl, pH 8.0 150 mM NaCl 1% NP-40 0.5% deoxycholate 0.1% SDS Complete protease inhibitors (Roche), freshly added Store stock solution without protease inhibitors up to 1 month at room temperature

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30% sucrose/1× PBS Weigh 30 g of sucrose Add 1× PBS (see recipe) up to 100 ml Dissolve under agitation at 37◦ C Store indefinitely at 4◦ C Tissue homogenization buffer 100 mM NaCl 50 mM Tris·Cl, pH 7.5 1 mM disodium EDTA 0.1% Triton X-100 10 mM NaF Complete protease inhibitors (Roche), freshly added Store stock solution without protease inhibitors up to 1 month at room temperature COMMENTARY Background Information Mouse genetics and visual system for the study of human eye diseases Mice are nocturnal animals and rely principally on olfactory, auditory, and tactile information. Because of their reduced visual function in general, and their low visual acuity in particular, mice were therefore not widely used in vision research. However, the power of mouse genetics based on the availability of inbred strains and, more recently, of genetically modified animals, stimulated vision research in this small rodent (Chalupa and Williams, 2008). Sequencing of the mouse and human genomes showed that ∼99% of mouse genes had human homologues and that the gene order was highly conserved between these two species (Taft et al., 2006). Furthermore, human diseases caused by mutations, including genetic eye diseases, are often recapitulated by similar mutations in mice.

Critical Parameters Ophthalmic phenotypes of common mouse laboratory strains The genetic background of a given mouse can greatly influence on its visual properties. Albino mice (e.g., A/J, AKR/J, BALB/cByJ, BALB/cJ) suffer from light-induced retinal degeneration in bright-light housing conditions. Anophthalmia and asymmetric microphthalmia are frequent (1% to 10%) in the commonly used inbred black C57BL strain. DBA/2J mice typically develop glaucoma after 9 months of age, due in part to iris stromal atrophy caused by a mutant allele of the tyrosinaserelated protein 1 gene (Tyrp1b ). Finally, a socalled rd1 (retinal degeneration 1) mutation

in the phosphodiesterase 6b gene (Pde6brd1 ) causes a rapidly progressing retinal degeneration and is especially frequent in mice with either C3H or FVB ancestry (e.g., C3H/HeJ, FVB/NJ, MOLF/EiJ, SJL/J). Housing conditions Because of the differential sensitivity of different mouse strains to light-induced retinal degeneration, it may be necessary to adapt the lighting conditions. For instance, different light intensities may be used in separate animal stock rooms.

Troubleshooting RNA preparation Lack of detectable mRNA isolated from dissected tissues may be due to RNA degradation, or failure to generate cDNA or detect cDNA by real-time PCR. Procedural controls can rule out cDNA amplification and RT-PCR as possible causes. RNA degradation may occur due to improper sample isolation and/or storage, or during the RNA isolation procedure. Protein preparation Protein degradation may occur if the samples are not processed on ice and the protease inhibitors are no longer active. For immunoblotting, protein loading and transfer can be determined through the use of the Ponceau stain, which reversibly binds to the proteins. The absence of red staining on the membrane indicates one of these first two steps failed. Immunohistochemistry Antibodies with nonspecific binding are the common cause of non-interpretable results.

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Anticipated Results RNA preparation From a single lens, iris, optic nerve, RPE and cornea, 20 to 30 μg of total RNA is at least recovered. From a retina, about 60 to 100 μg of total RNA is typically isolated. Protein preparation For all dissected eye tissues, from a single eye, 200 to 400 μg of proteins is routinely isolated.

Time Considerations Dissection Removing solely the retina from a mouse eye typically requires 5 min. To properly dissect all of the described eye tissues requires about 15 min per mouse eye. RNA preparation Standard RNA preparation procedures require ∼90 min. Homogenization of the cornea may require additional time. Protein preparation Standard protein preparation procedures require ∼5 min. Homogenization of the cornea may require additional time. Immunohistochemistry The entire protocol starting from dissection to observation of histological sections takes 4 working days.

Literature Cited Braissant, O. and Wahli, W. 1998. A simplified in situ hybridization protocol using nonradioactive labeled probes to detect abundant and rare mRNAs on tissue sections. Biochemica 1:10-16. Chalupa, L.M. and Williams, R.W. 2008. Eye, Retina, and Visual System of the Mouse. The MIT Press, Cambridge, Mass. Donovan, J. and Brown, P. 2006. Euthanasia. Curr. Protoc. Immunol. 73:1.8.1-1.8.4. Eldred, W.D., Zucker, C., Karten, H.J., and Yazulla, S. 1983. Comparison of fixation and penetration enhancement techniques for use in ultrastructural immunocytochemistry. J. Histochem. Cytochem. 31:285-292. Escher, P., Lacazette, E., Courtet, M., Blindenbacher, A., Landmann, L., Bezakova, G., Lloyd, K.C., Mueller, U., and Brenner, H.R. 2005. Synapses form in skeletal muscles lacking neuregulin receptors. Science 308:1920-1923. Haverkamp, S. and W¨assle, H. 2000. Immunocytochemical analysis of the mouse retina. J. Comp. Neurol. 424:1-23. Smith, R.S., John, S.W.M., Nishina, P.M., and Sundberg, J.P. 2002. Systematic evaluation of the mouse eye: Anatomy, pathology, and biomethods. CRC Press, Boca Raton, Fla. Sugita, S. and Streilein, J.W. 2003. Iris pigment epithelium expressing CD86 (B7-2) directly suppresses T cell activation in vitro via binding to cytotoxic T lymphocyte-associated antigen 4. J. Exp. Med. 198:161-171. Taft, R.A., Davisson, M., and Wiles, M.V. 2006. Know thy mouse. Trends Genet. 22:649-653.

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Myofiber Damage Evaluation by Evans Blue Dye Injection Christine I. Wooddell,1 Hannah G. Radley-Crabb,2 Jacob B. Griffin,1 and Guofeng Zhang2 1

Roche Madison Inc., Madison, Wisconsin School of Anatomy and Human Biology, The University of Western Australia, Crawley, Australia 2

ABSTRACT Evans blue dye (EBD) can be used in live mice to study muscle pathology or injury, including exercise-induced muscle damage. EBD is excluded from intact cell membranes but leaks into cells, including muscle fibers, when the cell membrane is ruptured. EBD can be visualized by its autofluorescence under a fluorescence microscope. EBD-stained myofibers can be quantified from microscope images of muscle cross-sections. These myofibers are often in clusters that lend themselves to morphometric analysis. When the damaged myofibers are interspersed among intact myofibers, however, a more suitable approach is to count individual myofibers in the field of view. A much faster approach to measure EBD in muscles from different strains of mice or between treatment groups is to extract the EBD from muscle samples and quantitate it using a spectrophotometric microplate reader. The advantages and disadvantages of using each of these approaches C 2011 by John Wiley & Sons, are discussed here. Curr. Protoc. Mouse Biol. 1:463-488  Inc. Keywords: Evans blue dye r skeletal muscle r mouse model r mdx mouse

INTRODUCTION Evans blue dye (EBD) is a cell membrane–impermeable tetrasodium diazo salt (mol. wt. 960.82 g/mol) that has been used as a tracer to study the vasculature of living animals and to evaluate the integrity of cell membranes in vivo (Reeve, 1957). EBD is usually delivered into the vasculature of mice by injecting it into the tail vein or into the peritoneum (Hamer et al., 2002). The injected EBD binds to albumin in the blood and the EBD-albumin conjugate circulates through the vasculature. The EBD-albumin conjugate is excluded from cells by membranes that are intact, but it leaks into and accumulates in myofibers that are damaged by rupture of the plasma membrane (Straub et al., 1997). Thus, EBD can be used for phenotypic characterization of mouse strains that have muscle pathologies such as muscular dystrophy (Straub et al., 1997). Leaky myofibers may also be the result of muscle injury in otherwise healthy animals. EBD is, therefore, useful for studying muscle pathology, muscle injury, and the effects of exercise on myofibers. The protocols describe procedures for measuring skeletal muscle marked by EBD caused by permeable (damaged) myofiber membranes. Basic Protocol 1 describes two options for delivering EBD into the mouse. Basic Protocols 2 through 4 describe separate approaches for quantitation of EBD staining. More specifically, Basic Protocols 2 and 3 present two different procedures based upon morphometrical analysis for collecting and freezing muscle samples suitable for sectioning. One procedure involves the stabilization and orientation of muscle samples in tragacanth gum, followed by freezing in a slurry of isopentane in liquid nitrogen (see Basic Protocol 2). The other procedure involves muscle stabilization and orientation in embedding molds with O.C.T. (see Basic Protocol 3 and Support Protocols 1 and 2). Both procedures ultimately arrive at the same end point (a muscle sample that can be cut on the cryostat to produce muscle sections) and could be Current Protocols in Mouse Biology 1: 463-488, December 2011 Published online December 2011 in Wiley Online Library (wileyonlinelibrary.com). DOI: 10.1002/9780470942390.mo110141 C 2011 John Wiley & Sons, Inc. Copyright 

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interchanged, depending on laboratory facilities. However, for consistency, all muscle samples from an experiment should be prepared the same way. Finally, Basic Protocol 4 details a rapid alternative quantitative method for EBD measurement, which is suitable for analysis when total uptake of EBD is expected to be significant between treatment groups. In the Commentary section, some of the considerations that will help researchers select the most suitable approach for their intended application are discussed. The protocols presented here focus on evaluation of skeletal muscles of the mouse hind leg as a model system. The hind legs contain the largest muscle mass in the mouse. Another advantage of evaluating the leg muscles is that these can be challenged by exercise protocols. Most of the methods used to evaluate the leg muscles could likely be applied to other muscles. BASIC PROTOCOL 1

DELIVERY OF EBD There are two commonly used approaches to delivering EBD into the mouse vasculature. The EBD solution can be injected intravenously (i.v.) into either the lateral vein of the tail or it can be injected intraperitoneally (i.p.). The advantage of i.v. injection is that the EBD solution is immediately available through the vasculature. Intravenous injection of EBD is the more technically challenging delivery technique, but carries the advantage of immediate availability through the vasculature, which may be desirable for timecritical studies. Intraperitoneal EBD injection, while less demanding of user skill and precision, necessitates a several-hour waiting period to allow for appropriate systemic dye dispersion, before animals can be evaluated. Approximately twice as much EBD is used for the i.p. injection compared to i.v. injection. The mouse should be injected in the lower portion of the abdomen, to avoid damaging the liver, and also away from the midline to avoid the bladder.

Materials Mice Evans blue dye (Sigma-Aldrich, cat. no. E2129) prepared at 5 mg/ml with sterile physiological saline (0.90% NaCl) (see recipe) 1% to 2% isoflurane Phosphate buffered saline (PBS) Heat lamp or 50-ml conical tube containing warm water (50◦ C) 1-ml syringe with 30-G needle for i.v. injection or 500-μl or 1-ml syringe with 27-G needle for i.p. injection Anesthesia machine with animal chamber or an animal holder to restrain the mouse during injection Injection of EBD solution into the tail vein (i.v.) Injecting into the tail vein requires some skill because the needle must puncture the vein in only one spot and remain in the lumen of the vein during the injection. The tail vein is more easily visualized in white mice than in darkly pigmented mice, which may be useful for training purposes. 1a. Weigh each mouse and record its weight along with identifying information, e.g., ear tag number. Calculate how much EBD solution to inject into each animal (50 μl of 5 mg/ml EBD solution per 10 g body weight). Myofiber Damage Evaluation by Evans Blue Dye Injection

2a. At least 30 min prior to injection, place the cage containing the mouse to be injected under a warming lamp to dilate its blood vessels. If a heat lamp is not available, place the tail into a tube of warm water (∼50◦ C) for 1 to 2 min to dilate the blood vessels.

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3a. Using a 1-ml syringe with a 30-G needle, draw the 5 mg/ml EBD solution into the syringe. Remove all air bubbles from the syringe by flicking the sides of the syringe. 4a. Anesthetize mouse with 1% to 2% isoflurane. If an anesthesia machine is not available, a mouse restraining structure (e.g., tube) can be prepared to facilitate the injection. This is made from a 50-ml conical polypropylene centrifuge tube. Cut a breathing hole with a diameter of 0.5 to 1 cm in the bottom of the conical tube. Make a 3-cm long and 0.3-cm wide notch on the upper edge of the tube, perpendicular to the rim. Place the mouse inside the tube with its tail sticking out of the notch. Continue to hold the tail while performing the injection.

5a. Lay the mouse on its side and hold the distal part of the tail with thumb and fourth finger. Position the tail between the index and middle fingers. 6a. Using index and middle fingers, put some pressure on the proximal part of tail to block blood flow in the tail, which causes the tail to brim with blood and allows for easier injection. 7a. Inject the calculated quantity of EBD solution slowly into the tail vein. 8a. Place the mouse back into its cage.

Injection of EBD solution into the peritoneum (i.p.) 1b. Weigh each mouse and record its weight along with identifying information, e.g., ear tag number. Calculate how much EBD solution to inject into each animal (100 μl of 5 mg/ml EBD solution per 10 g body weight). 2b. Using a 500-μl or 1-ml syringe with a 27-G needle, draw the EBD solution into the syringe. Remove all air bubbles by flicking the side of the syringe. 3b. Anesthetize mouse as in step 4a. 4b. Holding the abdominal skin away from its body using fingers or forceps, slowly insert the needle into the lower area of the peritoneal cavity from the side and in the direction of the midline, being careful not to damage any organs by inserting the needle too far. 5b. Slowly inject the calculated amount of EBD solution. 6b. Place mouse back into its cage.

Alternative approach for i.p. injection of EBD solution without anesthesia 1c. Freshly prepare solution of 1% EBD in PBS. 2c. Weigh each mouse and record its weight along with identifying information, e.g., ear tag number. Calculate how much EBD solution to inject into each animal (100 μl sterile EBD solution per 10 g body weight). 3c. Using a 500-μl syringe with a 27-G needle, draw the sterile EBD solution into the syringe. Remove all air bubbles from the syringe by flicking the sides of the syringe. 4c. Pick up and immobilize the mouse by pinching across the shoulder blades (use thumb and forefinger). Expose the abdomen by holding the tail out of the way (use little finger) and insert the needle slowly into the peritoneal cavity from the side, aiming toward the vicinity of the nipples. If right handed, use left hand to hold mouse and right to hold syringe. If you inadvertently insert the needle into the bladder during i.p. injection of the EBD, blue dye will be evident in the urine within ∼5 min.

5c. Slowly inject the calculated amount of EBD solution.

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6c. Return mouse to its cage. This protocol is appropriate for adult mice when the researcher desires to limit the exposure of mice to anesthesia. Use of gaseous anesthesia is preferable for performing i.p. injections on young mice (<4 weeks old) as it reduces the risk of injury in very small mice. BASIC PROTOCOL 2

MORPHOMETRIC QUANTITATION OF AREAS WITHIN THE MUSCLE The presence of EBD in skeletal muscle can be formally quantitated in transverse muscle sections. EBD-positive myofibers are visualized using fluorescence microscopy and the percentages quantitated by morphometric analysis (Fig. 1) as per Piers et al. (2011) and Shavlakadze et al. (2004). This procedure utilizes frozen histology methods and thus allows for a relatively quick evaluation by avoiding paraffin processing. Multiple sections can be cut from the same frozen muscle sample, allowing serial sections to be stained with hematoxylin and eosin (H&E) stain to assess muscle morphology or by immunohistochemistry to identify other proteins of interest. One limitation of this technique, as with all frozen histology, is the minor risk of ending up with a freeze fracture in the muscle samples.

Materials Tragacanth gum (Sigma Aldrich, cat. no. G1128) Isopentane Liquid nitrogen Isoflurane Acetone Xylene DPX mountant glue (BDH, cat. no. 36029.4H) Cork (∼1 cm3 per sample) Cryostat Glass histology slides (VWR Superfrost Plus Micro slides, cat. no. 48311-703) Glass cover slips Fluorescence microscope (band pass: excitation 515 to 560 nm; low pass: emission 590 nm) with camera and image capture software (e.g., Leica DM RBE microscope, a personal computer, a Hitachi HVC2OM digital camera, Image Pro Plus 6.2 software, and Vexta stage movement software)

Myofiber Damage Evaluation by Evans Blue Dye Injection

Figure 1 Morphometric quantitation of EBD-stained myofibers in the tibialis anterior muscle from dystrophic mice (mdx, left; and mdx/IGF.1, right) given 1% EBD in PBS (pH 7.5) by i.p. injection 24 hr prior to sacrifice. Adapted from Shavlakadze et al. (2004). Reprinted with permission from Macmillan Publishers.

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Prepare reagents and equipment 1. Mix the tragacanth gum solution 1 day before sampling and refrigerate overnight (prepare as per manufacturer’s instructions). Ideally, the tragacanth gum should be the consistency of toothpaste.

2. Cut cork into ∼1-cm3 squares prior to harvesting muscle. 3. Just before harvesting muscles, prepare a slurry of isopentane cooled in liquid nitrogen. Place the small container of isopentane into a larger container of liquid nitrogen, allowing the isopentane to be cooled without mixing with the liquid nitrogen.

Harvest muscle 4. Sacrifice mice as per institutionally approved animal ethics committee protocols (e.g., anesthetize with isoflurane followed by cervical dislocation). 5. Peel away and remove the skin from the limbs to expose the muscles underneath. 6. Dissect out the muscles of interest (e.g., tibialis anterior or quadriceps). This can be done by removing any covering connective tissue, cutting the appropriate tendons, and lifting out the whole muscle, or by sliding a surgical blade under the belly of the muscle and slicing it away from the bone.

7. Using a surgical scalpel blade, bisect the fresh muscle transversely (in the center and perpendicular to the grain). Cover the top of the cork square with a small layer (8-mm) of tragacanth gum. Mount the pieces of muscle in the tragacanth gum. The tragacanth gum stabilizes the muscle and maintains myofiber orientation. A 1-cm3 piece of cork is large enough to fit both halves of the tibialis anterior muscle side by side, whereas the quadriceps muscle will require two pieces of cork.

8. Freeze the muscles in a slurry of isopentane cooled in liquid nitrogen by placing the muscles into the isopentane slurry for ∼20 sec or until the tragacanth gum turns white. Isopentane reduces surface tension, thus producing a frozen sample that is excellent for histological evaluation with little or no freezing artifacts.

Process muscle 9. Using a cryostat, cut frozen muscle sections ∼8-μm thick directly onto uncoated or silinated glass histology slides. Slides can be stored for ∼2 weeks at −20◦ C until ready to visualize.

10. Fix muscle sections 1 min using cold acetone (–20◦ C) and then air dry for 2 to 3 min at ambient temperature (25◦ C). 11. Dip slides in three changes of xylene, 3 min each time. 12. Cover muscle sections with a small amount of DPX mountant glue (∼0.2 ml, depending on size of cover slip) and then mount a glass cover slip over the glue; avoid creating air bubbles.

Collect images 13. View unstained frozen sections by fluorescence microscopy. EBD autofluoresces red under green light (band pass: excitation 515 to 560 nm; low pass: emission 590 nm).

14. Non-overlapping images of a transverse muscle section can be tiled/stitched together to provide a single digital image of the entire muscle cross-sectional area. To do this, acquire images using a fluorescence microscope, digital camera, and image capture

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software (e.g., Leica DM RBE microscope, a personal computer, a Hitachi HVC2OM digital camera, Image Pro Plus 6.2 software, and Vexta stage movement software). More sophisticated digital slide scanners (e.g., Aperio Scanscope) can be used to collect images, although fewer researchers are likely to have access to such machines. Morphological analysis can be performed on multiple single frame images taken from the same muscle section (multiple fields of view) rather than analyzing the entire crosssectional area. However, this method assumes that EBD+ myofibers are homogeneous throughout the muscle cross-section, which may not always be true. In the absence of software that allows stitching together images, sequential images can be taken across the diameter of the muscle and evaluated separately.

Analyze images and quantitate EBD 15. Identify the EBD+ myofibers manually and then quantify them using image analysis software (e.g., Image Pro Plus). To quantitate the abundance of EBD+ myofibers per muscle cross-sectional area, first open the digital image in the image analysis software (e.g., Image Pro Plus) and measure (draw around) the entire cross-section area of the muscle. This can be done in pixels (sufficient when determining a percentage) or the software can be calibrated and features measured as an absolute value (e.g., μm2 ). Second, identify and manually measure (draw around) all the EBD+ myofibers. The data can be exported directly into Microsoft Excel and the percentage of EBD+ myofibers calculated as: (total area of EBD+ myofibers/total area of muscle cross-section) × 100 = % EBD+ Then perform appropriate statistical analyses. BASIC PROTOCOL 3

DETERMINING PERCENTAGES OF EBD-POSITIVE MYOFIBERS To determine as accurately as possible the percentages of EBD+ myofibers in the mouse leg, an investigator will need to select a sufficient number of muscle sections to evaluate. These should be representative of the whole. Many of the individual muscles in the mouse leg are small. Rather than attempting to evaluate each individual muscle, the mouse leg can be divided into five large groups of muscles: muscles in the anterior, posterior, and

ventral and medial muscles

lateral superficial muscles Group 1

Group 1

Group 3

Group 3

Group 2

Group 5

Group 5 Group 4

Group 4 Group 5

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Group 1 - anterior thigh Group 2 - posterior thigh Group 3 - medial thigh Group 4 - posterior lower Group 5 - anterior lower

Figure 2 Dissection of the mouse leg into groups of muscles. Muscle groups 1 to 5 are defined by their anatomical location as shown in this color-coded schematic.

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medial regions of the upper leg and the anterior and posterior regions of the lower leg (Fig. 2). Multiple cross-sections (specimens) are then cut from each group of muscles and microscope images captured along the entire diameter of the specimen. Dividing the leg into five muscle groups allows sufficient sample to be evaluated by multiple methods. For example, it may be preferred to evaluate the leg muscles by counting how many muscle fibers are positive for EBD and how many muscle fibers are positive for a protein of interest as determined by immunohistochemistry (IHC) or to determine the concentration of a protein of interest by immunoblotting, e.g., dystrophin (Fig. 3). Preparation of multiple cross-sections is described in this protocol, while determining the percentage of EBD-positive fibers within these sections is detailed in Support Protocol 1. Due to the difficulty of visualizing individual fibers that are not stained by EBD, immunohistochemistry is used to outline fibers. For this purpose, staining for dystrophin is implemented, and the method for generating anti-dystrophin antibodies is described in Support Protocol 2. While success has been achieved using anti-dystophin antibodies generated by this method, it is expected that polyclonal anti-dystrophin antibodies obtained by other methods and possibly mouse monoclonal antibodies to mouse dystrophin would serve equally well.

Figure 3 Overlay of EBD staining and dystrophin immunohistochemistry. (A, B) The mdx mouse was given six hydrodynamic limb vein injections with dystrophin-expressing pDNA into the hind legs. When it was 18 months old, it was exercised on a treadmill at 12 m/min for 10 min, injected with EBD the next day, and sacrificed for evaluation 24 hr after this. Consecutive muscle sections were stained for dystrophin (FITC), left panels, or stained with hematoxylin and eosin for evaluation of the histology, right panels. The FITC images were overlapped with images of EBD auto-fluorescence (in the CY3 channel) from the same muscle sections. The FITC-labeled anti-mouse IgG secondary antibody labeled dystrophin expressing myofibers (arrows). This secondary antibody also labeled the endogenous IgG antibodies that accumulated within damaged myofibers (*) in A. Myofibers staining intensely with EBD (*) and those staining only lightly (L) are shown in the merged images and corresponding histology sections from the medial thigh muscles of the same mouse. This figure was adapted from Zhang et al. (2010).

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Materials Tissue-Tek O.C.T. compound (Sakura Finetek, cat. no. 4583) Mice 3% to 5% isoflurane Liquid nitrogen Styrofoam container for holding liquid nitrogen and Styrofoam float for holding embedding molds while freezing in liquid nitrogen (Fig. 4) Peel-A-Way disposable embedding molds, truncated (8 × 8 × 20–mm; PolySciences, cat. no. 18985) 1. Produce a Styrofoam float rack by cutting off the top of a Styrofoam rack that comes with 50-ml conical polypropylene tubes (Fig. 4), making the Styrofoam float between 2.0- and 2.5-cm thick to ensure that the embedding molds come into contact with the liquid nitrogen and the muscle freezes quickly. 2. Prepare embedding molds before harvesting muscles as follows. a. Label an embedding mold for each muscle sample that will be harvested (one group of muscles), including animal number, muscle group number, and left (L) or right (R) leg. b. Right before embedding, fill the embedding molds 5- to 7-mm deep with O.C.T. freezing compound. Blocks of O.C.T. containing specimens frozen in embedding molds that are not truncated at the end (such as those shown in Fig. 4) can be truncated by hand to resemble blocks frozen in the truncated molds.

3. Anesthetize the mouse with 3% to 5% isoflurane, then euthanize by cervical dislocation (or per approved institutional animal ethics protocols). 4. Remove the skin from the hind legs to expose the muscles. 5. Divide the muscles into five groups as indicated by the color coding in Figure 2:

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Figure 4 Preparation for embedding muscle in freezing medium. Cut out Styrofoam float racks from the Styrofoam tube-holding racks that come with 50-ml conical polypropylene tubes. Draw a line on the Styrofoam rack as shown so that the float rack will be 2.0 to 2.5 cm in height. Embedding molds fit into the holes in the float rack, allowing liquid nitrogen to contact the embedding mold containing O.C.T. freezing compound when the float rack is placed into a Styrofoam box containing liquid nitrogen. Photograph provided by Julia Hegge and Tracie Milarch (Roche Madison). Current Protocols in Mouse Biology

a. Muscle group no. 1: the anterior group of muscles of the upper leg. This group includes the quadriceps. It comprises ∼20% of the leg muscle mass. b. Muscle group no. 2: the posterior group of muscles of the upper leg. This group includes the biceps femoris and comprises ∼25% of leg. c. Muscle group no. 3: the medial group of muscles of the upper leg. This group includes the gracilis and adductor muscles. It comprises ∼25% of the leg muscle mass. d. Muscle group no. 4: the posterior group of muscles of the lower leg. This group includes the gastrocnemius and soleus. It comprises ∼20% of the leg muscle mass. e. Muscle group no. 5: the anterior group of muscles of the lower leg that is comprised primarily of the tibialis anterior and extensor digitorum longus. This group includes ∼10% of the leg muscle mass. 6. Freeze the muscle immediately after harvesting in Tissue-Tek O.C.T. compound. The integrity of muscle samples is preserved by rapidly freezing them in tissue freezing medium. The muscle pieces must be oriented in a consistent manner to obtain high-quality cross-sections. This can be achieved by freezing each muscle sample immediately after adding it to the freezing medium in an embedding cup. a. Embed each muscle with the grain of the muscle in a vertical position within the embedding cup, taking care not to trap air bubbles around the muscle. b. Add O.C.T. to cover the muscle sample by 3 to 5 mm. c. Fill the Styrofoam container with enough liquid nitrogen that the Styrofoam float rack will float. d. Place each embedding mold into the Styrofoam float rack, and then set the float rack into the liquid nitrogen. Take care to keep the muscle piece in a vertical orientation while it is freezing to allow for high-quality cross-sectioning. e. Wait until the O.C.T. turns hard and solid white. Then, move the embedding molds to a container of dry ice. Place muscle samples from each animal into a separate plastic storage bag. Store the frozen O.C.T. blocks preserved up to 1 year at −80◦ C. 7. Section the frozen tissue when ready to proceed with immunostaining (see Support Protocol 1).

COUNTING PERCENTAGE OF EBD+ MYOFIBERS EBD+

Determining the percentage of myofibers requires counting the total number of myofibers in the muscle cross-sections. The EBD autofluoresces, so EBD+ myofibers can easily be counted when they are interspersed among the non-stained myofibers. They are more difficult to distinguish when groups of them are clustered together as can be seen in Figure 5. Individual myofibers within the specimen that are not EBD-stained can also be difficult to distinguish unless a method to outline individual myofibers is used, e.g., an immunohistochemistry staining method utilizing antibodies to label the dystrophin that is associated with the cell membrane, which results in a crisp outline of each myofiber.

SUPPORT PROTOCOL 1

In mice such as mdx that do not produce dystrophin, anti-mouse IgG antibodies can be used instead to outline the myofiber as shown in Figure 6B. Antibodies to other blood proteins serve the same function (see Fig. 6C,D), although the myofiber membrane is generally not outlined as distinctly when targeting blood proteins as it is when labeling the dystrophin in mice that have the wild-type dystrophin protein.

Materials Frozen tissue (see Basic Protocol 3) 2% to 4% formalin in PBS

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PBS Mouse anti-dystrophin polyclonal antibodies diluted as needed in PBS (see Support Protocol 2) FITC-conjugated goat anti-mouse IgG (FAB-specific; Sigma-Aldrich, cat. no. F8771) diluted 1:400 in PBS Cryostat Glass histology slides (VWR Superfrost Plus Micro slides, cat. no. 48311-703) Paraffin pen Microscope, excitation filters (FITC and CY3), camera Prepare sections and fix 1. Using a cryostat, begin cutting 8- to 10-μm cross-sections from the frozen tissue. Section approximately one-third off the end of each large muscle piece frozen in O.C.T. and less off small muscle pieces prior to collecting specimens. 2. Cut two adjacent sections and place these sections onto a slide. Discard the subsequent ten sections. Cut two more adjacent sections and place on same slide. Repeat this procedure until four to six sections from one muscle sample (a group of muscles) is placed onto the slide. Repeat this process for each muscle sample. 3. Fix tissue slices in 2% to 4% formalin in PBS for 5 to 10 min. 4. Rinse sections three times with PBS for 2 min each time.

Immunostain samples 5. Circle each specimen on the slide with a paraffin pen. 6. Pipet enough polyclonal anti-dystrophin antibodies in PBS onto the specimen to cover the sample (the paraffin circle made around the sample contains the fluid). The proper dilution of the mouse anti-dystrophin polyclonal antibodies in PBS is the one that provides clear staining without excessive background.

7. Incubate 40 to 60 min at 25◦ C.

Myofiber Damage Evaluation by Evans Blue Dye Injection

Figure 5 Dark and light EBD staining compared to histological staining. Myofibers containing dark (intense) or light EBD fluorescence were compared by histological H&E staining on sections of hind leg muscle from a 2-month-old mdx mouse that had been exercised on a treadmill. Consecutive frozen muscle sections from the gastrocnemius were evaluated for EBD fluorescence (A), or stained with H&E for histological evaluation (B). A few of the dark (D) and light (L) EBD-stained myofibers are indicated. Scale bar indicates 100 μm. This figure is from Wooddell et al. (2010).

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Figure 6 Co-localization of EBD, IgG, IgM, and C3 complement in damaged myofibers. Staining was performed on consecutive frozen sections from a 2.4-month-old mdx mouse that had been exercised on a treadmill, injected with EBD, and sacrificed for evaluation 24 hr later. (A) EBD fluorescence showing both lightly (L) and darkly (D) stained myofibers. Dark EBD staining appears as brighter fluorescence (more white) than the light EBD staining. (B) IgG immunostaining, fluorescein isothiocyanate (FITC)-labeled. (C) C3 complement immunostaining (FITC). (D) IgM immunostaining (FITC). The same myofibers are marked in consecutive sections. This figure is from Wooddell et al. (2010).

8. Rinse the sections three times with PBS for 2 min each time. 9. Incubate the sections in FITC-conjugated goat anti-mouse IgG in PBS for 40 min at 25◦ C. 10. Rinse sections three times with PBS, 2 min each time. 11. Keep sections in the dark until ready to image. Typically, images are collected from the specimen covered with some PBS to keep specimen moist, but without adding a cover slip. If images will not be collected immediately, then place a cover slip over the specimen to prevent the sample from drying, which would result in high background, and store for up to 2 days at 4◦ C.

Count myofibers 12. Take a series of images from one representative specimen from each slide that corresponds to one muscle group from one leg. At each microscope position take two images by using two different excitation filters, one for FITC (appears green) and one for CY3 (appears red). When looking at the specimen on the slide under the microscope, begin taking images at the edge farthest from you (or closest to the top of the image). Take the next image right below the first. Continue taking sequential images across the diameter of the muscle specimen until the bottom of the specimen is reached.

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13. Merge the two images taken at each microscope position, those taken with the FITC and CY3 filters. 14. Print out the merged images from each microscope position. Count the total number of myofibers in each image. 15. Count EBD+ myofibers on the computer screen. The bright yellow-green myofibers are those that stain intensely with EBD and contain IgG that leaked in through the damaged cell membrane. The myofibers that are only lightly EBD+ appear to be red. See Figure 3 as an example. Decide whether to count all EBD+ myofibers together, only the intensely EBD+ myofibers, or to separately count the intensely and lightly stained myofibers. 16. Combine in data tables for each muscle group the total number of myofibers, the total number of intensely stained EBD+ myofibers, and the total number of lightly EBD+ myofibers (if desired). For each muscle group, the percentage of EBD+ myofibers is determined by dividing the total EBD+ myofibers by the total number of myofibers counted.

Analyze + 17. Determine the weighted  percentages of EBD myofibers in the entire hind limb using the formula: ( A1 M1 + A2 M2 + A3 M3 + A4 M4 + A5 M5 )/Mt , where Ai is the average number of EBD+ myofibers counted in muscle group i and Mi is the weight of that muscle group, and Mt is the total weight of all groups of muscles in the leg. These averages can be used to compare groups of mice or different treatments. Alternative approaches The amount of labor involved in comparing the number of EBD+ myofibers between groups of animals and treatments can be reduced by the following approach. Instead of counting the total myofibers in each microscope view, count only the EBD+ myofibers

B 25 80 20

EBD+ myofibers/view in muscle groups

EBD+ myofibers/view in limb

A

15 10 5 0 unexercised exercised limbs limbs Treatment of old mdx mice

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70 60 50 40 30 20 10 0 Young ( E) Young ( E)

Old ( E)

Old ( E)

Figure 7 Counting EBD+ myofibers. (A) Total EBD+ myofibers (dark and light) were counted in ten to eleven microscope views across the diameter of muscle specimens from each of the 5 groups of muscles from the hind limbs of 13- to 19-month-old mdx mice that were either unexercised or had been exercised by running on a treadmill or a rotarod. A weighted average of EBD+ myofibers/view in the entire mouse limb was determined by taking into account the weight of each muscle group. (B) The total numbers of EBD+ myofibers in each of the 5 groups of muscles of the hind limbs of young (4- to 6-month-old) and 13- to 19-month-old mdx mice are evaluated as collectives of muscle samples. Young and old mice were unexercised (E–) or exercised (E+), p = 0.434 for young adult and p < 0.0001 for old animals. Each point represents the average number of EBD+ myofibers per microscope view for one group of muscles (one sample). Figure adapted from Wooddell et al. (2010).

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Figure 8 Use of EBD for a gene therapy experiment. (A, B) In a gene therapy study, mdx mice were given 6 hydrodynamic limb vein injections with a dystrophin-expressing plasmid (Dys) in one leg and a control plasmid in the other leg. At the end of the study, the mice were exercised on a treadmill for 15 to 30 min at 12 m/min and then evaluated for EBD+ myofibers in each muscle group of the legs. (A) The percentages of EBD+ myofibers in the control leg muscles were arbitrarily set at 100% to determine the relative percentages of EBD+ myofibers in the contra-lateral Dysinjected leg muscles. The average normalized EBD levels are shown for each group of muscles. (B) Each data point represents the weighted average percentage of EBD+ myofibers in one leg. Statistical analysis comparing contra-lateral pairs of muscles was performed with a two-tailed, Wilcoxon signed rank test (**p < 0.01). These data are from Zhang et al. (2010).

per microscope view. The weighted average of EBD+ myofibers in the whole limb can be determined using the same formula as in step 17, except in this case Ai refers to the average number of EBD+ myofibers per microscope view in muscle group i (Fig. 7A). When the variability in EBD staining is high within the animals that are being compared, as is the case with mdx mice, the power of analysis can be improved by comparing samples as collective groups of muscles. See Figure 7B; in this case, the weight of the samples is not taken into account. All samples contribute equally to the resulting average. If one particular group of muscles skews the results, perform the analysis with and without that group of muscles. For example, the tibialis anterior of muscle group no. 5 may be much more affected by an exercise protocol than the rest of the leg, although all of muscle group no. 5 comprises only 10% of the whole leg. When one leg of the mouse can be used as a control for a treatment given to the other leg, then pairs of muscle groups from the left and right legs can be compared using a two-tailed, Wilcoxon signed rank test for statistical analysis using GraphPad Prism software (e.g., Fig. 8).

PRODUCTION OF MOUSE POLYCLONAL ANTIBODIES TO DYSTROPHIN Naked plasmid DNA expressing a protein of interest can be delivered to mouse muscle to generate polyclonal antibodies against the expressed gene product, as described in Bates et al. (2006). Hydrodynamic limb vein (HLV) injection delivers the pDNA to skeletal muscles (Hagstrom et al., 2004). This procedure is referred to as genetic immunization. The reader is referred to Bates et al. (2006) for details regarding this method of antibody generation. Here, specific details for generating anti-dystrophin antibodies by HLV injection of dystrophin-expressing pDNA into the saphenous vein of an mdx mouse are described. The dystrophin protein is large (∼500 kD) and has conserved epitopes between mouse and human dystrophin. Therefore, pDNA expressing either human or mouse Current Protocols in Mouse Biology

SUPPORT PROTOCOL 2

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dystrophin can be used to generate antibodies in a dystrophin-deficient mouse that will cross-react with mouse dystrophin. Anti-dystrophin antibodies can also be produced in rats using the same procedure (Bates et al., 2006). Anti-dystrophin antibodies are usually produced after two HLV injections of the dystrophin pDNA. Sometimes a single injection is sufficient. Antibody production is confirmed using either an enzyme-linked immunosorbent assay (ELISA) or by immunohistochemistry (IHC), as described in Bates et al. (2006). The IHC test is described below.

Materials pDNA that produces human or mouse dystrophin (prepare or order endotoxin-free plasmid DNA) Sterile, physiological saline solution C57Bl/10ScSn-Dmdmdx /J (mdx-10ScSn) mice (Jackson Laboratories) 1% to 2% isoflurane anesthesia Ketoprofen Serum tubes with gel and clot activator 50-ml polypropylene tubes Heating pad at 37◦ C Scalpel Latex tourniquet 4-0 absorbable sutures 1-ml syringes Additional reagents and equipment for muscle harvesting, processing, and immunostaining (see Basic Protocol 3 and Support Protocol 1) and HLV injection (Hagstrom et al., 2004) Immunize mice 1. In a 50-ml tube, prepare 50 to 100 μg of dystrophin expression pDNA in a physiological saline solution at 25◦ C such that the final volume is 1 ml per injection. The commonly used cytomegalovirus (CMV) promoter works well for driving expression of this protein from pDNA in muscle (Bates et al., 2006; Zhang et al., 2010).

2. Anesthetize the mouse with 1% to 2% isoflurane and then place mouse on a 37◦ C heating pad for surgery. 3. Using a scalpel, make a small incision near the ankle of the C57Bl/10ScSn-Dmdmdx /J (mdx-10ScSn) mouse to visualize the saphenous vein. Other strains of mdx mice can be used as well, but these are less readily available.

4. Using a latex tourniquet around the upper part of the mouse hind limb to block blood flow as described (Bates et al., 2006), perform an HLV injection with 1 ml of pDNA/saline solution into the saphenous vein of the mdx mouse hind limb at a rate of 8 ml/min. For the HLV injection, refer to Hagstrom et al. (2004) for a detailed description. 5. Close the incision with a 4-0 suture. Give the mouse a subcutaneous injection of ketoprofen (5.0 mg/kg) as analgesia while it is still anesthetized. Keep the mouse on the heating pad until it awakens. 6. Repeat the HLV injection 2 weeks later. Myofiber Damage Evaluation by Evans Blue Dye Injection

7. Ten days after the second pDNA injection, collect blood from the mouse by retroorbital bleed. Isolate serum using serum tubes with gel and clot activator according to the manufacturer’s instructions.

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Immunohistochemistry test for antibody production 8. Use a wild-type mouse for IHC testing of the antibody. Sacrifice the mouse, harvest hind leg muscle, freeze muscle in O.C.T., and cut 8-μm thick cross-sections of the muscle as described for muscle sample preparation for myofiber counting. 9. As a negative control, use wild-type mouse serum. This control will demonstrate the background labeling from the secondary antibody.

10. Prepare the following dilutions of serum from the pDNA-injected mouse in PBS: 1:20, 1:40, 1:100, 1:200, 1:400, and 1:800. 11. Follow the steps described for dystrophin immunostaining in Support Protocol 1, including a 1:400 dilution of FITC-goat anti-mouse IgG. 12. Use the dilution that shows clear outlines around each myofiber with a minimum of background fluorescence for immunostaining. If the antibodies do not give adequate immunostaining when diluted at 1:100, then give the mouse an additional HLV injection of the dystrophin pDNA.

13. When the antibody titer is adequate for use at a 1:100 or greater dilution, then collect additional serum from the mouse as follows: a. Anesthetize the mouse with 3% to 5% isoflurane until it is fully unconscious. b. Open the chest cavity of the mouse by making an incision through the abdominal wall and extending it to the upper chest. Avoid cutting the diaphragm. c. Hold the right lung and cut it off through the root of the lung. Blood will then flow into the chest cavity. d. Collect the blood by aspirating slowly into a 1-ml syringe. e. Sacrifice the mouse promptly by cervical dislocation. Do not allow it to regain consciousness. f. Allow 15 min for blood to clot. Separate the serum using serum tubes with gel and clot activator. Centrifuge 2 min at 8000 × g, 4◦ C. g. Dispense the serum into 5- to 10-μl aliquots per tube and freeze for up to at least 3 years at −80◦ C.

SPECTROPHOTOMETRIC QUANTITATION Muscle samples from which EBD will be extracted may be fresh, snap frozen without freezing medium, or frozen in O.C.T. freezing compound. If muscles are frozen in O.C.T., rinse under cold running water just until the frozen O.C.T. melts off, taking care to keep the muscle specimen frozen as much as possible. Muscle samples must be frozen prior to grinding to generate a fine powder from which the EBD is extracted using N,N-dimethyl formamide (DMF).

BASIC PROTOCOL 4

The extracted EBD can be quantitated using either a spectrophotometer or a fluorimeter. Using a spectrophotometric microplate reader is straightforward, fast, and convenient. In this case, absorbance is measured at 630 nm. Standards should be prepared such that the samples will fall within range of the standards. This protocol describes the range that is convenient for studies with leg muscle samples from mdx and wild-type mice.

Materials EBD (see recipe) N,N-dimethyl formamide (DMF) Liquid nitrogen Muscle samples Current Protocols in Mouse Biology

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Spreadsheet (e.g., Microsoft Excel) Mortar and pestle (mortar must be cooled with liquid N2 ) 1.5-ml microcentrifuge tubes Vortex mixer Rotator (e.g., Rugged rotator, Glas-Col) or equivalent Microcentrifuge 96-well UV-transparent microplates (BD Falcon, cat. no. 353261) Microplate reader capable of reading absorbance at 630 nm (SpectraMAX Plus, Molecular Devices) Prepare for EBD extraction 1. Generate a spreadsheet with all sample numbers and spaces to record the whole muscle sample weight as well as the quantity of powdered muscle used for the extraction. 2. Prepare dilutions of EBD in DMF for a standard curve as shown in Figure 9, beginning with 8.0 μg/ml EBD. Prepare serial two-fold dilutions down to 0.061 μg/ml. Prepare additional standards in 0.5-μg/ml increments to cover the range from 8.0 μg/ml down to 3.0 μg/ml. If absorbance from samples falls outside the range of standards used, then prepare additional concentrations of EBD as needed.

3. Set up a mortar and pestle with a container of liquid nitrogen that can be poured into the chilling chamber under the mortar as needed to keep tissue samples extremely cold. 4. Pre-chill 1.5-ml microcentrifuge tubes on dry ice.

Perform EBD extraction 5. Weigh and record each muscle sample as fast as possible to keep cold. 6. Freeze muscle samples in liquid nitrogen or at −80◦ C. When a tissue sample is placed into a tube prior to freezing, it often sticks to the side of the tube, making it difficult to remove. Dropping the piece of tissue directly into liquid nitrogen allows it to freeze without sticking to the tube and then remove from liquid nitrogen with forceps.

7. Grind the frozen muscle sample into powder using a mortar and pestle. Place ground sample into a pre-chilled 1.5-ml microcentrifuge tube (or larger if needed), keeping tissue as cold as possible.

Evans blue dye ( g/ml)

10 8 6 4

0.024

2 0 0.0

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y 26.858x R2 0.9998

0.1

0.2

0.3

0.4

Absorbance (630 nm)

Figure 9 Standard curve prepared with EBD. Dilutions of EBD in DMF were used to generate a standard curve. A graph and standard curve formula can be generated by plotting the data using Microsoft Excel, although this graph was generated by GraphPad Prism.

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8. Weigh 50 mg of this muscle powder as quickly as possible to keep cold and place into a clean pre-chilled 1.5-ml microcentrifuge tube. Tare the tube before adding muscle powder and weigh afterward to determine the actual weight. 9. Add 1 ml DMF to each tube of muscle powder and mix well using a vortexer. 10. Incubate with continual rotational mixing (on a rotator) for 24 hr at 25◦ C. 11. Centrifuge tube 10 min at 6500 × g, 25◦ C. 12. Transfer the blue supernatant containing the EBD to a fresh 1.5-ml microcentrifuge tube.

Quantitate EBD in muscle samples 13. Prepare a 96-well UV-transparent microplate with duplicate samples by pipetting 200 μl sample to each well. Be sure to include the standard curve dilutions on the plate as well as a blank with DMF only. 14. Use a spectrophotometric microplate reader to measure the absorbance at 630 nm (Amax ). 15. Use Excel to determine the concentration of EBD in each sample using the formula generated from the standard curve (see Fig. 9).

Analyze EBD in muscle samples 16. Compare whole limbs (see example in Fig. 10A) as follows: a. Determinethe weighted percentages of EBD in the entire hind limb using the formula: ( A1 M1 + A2 M2 + A3 M3 + A4 M4 + A5 M5 )/Mt , where Ai is the amount of EBD extracted per mass of muscle from muscle group i and Mi is the original weight of that muscle group, and Mt is the total weight of all limb muscles. b. Calculate the weighted average for each leg and then calculate the average value for the treatment group (all the legs in the group). Even if some of the muscle was first used for immunohistochemistry or some other procedure, make sure to use the original muscle sample weight when calculating the weighted average of EBD in the leg.

A

EBD in whole limb

B

all muscle groups 120

g EBD/g muscle

g EBD/g muscle

40 30 20 10 0

100 80 60 40 20 0

unexercised exercised Treatment of mice

unexercised exercised Treatment of mice

Figure 10 Extracted EBD from hind leg muscles. The 15- to 17-month-old mdx mice were either unexercised or were exercised by running on a treadmill 30 min at a speed of 12 m/min, n = 12 legs for each group of mice. (A) The weighted average amount of EBD in the whole leg is shown for each group of mice. EBD extracted from the exercised legs was 26% higher than from the unexercised legs (p = 0.0026). (B) Each data point indicates the amount of EBD in one muscle sample. The average amount of EBD extracted from muscle samples of the exercised mice was 39% higher than from those of the unexercised mice (p < 0.0001). Averages are indicated by a bar ± SD. See Wooddell et al. (2010) for additional details.

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17. Compare muscle samples as collectives (Fig. 10B). As described for analysis of EBD+ myofibers, the power of analysis can be improved by comparing samples as collective groups of muscles. In this case, the weight of the samples is not taken into account and all samples contribute equally to the resulting average.

18. In cases where one leg of the mouse can be used as a control for a treatment given to the other leg, then pairs of muscle groups from the left and right legs can be compared using a two-tailed, Wilcoxon signed rank test for statistical analysis.

REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps.

Evans blue dye (EBD) in PBS (1% weight/volume) Dissolve 1 g Evans blue dye (EBD; Sigma-Aldrich, cat. no. E2129) in 100 ml sterile, filtered, phosphate buffered saline (PBS), pH 7.5, and use fresh.

Evans blue dye (EBD) in saline (5 mg/ml) Dissolve 200 mg Evans blue dye (EBD; Sigma-Aldrich, cat. no. E2129) in 40 ml sterile, commercially-prepared physiological saline (0.90% NaCl). Filter through a 0.22-μm filter. Store at 4◦ C.

COMMENTARY Background Information

Myofiber Damage Evaluation by Evans Blue Dye Injection

Evans blue dye is an exclusion dye that is widely used to assess cellular integrity, due to its inability to enter (and, therefore, stain) intact cells. The propensity of the dye to bind to serum albumin following injection makes it a useful tool for assessing a number of physiological parameters, and the spectral characteristics of the dye facilitate tissue- and cellularlevel visualization. In this manuscript, several methods for injecting and quantitatively measuring uptake of Evans blue dye in mouse hindlimb skeletal muscle are described to provide the researcher with a flexible set of tools for assessment of skeletal muscle damage. The different quantitation protocols balance workload with specificity and resolution, and further discussion of these qualities is presented in the Critical Parameters and Troubleshooting section. Damaged muscles containing EBD can be observed macroscopically due to the bright blue stain (Fig. 11) or microscopically by the red auto-fluorescence of the dye in muscle sections (Fig. 12). Myofibers with histologically distinct necrosis (cell death) always stain positive for EBD and myofibers with an intact sarcolemma do not stain (Matsuda et al., 1995; Brussee et al., 1997; Straub et al., 1997; Archer et al., 2006). Blood proteins can be detected in spaces occupied by the necrotic myofibers (Fig. 6). EBD can also be present within myofibers that appear morphologically nor-

mal in hematoxylin and eosin (H&E)-stained sections (Brussee et al., 1997; Hamer et al., 2002) and EBD uptake into leaky myofibers does not always reflect severe myofiber damage (necrosis) that will provoke regeneration and require myogenesis (reviewed in Grounds et al., 2008). Temporary lesions do not necessarily cause myofiber death because the membrane of the myofiber can reseal (Doherty and McNally, 2003; McNeil and Kirchhausen, 2005). Most EBD-stained myofibers in mdx mice are intensely stained and clearly necrotic, but those that resemble normal (non-necrotic) myofibers when viewed by H&E stain only lightly with EBD (Figs. 5 and 6). EBD is also used as an end-point to demonstrate an efficacious therapy in dystrophic mdx mice (Matsuda et al., 1995; Straub et al., 1997; Grounds et al., 2008; Zhang et al., 2010). However, demonstration of a therapeutic effect requires that one first determines the natural variation between animals, which is extremely high in dystrophic mdx mice (for an example, see Fig. 11; Straub et al., 1997; Grounds et al., 2008). The therapeutic effect must be significantly greater than the natural variation between animals in order to detect and quantify efficacy (Wooddell et al., 2010). If one leg of the animal receives a therapeutic treatment and the other leg receives a control treatment, then the variation between the left and right legs of individual mice must also be determined.

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Figure 11 Macroscopic evaluation of two 9-week-old dystrophic (mdx) litter-mates given i.v. injections of EBD (mouse 1, a and b; mouse 2, c and d). Mice were sacrificed 6 hr after EBD injection and fixed in 8% formaldehyde solution. See original article for additional information (Straub et al., 1997; Rockefeller University Press. Originally published in J. Cell Biol. 139:375-385. doi: 10.1083/jcb.139.2.375).

Figure 12 Tibialis anterior muscle in cross section showing positive staining for EBD in dystrophic (mdx) myofibers 48 hr after an exercise-induced muscle damage protocol. EBD was administered via i.p. injection 24 hr prior to sacrifice.

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Critical Parameters and Troubleshooting Timing of muscle evaluation relative to EBD delivery In published reports of EBD studies, dystrophic mice have been sacrificed at varying times between 3 and 96 hr after injection of the EBD, most commonly at 12 to 24 hr (Straub et al., 1997; Richard et al., 2000; Hamer et al., 2002; Kobinger et al., 2003; Chakkalakal et al., 2004; Sokolow et al., 2004; Sher et al., 2006). In addition to the EBD that leaks into muscle fibers with permeable cell membranes and can be visualized for at least 96 hr, some EBD can also be visualized in the interstitial space for just as long (Fig. 13). In the authors’ studies, very little EBD circulating in the blood was detected 24 hr after it was intravenously injected (Wooddell et al., 2010). In muscle sections taken during the first few hours after EBD delivery, up to ∼6 hr, EBD appears to have incompletely penetrated into the myofibers (Straub et al., 1997; Wooddell et al., unpub. observ.). Even with this characteristic, EBD-positive (EBD+ ) myofibers can be distinguished as early as 1 hr after i.v. injection of EBD (Wooddell et al., unpub. observ.). Exercise protocols to challenge the muscles Muscle contractions can induce injury in the fragile myofiber plasma membranes of dystrophic mdx mice. When subjected to sustained exercise and then injected with EBD, the muscles of mdx mice accumulate the dye much more than those of normal mice (Brussee

et al., 1997; Tinsley et al., 1998; Vilquin et al., 1998; Hamer et al., 2002). Some mouse strains with muscle defects do not exhibit a particularly strong myopathy when the mice are sedentary. This is the case for mdx mice. In these mice, dystropathology can be exacerbated by subjecting the mice to an exercise protocol. The most commonly used exercise protocols are treadmill running under a number of conditions and balancing on a rotating rod (rotarod) (Grounds et al., 2008; Wooddell et al., 2010; Marcaletti et al., 2011). Exercise protocols are beyond the scope of this paper and are described elsewhere; here, treadmill running is used in some examples. Additional protocols (SOPs) for exercising mdx mice are available on the TREAT-NMD Neuromuscular Network Website http://www.treatnmd.eu/research/preclinical/dmd-sops/. The mdx mice may be reluctant to participate in exercise protocols, such as treadmill running, after being injected with EBD (Wooddell et al., unpub. observ.). For this reason it is more suitable to perform the exercise regimen prior to EBD injection, generally followed 20 to 30 min later by an EBD injection. If mdx mice must be injected with EBD prior to an exercise regimen, allowing them at least a 30-min rest may make them better able to perform the exercise. Quantitative methods for assessment of EBD uptake As demonstrated by Straub et al. (1997), macroscopic evaluation of EBD in the whole

6 myofiber contrast

5 EBD signal strength

interstitium 4

3

2

1

0 88

80

72

64

56

48

40

32

24

16

8

0

Time of EBD injection prior to sampling (hr)

Myofiber Damage Evaluation by Evans Blue Dye Injection

Figure 13 Strength of EBD fluorescent signal in skeletal myofibers and interstitium of mdx mice given an i.p. injection of 1% EBD in PBS at various times before sacrifice and muscle harvest. Adapted from Hamer et al. (2002).

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mouse is highly useful for a qualitative assessment of phenotypes and to visualize the variation between animals in a group (Fig. 11). Some other approaches that allow quantitation of EBD in the muscles are based on microscopic evaluation or spectrophotometric detection of EBD that is extracted from the muscles. Microscope-based methods of analysis Muscle damage in dystrophic mdx mice tends to be in regions composed of multiple damaged muscle fibers. For example, see the myofibers labeled as darkly (intensely) EBD-stained in the sectioned muscle specimen shown in Figure 5A. Such regions in images captured by fluorescence microscopy can be quantitated by morphometric analysis (Fig. 1). Another option for quantitating EBDstained muscle is to count EBD+ myofibers in cross-sections of muscle samples. This method is suitable for determining a therapeutic effect from a treatment that affects some but not all muscle fibers within the leg, e.g., following gene therapy (Fig. 3). In this example, the muscle fibers that received a dystrophin-expressing plasmid DNA (pDNA) were protected from exercise-induced muscle damage. The dystrophin-positive muscle fibers excluded EBD while neighboring muscle fibers that did not receive the therapeutic plasmid were found to be leaky and EBDstained. This method of visualizing the EBD+ myofibers also allows one to distinguish between those that stain intensely with EBD (generally necrotic) and those that stain only lightly. Microscopic analyses are labor intensive. Multiple representative muscle specimens from throughout the limb need to be examined to assess EBD staining in the whole limb. For mice in which morphological features appear to be equally divided throughout the muscle from tendon to tendon, such as reported for unexercised mdx mice, it is sufficient to take specimens from one location within the muscle, i.e., the middle (van Putten et al., 2010). Nevertheless, the leg is composed of many muscles. A researcher may choose to use microscopic techniques to visualize specific EBD-stained muscles in detail rather than evaluating muscle specimens sampled from the whole leg. Myofibers are long, thin muscle cells. Determining the percentages of them that are EBD-positive requires the researcher to quantitate a representative number in cross-sections

of muscle specimens. The sections should be cut perpendicular to the muscle fibers to obtain high-quality cross-sectional images. The easiest way to accomplish this is by taking great care to keep the muscle specimens properly oriented while freezing them. EBD staining of the myofibers is better preserved when muscle specimens are frozen in a freezing compound such as O.C.T. than when they are embedded in paraffin and processed for histological evaluation (Hamer et al., 2002). EBD causes high background fluorescence throughout paraffin-embedded muscle specimens (Wooddell et al., unpub. observ.). Muscle specimens frozen in O.C.T. and then sectioned are ideal for immunohistochemistry (Figs. 3 and 6). Additionally, sections of muscle can be cut from the frozen blocks of O.C.T. and then stained for histological evaluation with H&E (Fig. 5). Although the quality of the H&E staining is not as sharp as it is when sections are cut from paraffin blocks, this approach allows adjacent sections to be evaluated by H&E as well as by EBD fluorescence and IHC, even with multiple antibodies in sequential sections (Fig. 6). Immunohistochemistry EBD staining is commonly used to evaluate mouse models for myopathies, which may be caused by defects in myofiber cell membrane proteins such as dystrophin. Immunostaining of such membrane proteins in wild-type mice allows the researcher to distinguish the individual myofibers. However, genetic defects that result in muscle myopathies often also cause disruption of the dystrophinglycoprotein complex. As a consequence, proteins that are localized to the myofiber membrane in wild-type mice may not be similarly located in mice that have myopathies. In this case, immunostaining of blood proteins can serve the purpose of outlining individual myofibers. Use of an antibody to detect a blood protein serves another function in conjunction with EBD staining. Some of the EBD+ myofibers are clearly damaged and stain intensely (Figs. 5 and 6). Blood proteins can be found in these myofibers (Fig. 6). Overlaying the signals from the red-fluorescing EBD with the green signal from fluorescein isothiocyanate (FITC)-labeled antibodies results in a bright yellow-green signal in the severely damaged myofibers (Fig. 3A). The myofibers that stain lightly with EBD exclude the blood proteins. They appear red in the microscope images rather than yellow-green (Fig. 3B).

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g EBD/g muscle

60

40

20

o mdx ( E)

o mdx ( E)

y mdx ( E)

y mdx ( E)

C57 ( E)

C57 ( E)

0

Groups of unexercised and exercised mice

Figure 14 EBD extracted from the whole leg of unexercised and exercised young and old mdx mice and wild-type mice. Groups of mice were unexercised or exercised by running on a treadmill for 10 to 30 min at 12 m/min. The average amount of EBD extracted per gram muscle in each leg is shown for unexercised (E–) and exercised (E+) mice: male wild-type C57Bl/6 mice (C57, n = 4 mice, 8 legs); male and female young adult mdx4Cv mice, 3 to 7 months old (y mdx, n = 5 mice, 10 legs); and male and female 15 to 17 months old mdx-10ScSn mice (o mdx, n = 6 mice, 12 legs). Figure taken from Wooddell et al. (2010).

The researcher must decide whether to count only the intensely-stained EBD+ myofibers or to include all EBD+ myofibers, the intensely and lightly stained ones. It was observed that the intensely-stained myofibers increased following exercise in old mdx mice, but the lightly stained ones did not (Wooddell et al., 2010).

Myofiber Damage Evaluation by Evans Blue Dye Injection

Spectrophotometric quantitation An alternative, faster, simpler, and highly quantitative method of evaluating EBD in the muscles is to grind up the muscle samples, extract EBD from them, and quantitate the EBD spectrophotometrically using a microplate reader. This approach reduces the sampling variation inherent in cutting sections from specimens. This spectrophotometric method may be employed when the amount of EBD taken up by muscles in one group of mice differs significantly from that in another, as shown in Figure 14. The method is not suitable for small differences between experimental groups because EBD in the vasculature and especially that in the interstitium between muscle fibers causes high background signal in all animals. Care should be taken to evaluate the muscles at a time when the EBD signal in the myofibers is sufficiently high and the background of EBD in the interstitial space is sufficiently low (Fig. 13). The timing of muscle harvest relative to the time of EBD injection affects this background. Sacrificing mice 24 hr after EBD injection is recommended when EBD is

to be extracted from the muscles and evaluated by the spectrophotometric method. A fluorimeter can also be used to quantitate the extracted EBD, but the conditions would need to be worked out for individual instruments. Only a cuvette-reading fluorimeter (VARIAN model Cary Eclipse) was used by the authors, a much slower process than using a microplate reader. With this fluorimeter, excitation was set at 640 nm; emission at 680 nm; slit width at 5 nm and the voltage at 600 V. DMF will dissolve some plastic cuvettes, so plastic ware must be tested prior to adding samples containing DMF. Diluting the DMFextracted EBD samples in water resulted in an opaque liquid that caused scatter during fluorimeter reading, so this option for decreasing the DMF concentration is not recommended. DMF can be handled in polypropylene tubes and in the microplates described in Basic Protocol 4 for spectrophotometric microplate reading. When an experiment contains a small number of animals and the variability between them in terms of muscle uptake of EBD is expected to be broad, the sample size must be large enough to determine if there are any statistically significant differences between treatment groups. This is a common dilemma when using mdx mice. The statistical power of analysis can be increased by increasing the sample size, either by using more animals or by evaluating more samples within animals. In Figure 10, for example, all of the five groups of muscles from all of the mice in one treatment

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group were compared as collectives to all the muscle samples of all the mice in another treatment group.

Anticipated Results EBD has been used to characterize mice with muscle defects and to evaluate therapies, frequently with quantitation performed by the morphometric method (Brussee et al., 1997; Straub et al., 1997; Richard et al., 2000; Kobinger et al., 2003; Chakkalakal et al., 2004; Shavlakadze et al., 2004; Sokolow et al., 2004; Archer et al., 2006; Minetti et al., 2006; Li et al., 2008; Miura et al., 2009; Piers et al., 2011). See Figure 1 for an example. EBD is also used without quantitation for qualitative examination of muscle. In this case, it is often used to visualize co-localization or lack thereof with other cellular proteins or with histological features (Straub et al., 1997; Cifuentes-Diaz et al., 2001; Hamer et al., 2002; Durbeej et al., 2003; Goyenvalle et al., 2004; Sher et al., 2006; Fougerousse et al., 2007; Wang et al., 2008). When EBD is used as an assay for therapy studies, the evaluation can be quantitative and/or qualitative (Shavlakadze et al., 2004; Wang et al., 2008; Zhang et al., 2010; Figs. 3 and 8). The hind limb muscles are commonly evaluated, but EBD can also be used to evaluate other mouse muscles such as those in the diaphragm (Voisin et al., 2005) or the heart (Bostick et al., 2009). In some cases, macroscopic visualization of the blue stain can be used to characterize a mouse strain or evaluate a therapeutic (Straub et al., 1997; Voisin et al., 2005). See Figure 11 for example. Use wild-type na¨ıve mice as a control when working with other mouse strains or experimental conditions expected to cause muscle damage. Particularly when using the spectrophotometric method to quantitate EBD in muscles, it is important to remember that much of the EBD is in the interstitial space. The consequence of this is that a substantial amount of EBD can be extracted from wild-type mouse muscle, as shown in Figure 14. Approximately 10 μg EBD/g leg muscle was extracted from wild-type mice, whereas >30 μg EBD/g muscle was extracted from 3- to 7-month-old mdx mice. The amount of EBD in the interstitial space decreases over time as Hamer et al. (2002) showed (Fig. 13), but the EBD within the damaged myofibers also decreases over time. Damaged myofibers are replaced by newly formed myofibers in the course of a few days.

Mdx mice are notorious for exhibiting high levels of variation in the extent of dystropathology. This will be reflected in the analysis of EBD+ myofibers (percent crosssectional area or percent counted myofibers). It must also be noted that if analyzing myofiber necrosis and EBD+ myofibers in the same muscle section, results for EBD+ myofibers will often be higher, as not all leaky myofibers (which will stain positive for EBD) will undergo necrosis. See Table 1 for examples of anticipated results in mdx mice. Mouse age and their spontaneous activity can complicate experimental design. The physiology of dystrophic mdx mouse muscle, for example, changes as the mice age (Coulton et al., 1988; McGeachie et al., 1993; Lefaucheur et al., 1995). Measuring a significant difference in muscles of the whole legs of exercised compared to unexercised mdx mice can be more challenging in 2- to 6-month-old compared to 13- to 19-month-old mice (Fig. 7 and Wooddell et al., 2010). The younger mice have much more variable levels of EBD staining than the older mice, even from one leg to the other. This appears to be due to the higher spontaneous activity level of young mice compared to sedate older mice.

Time Considerations As with any animal experiment, the greatest time consideration is that of breeding or otherwise obtaining the desired number of mice of the appropriate age and gender. If antibodies need to be produced for immunohistochemistry, allow at least 4 weeks for this process. Injection of EBD (see Basic Protocol 1) is a quick procedure, but allow ∼30 min prior to injection for mice to be placed under a heat lamp when performing the intravenous injection. Anesthetizing the mouse takes only 2 to 3 min. The amount of time required to produce skeletal muscle sections on glass slides suitable for morphometric EBD imaging without immunohistochemistry (see Basic Protocol 2) is fairly short, depending on the number of samples. Immediately after freezing the muscle samples, they can be cut on a cryostat and tissue sections produced. Preparing frozen muscle samples avoids waiting for paraformaldehyde fixation and overnight paraffin processing. The speed of cryostat cutting depends on the individual researcher, but generally a group of eight muscle samples can easily be cut in a 3-hr cryostat session. EBD is autofluorescent and, thus, no additional

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Table 1 Examples of Anticipated Results from mdx Mice

Results in specified muscle (% EBD+ cross-sectional area)

Reference

Description of mice

Piers et al. (2011)

10- to 13-week-old male mdx mice, subjected to exercise-induced muscle damage

Tibialis anterior (up to 20%)

Shavlakadze et al. (2004)

19- to 26-day-old mdx mice

Tibialis anterior (up to 60%); Quadriceps (up to 30%); Diaphragm (up to 40%)

Radley-Crabb (unpub. observ.)

12-week-old male mdx mice, unexercised

Quadriceps (up to 20%); Gastrocnemius (up to 23%)

Radley-Crabb (unpub. observ.)

12-week-old male mdx mice, subjected to one 30-min treadmill running exercise

Quadriceps (up to 30%); Gastrocnemius (up to 32%)

Myofiber Damage Evaluation by Evans Blue Dye Injection

staining is needed. Once cut, muscle sections are simply fixed with acetone and covered with glass cover slips. Image capture and image analysis, however, are two rather timeconsuming steps. The time taken to capture an image of a whole muscle in cross section depends on the size of the muscle, the capabilities of the image capture software, and the microscopic magnification. Capturing an entire mdx quadriceps muscle section in cross section at 10× magnification requires ∼20 min. Image analysis is also a time-consuming step and must be done accurately. This step also depends on the size of the muscle and the amount of muscle damage. Analyzing an entire mdx quadriceps muscle generally takes 20 to 30 min, whereas a smaller tibialis anterior muscle may take 10 to 15 min. Basic Protocol 3 is fairly quick once the researcher has a feel for the dissection. Mice are anesthetized and sacrificed. Then the leg muscles are cut into pieces, weighed, and frozen. This is a convenient stopping point before proceeding with Support Protocol 1. Support Protocol 1 is the most laborintensive of these protocols, especially counting of the myofibers. In addition to the time required for cryostat sectioning, ∼2 hr are required for immunostaining a batch of slides. Then allow 2 to 3 hr per leg to take ten images across the diameters of each of the five groups of muscles using two different filters. Image capture should be performed within 2 days of immunostaining. Once the images are acquired, positive myofibers are counted on the computer screen. This takes more time when there are many EBD+ myofibers, if the researcher wishes to distinguish the lightly from the intensely stained myofibers, or if specifically immunostained myofibers are to be counted. The total number of myofibers can

be counted from printed images for determination of the percentage of EBD+ myofibers. Allow 1 day per leg for counting all myofibers in the cross-sectional images for EBD and specific immunostaining staining. The time can be reduced if only the positive myofibers in each image are counted and these are reported as EBD+ myofibers/view (Fig. 7) rather than as a percentage of the total. The time can be further reduced by selecting just a subset of the muscle pieces for evaluation. Support Protocol 2 is for production of antibodies. After a suitable plasmid and animals are acquired, allow at least 4 weeks for the animals to develop antibodies. Hands-on time is minimal. The plasmid injection procedure requires some skill, but each injection typically takes <10 min. Immunohistochemistry testing can be performed in 1 day. It does not involve myofiber counting. Tissue grinding accounts for the majority of hands-on time required to perform Basic Protocol 4. After samples are ground, EBD is extracted over 24 hr. The microplate reader assay is quick. Overall, nine samples can be processed in an average of 4 hr hands-on time.

Literature Cited Archer, J.D., Vargas, C.C., and Anderson, J.E. 2006. Persistent and improved functional gain in mdx dystrophic mice after treatment with L-arginine and deflazacort. Faseb J. 20:738-740. Bates, M.K., Zhang, G., Sebestyen, M.G., Neal, Z.C., Wolff, J.A., and Herweijer, H. 2006. Genetic immunization for antibody generation in research animals by intravenous delivery of plasmid DNA. Biotechniques 40:199-208. Bostick, B., Yue, Y., Long, C., Marschalk, N., Fine, D.M., Chen, J., and Duan, D. 2009. Cardiac expression of a mini-dystrophin that normalizes skeletal muscle force only partially restores

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heart function in aged mdx mice. Mol. Ther. 17:253-261. Brussee, V., Tardif, F., and Tremblay, J.P. 1997. Muscle fibers of mdx mice are more vulnerable to exercise than those of normal mice. Neuromuscul. Disord. 7:487-492. Chakkalakal, J.V., Harrison, M.A., Carbonetto, S., Chin, E., Michel, R.N., and Jasmin, B.J. 2004. Stimulation of calcineurin signaling attenuates the dystrophic pathology in mdx mice. Hum. Mol. Genet. 13:379-388. Cifuentes-Diaz, C., Frugier, T., Tiziano, F.D., Lacene, E., Roblot, N., Joshi, V., Moreau, M.H., and Melki, J. 2001. Deletion of murine SMN exon 7 directed to skeletal muscle leads to severe muscular dystrophy. J. Cell Biol. 152:11071114. Coulton, G.R., Morgan, J.E., Partridge, T.A., and Sloper, J.C. 1988. The mdx mouse skeletal muscle myopathy: I. A histological, morphometric and biochemical investigation. Neuropathol. Appl. Neurobiol. 14:53-70. Doherty, K.R. and McNally, E.M. 2003. Repairing the tears: Dysferlin in muscle membrane repair. Trends Mol. Med. 9:327-330. Durbeej, M., Sawatzki, S.M., Barresi, R., Schmainda, K.M., Allamand, V., Michele, D.E., and Campbell, K.P. 2003. Gene transfer establishes primacy of striated vs. smooth muscle sarcoglycan complex in limb-girdle muscular dystrophy. Proc. Natl. Acad. Sci. U.S.A. 100:8910-8915. Fougerousse, F., Bartoli, M., Poupiot, J., Arandel, L., Durand, M., Guerchet, N., Gicquel, E., Danos, O., and Richard, I. 2007. Phenotypic correction of alpha-sarcoglycan deficiency by intraarterial injection of a muscle-specific serotype 1 rAAV vector. Mol. Ther. 15:53-61. Goyenvalle, A., Vulin, A., Fougerousse, F., Leturcq, F., Kaplan, J.C., Garcia, L., and Danos, O. 2004. Rescue of dystrophic muscle through U7 snRNA-mediated exon skipping. Science 306:1796-1799. Grounds, M.D., Radley, H.G., Lynch, G.S., Nagaraju, K., and De Luca, A. 2008. Towards developing standard operating procedures for pre-clinical testing in the mdx mouse model of Duchenne muscular dystrophy. Neurobiol. Dis. 31:1-19. Hagstrom, J.E., Hegge, J., Zhang, G., Noble, M., Budker, V., Lewis, D.L., Herweijer, H., and Wolff, J.A. 2004. A facile nonviral method for delivering genes and siRNAs to skeletal muscle of mammalian limbs. Mol. Ther. 10:386-398.

Lefaucheur, J.P., Pastoret, C., and Sebille, A. 1995. Phenotype of dystrophinopathy in old mdx mice. Anat. Rec. 242:70-76. Li, D., Yue, Y., and Duan, D. 2008. Preservation of muscle force in Mdx3cv mice correlates with low-level expression of a near full-length dystrophin protein. Am. J. Pathol. 172:13321341. Marcaletti, S., Thomas, C., and Feige, J.N. 2011. Exercise performance tests in mice. Curr. Protoc. Mouse Biol. 1:141-154. Matsuda, R., Nishikawa, A., and Tanaka, H. 1995. Visualization of dystrophic muscle fibers in mdx mouse by vital staining with Evans blue: Evidence of apoptosis in dystrophin-deficient muscle. J. Biochem. 118:959-964. McGeachie, J.K., Grounds, M.D., Partridge, T.A., and Morgan, J.E. 1993. Age-related changes in replication of myogenic cells in mdx mice: Quantitative autoradiographic studies. J. Neurol. Sci. 119:169-179. McNeil, P.L. and Kirchhausen, T. 2005. An emergency response team for membrane repair. Nat. Rev. Mol. Cell Biol. 6:499-505. Minetti, G.C., Colussi, C., Adami, R., Serra, C., Mozzetta, C., Parente, V., Fortuni, S., Straino, S., Sampaolesi, M., Di Padova, M., Illi, B., Gallinari, P., Steinkuhler, C., Capogrossi, M.C., Sartorelli, V., Bottinelli, R., Gaetano, C., and Puri, P.L. 2006. Functional and morphological recovery of dystrophic muscles in mice treated with deacetylase inhibitors. Nat. Med. 12:11471150. Miura, P., Chakkalakal, J.V., Boudreault, L., Belanger, G., Hebert, R.L., Renaud, J.M., and Jasmin, B.J. 2009. Pharmacological activation of PPARbeta/delta stimulates utrophin A expression in skeletal muscle fibers and restores sarcolemmal integrity in mature mdx mice. Hum. Mol. Genet. 18:4640-4649. Piers, A.T., Lavin, T., Radley-Crabb, H.G., Bakker, A.J., Grounds, M.D., and Pinniger, G.J. 2011. Blockade of TNF in vivo using cV1q antibody reduces contractile dysfunction of skeletal muscle in response to eccentric exercise in dystrophic mdx and normal mice. Neuromuscul. Disord. 21:132-141. Reeve, E.B. 1957. The contribution of I 131-labeled proteins to measurements of blood volume. Ann. N.Y. Acad. Sci. 70:137-147.

Hamer, P.W., McGeachie, J.M., Davies, M.J., and Grounds, M.D. 2002. Evans blue dye as an in vivo marker of myofibre damage: Optimizing parameters for detecting initial myofibre membrane permeability. J. Anat. 200:69-79.

Richard, I., Roudaut, C., Marchand, S., Baghdiguian, S., Herasse, M., Stockholm, D., Ono, Y., Suel, L., Bourg, N., Sorimachi, H., Lefranc, G., Fardeau, M., Sebille, A., and Beckmann, J.S. 2000. Loss of calpain 3 proteolytic activity leads to muscular dystrophy and to apoptosisassociated IkappaBalpha/nuclear factor kappaB pathway perturbation in mice. J. Cell Biol. 151:1583-1590.

Kobinger, G.P., Louboutin, J.P., Barton, E.R., Sweeney, H.L., and Wilson, J.M. 2003. Correction of the dystrophic phenotype by in vivo targeting of muscle progenitor cells. Hum. Gene Ther. 14:1441-1449.

Shavlakadze, T., White, J., Hoh, J.F. Rosenthal, N., and Grounds, M.D. 2004. Targeted expression of insulin-like growth factor-I reduces early myofiber necrosis in dystrophic mdx mice. Mol. Ther. 10:829-843.

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Sher, R.B., Aoyama, C., Huebsch, K.A., Ji, S., Kerner, J., Yang, Y., Frankel, W.N., Hoppel, C.L., Wood, P.A., Vance, D.E., and Cox, G.A. 2006. A rostrocaudal muscular dystrophy caused by a defect in choline kinase beta, the first enzyme in phosphatidylcholine biosynthesis. J. Biol. Chem. 281:4938-4948. Sokolow, S., Manto, M., Gailly, P., Molgo, J., Vandebrouck, C., Vanderwinden, J.M., Herchuelz, A., and Schurmans, S. 2004. Impaired neuromuscular transmission and skeletal muscle fiber necrosis in mice lacking Na/Ca exchanger 3. J. Clin. Invest. 113:265-273. Straub, V., Rafael, J.A., Chamberlain, J.S., and Campbell, K.P. 1997. Animal models for muscular dystrophy show different patterns of sarcolemmal disruption. J. Cell Biol. 139:375385. Tinsley, J., Deconinck, N., Fisher, R., Kahn, D., Phelps, S., Gillis, J.M., and Davies, K. 1998. Expression of full-length utrophin prevents muscular dystrophy in mdx mice. Nature Med. 4:14411444. van Putten, M., de Winter, C., van Roon-Mom, W., van Ommen, G.J., ’t Hoen, P.A., and AartsmaRus, A. 2010. A 3 months mild functional test regime does not affect disease parameters in

young mdx mice. Neuromuscul. Disord. 20:273280. Vilquin, J.T., Brussee, V., Asselin, I., Kinoshita, I., Gingras, M., and Tremblay, J.P. 1998. Evidence of mdx mouse skeletal muscle fragility in vivo by eccentric running exercise. Muscle Nerve 21:567-576. Voisin, V., Sebrie, C., Matecki, S., Yu, H., Gillet, B., Ramonatxo, M., Israel, M., and De la Porte, S. 2005. l-Arginine improves dystrophic phenotype in mdx mice. Neurobiol. Dis. 20:123-130. Wang, B., Li, J., Qiao, C., Chen, C., Hu, P., Zhu, X., Zhou, L., Bogan, J., Kornegay, J., and Xiao, X. 2008. A canine minidystrophin is functional and therapeutic in mdx mice. Gene Ther. 15:10991106. Wooddell, C.I., Zhang, G., Griffin, J.B., Hegge, J.O., Huss, T., and Wolff, J.A. 2010. Use of Evans blue dye to compare limb muscles in exercised young and old mdx mice. Muscle Nerve. 41:487-499. Zhang, G., Wooddell, C.I., Hegge, J.O., Griffin, J.B., Huss, T., Braun, S., and Wolff, J.A. 2010. Functional efficacy of dystrophin expression from plasmids delivered to mdx mice by hydrodynamic limb vein injection. Hum. Gene Ther. 21:221-237.

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One-Step Vital Staining of Presynaptic Terminals and Post-Synaptic Receptors at Neuromuscular Junctions in Mouse Skeletal Muscle Richard R. Ribchester1 1

Euan MacDonald Centre for Motor Neurone Disease Research, University of Edinburgh, Edinburgh, Scotland, United Kingdom

ABSTRACT This protocol describes vital staining of neuromuscular junctions in the mouse triangularis sterni muscle in one incubation step, combining presynaptic, motor nerve terminal staining with the styryl dye FM1-43, which labels recycling synaptic vesicles, and TRITC-α-bungarotoxin, which labels acetylcholine receptors in the motor endplate C 2011 by John Wiley & Sons, Inc. membrane. Curr. Protoc. Mouse Biol. 1:489-496  Keywords: neuromuscular junction r fluorescence microscopy r vital staining r styryl dyes r α-bungarotoxin r acetylcholine receptor r synaptic vesicle

The large size and visibility of neuromuscular synapses enabled early light microscopists to define the basic cytological organization of neuromuscular junctions (NMJs). By the early 20th century, silver staining protocols had established a most remarkable feature of NMJs in most vertebrate striated muscle: the mononeuronal innervation of a discrete plaque, occupying 0.1% or less of the total surface area of a muscle fiber. Thus, it was subsequently established, and amply confirmed by all techniques that have superseded silver staining, that in mature animals each motor neuron projects to a discrete muscle, wherein each innervates many muscle fibers (a “motor unit”). However, each muscle fiber is innervated by a collateral branch and terminal derived from the axon of one and only one motor neuron. Moreover, presynaptic and post-synaptic specializations at neuromuscular junctions coincide: motor nerve terminals are intimately associated and align with gutters and folds of motor endplates, which contain a high density of acetylcholine receptors (Costanzo et al., 1999, 2000; Lu et al., 2009). In rodents, the presynaptic and post-synaptic specializations form prenatally (Pun et al., 2002), but the mature pattern of innervation finally emerges during the first three weeks of postnatal development from one of polyneuronal innervation (Brown et al., 1976; Walsh and Lichtman, 2003). In addition, the transformation may be driven partly by differences in synaptic strength and activity-dependent competition (Betz et al., 1980, 1990; Fladby and Jansen, 1987; Buffelli et al., 2003; Kasthuri and Lichtman, 2003). Neuromuscular synapses normally degenerate rapidly (within 18 hr) after peripheral nerve injury, but they are much more slowly removed after nerve injury in the WldS mutant mouse (Gillingwater et al., 2002) and in some forms of motor neuron disease (Schaefer et al., 2005; Murray et al., 2010). Taken together with developmental synaptic remodeling, such observations are consistent with a “compartmental neurodegeneration” hypothesis, according to which axons and synapses are maintained by cellular and molecular mechanisms that are distinct from those that maintain cell bodies (Gillingwater and Ribchester, 2001, 2003). Analysis of neuromuscular synaptic connectivity is therefore

Current Protocols in Mouse Biology 1: 489-496, December 2011 Published online December 2011 in Wiley Online Library (wileyonlinelibrary.com). DOI: 10.1002/9780470942390.mo110128 C 2011 John Wiley & Sons, Inc. Copyright 

BASIC PROTOCOL

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of interest and relevance in the context of development, maintenance, plasticity of motor units, and their vulnerability in different forms of neuromuscular disease. The “one-step” method for labeling of presynaptic and post-synaptic components of neuromuscular junctions given here is very simple, and it combines vital staining of motor nerve terminals with FM1-43 and staining of acetylcholine receptors at motor endplates with TRITC-conjugated α-bungarotoxin. This method is illustrated using isolated preparations of the triangularis sterni muscle, a muscle lining the pleural side of the thorax that is only one or two fibers thick. The results can readily be observed with conventional fluorescence microscopy, using standard FITC and TRITC filter cubes. NOTE: All protocols using live animals must first be reviewed and approved by an Institutional Animal Care and Use Committee (IACUC) and must follow officially approved procedures for care and use of laboratory animals.

Materials Physiological saline (see recipe) Depolarizing saline (see recipe) Adult mice (any laboratory strain, e.g., C57B16; any age or gender) 1 mg/ml FM1-43 or FM1-43FX (see recipe) 500 μg/ml TRITC-α-bungarotoxin stock solution (see recipe) 4% paraformaldehyde, optional Minutien pins Sylgard-lined petri dishes Dissecting microscope fitted with lamps or fiber-optic illumination for both transmitted and incident light Watchmaker’s forceps (nos. 3 and 5) Fine spring scissors Iris scissors Incubator with rocking platform Upright fluorescence microscope or confocal microscope fitted with water-dipping objectives Digital camera and driver/image-processing software/PC Additional reagents and equipment for euthanizing the animal (Donovan and Brown, 2006) 1. Prepare one liter of fresh “normal” physiological saline solution (i.e., normal saline) following the recipe in the Reagents and Solutions section. 2. Freshly prepare 100 ml of a depolarizing mammalian physiological saline solution following the recipe in the Reagents and Solutions section. 3. Sacrifice a mouse using an authorized, legal method approved by the institution where the research is to be conducted (e.g., see Donovan and Brown, 2006). Stunning, immediately followed by cervical dislocation is suitable, swift, and in accordance with approved methods listed under the UK Home Office’s Schedule 1.

4. Using iris scissors, make a circumferential incision through the abdominal skin. Grip the skin rostral to the incision and strip upwards, exposing the thoracic musculature and ribcage. Douse the exposed thorax with normal physiological saline. Pre/PostSynaptic NMJ Staining in Mouse Skeletal Muscle

5. Using iris scissors, carefully cut through the abdominal body wall following the line of the most caudal ribs. Cut through the ribcage laterally using iris scissors and separate from underlying pleura and connective tissue. Remove the ribcage and pin it to a Sylgard-lined petri dish.

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A

200 m

C

B

30 m

Figure 1 (A) Low-power micrograph taken in a dissecting microscope with both transmitted and incident illumination of triangularis sterni muscle preparation. Note the intact muscle fibers oriented vertically and the intramuscular nerve (arrow) running across it. (B) Examples of intramuscular axons and neuromuscular junctions visualized in a conventional fluorescence microscope. Left panels, upper TRITC-α-bungarotoxin staining; lower, FM1-43; Middle panel, merged channels showing overlay and alignment of motor nerve terminal and motor endplate. (C) Upper panel, FM143, middle, TRITC-α-bungarotoxin; lower, merged channels. Yellow fluorescent region indicates the alignment of presynaptic and post-synaptic components of the NMJ.

6. Exchange the bathing fluid with normal physiological saline frequently during the dissection. With the ribcage pinned external face uppermost, carefully cut through the intercostal muscles with watchmaker’s forceps and fine spring scissors, taking care not to damage the underlying triangularis sterni muscle. The dissection of triangularis sterni is best performed under a dissecting microscope that permits both transmitted and incident illumination simultaneously.

7. Snip the ribs at the sternum using small iris scissors or spring scissors, then reflect and remove the ribs, carefully trimming away adherent tissue and blood vessels. The triangularis sterni muscle should then be exposed as a semi-transparent sheet, only 1 to 3 muscle fibers thick (Fig. 1). Insert fine minutien pins through the lateral intercostal musculature and the sternum to secure the muscle, stretched at about its resting length. 8. Take 10 ml depolarizing saline and add 25 μl of either FM1-43 or FM1-43FX (or AM1-43) from an aqueous 1 mg/ml stock solution. To this solution add 20 μl

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from an aqueous 500 μg/ml stock solution of TRITC-α-bungarotoxin. The solution containing both fluorescent dyes will be referred to as staining saline. Both FM-dyes and labeled bungarotoxin can be divided into aliquots, frozen, and used after thawing. They may be stored in a refrigerator for a few days without loss of activity. An Alexa-Fluor 555 conjugate of α-bungarotoxin may be used if the preparation is intended for confocal microscopy although TRITC-α-bungarotoxin is suitable for confocal microscopy as well.

9. Drain the dissecting dish of all normal physiological saline and replace it with 5 to 10 ml staining saline containing the fluorescent cocktail. Incubate in this solution for 10 to 30 min on a rocking platform at room temperature. After the incubation, wash three to five times, each time with copious amounts of oxygenated normal saline for 30 to 60 min each time, and for the final wash immerse the dish containing the pinned preparation in 250 ml or more of oxygenated normal medium. You may need to adjust the time of the incubation depending on the nature of your preparation. Start with 10 min and increase the time, if necessary.

10. Transfer the preparation, still pinned to the Sylgard-lined dish and covered with fresh normal saline to an upright fluorescence compound microscope fitted with water-dipping 10× to 60× objectives. 11. Image and record the glowing pretzel-shape of the primary junctional folds at the motor endplates using the TRITC cube. Obtain images with a digital camera (e.g., a Hamamatsu Orca-12 camera, driven by Improvision/Perkin-Elmer OpenLab software, was used for the images presented in Fig. 1B-C). The broad emission spectrum of FM-dyes enables terminals to be visualized through standard FITC or TRITC filter cubes. However, better results are obtained using a custom filter cube containing a 435-nm excitation filter and a 515-nm, narrow bandpass (±10 nm) emission filter. A 495-nm dichroic mirror completes the custom cube.

12. If FM1-43fx (or AM1-43) is used instead of FM1-43, then fix the preparation for 30 min in 4% paraformaldehyde after the post-depolarization washes. However, the coloration of the nerve terminals may fade over the following 12 to 24 hr so images are best obtained as soon as possible after dye loading. 13. If the microscope is part of a confocal system, make a z-series at the diffraction limit and either combine these into a maximum-intensity projection or make rotating movies around x, y, or z axes. In good preparations, it is possible to discern the organization of the secondary junctional folds. ACh receptors are more concentrated at the crests of these folds. Quantification of end-plate size and fractional occupancy by motor nerve terminals can be accomplished using public-domain Image J software, downloadable from http://rsbweb.nih.gov/ij/.

REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps.

Depolarizing saline

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Freshly prepare 100 ml of a depolarizing mammalian physiological saline solution, the same composition as the normal physiological saline (see recipe) but with KCl elevated to 50 mM and NaCl reduced by 45 mM to 92 mM. Bubble to equilibration as with normal physiological saline. Do not store.

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FM1-43/FM1-43fx 1 mg/ml FM1-43/FM1-43FX (Invitrogen) in phosphate-buffered saline (PBS). Divide into 25- to 100-μl aliquots into microcentrifuge tubes and freeze. Thawed aliquots can be stored for a few days at 4◦ C without loss of activity.

Normal saline Freshly prepare one liter of normal physiological saline solution from stocks of NaCl (3 M), KCl (1 M), CaCl2 (0.2 M), MgCl2 (0.2 M), NaH2 PO4 (0.2 M). The following volumes of these, respectively, should be mixed and made up to 1 liter: 40 ml, 5 ml, 10 ml, 5 ml, and 2 ml. To this mix add 2 g NaHCO3 and 1 g Dglucose. After mixing, bubble through tubing or an aspirator with 95% O2 /5%CO2 gas mix for at least 10 min before use. Alternatively, instead of mixing NaH2 PO4 and NaHCO3 , add HEPES buffer to a final concentration of 5 mM, adjust pH to 7.2 to 7.4 and bubble either with air or 100 % O2 . Do not store.

TRITC-α-bungarotoxin 500 μg/ml TRITC-α-bungarotoxin (Invitrogen) in phosphate-buffered saline (PBS). Divide into 25- to 100-μl aliquots into microcentrifuge tubes and freeze. Thawed aliquots can be stored for a few days at 4◦ C without loss of activity.

COMMENTARY Background Information A potent high-affinity ligand for acetylcholine (ACh) receptors, known as αbungarotoxin, was originally extracted from the venom of a Taiwanese species of krait. The utilization of this toxin led to detailed molecular and biophysical characterization of these receptors at the mammalian NMJ and elsewhere (Chang and Lee, 1963; Changeux et al., 1970; Mishina et al., 1986). Conjugation of α-bungarotoxin with fluorescent ligands also enables simple, one-step labeling of ACh receptor-rich patches in the membranes of cultured myotubes, or motor endplates in situ (or strictly, the densely packed receptors located in the muscle fiber membranes at the electron dense crests of the junctional folds). Visualizing the distribution of ACh receptors in mammalian skeletal muscles is therefore a very simple procedure (Anderson and Cohen, 1977; Slater, 1982). Various fluorescent forms of αbungarotoxin are available: FITC-, TRITC-, or AlexaFluor variants all give good results. Good images can be obtained from any muscle but whole-mounts of thin muscles give the best overviews and images of endplate distribution and fine structure. Excellent preparations of this kind can be prepared using the triangularis sterni and the levator auris longus muscles, but fluorescent α-bungarotoxin also works well with tissue

sections or teased preparations of any muscle. Those we have tested and for which we have obtained uniformly good staining results include extensor digitorum longus, soleus, diaphragm, sternomastoid, intercostals, transverses abdominis, flexor digitorum brevis, interosseus, and lumbrical muscles. In fact, the lumbrical muscles are small and thin enough for excellent data to be obtained from whole-mounts using either conventional fluorescence of confocal microscopy. Classical silver-staining methods for motor axons and their terminals have the advantage of being high in contrast and durable: wellkept preparations can still be inspected and useful data gained years after the preparation has been made. However, the method is capricious and time consuming; and with the advent of confocal microscopy (useless with silverstained preparations) and other digital imaging technologies the need for storage of stably stained preparations has diminished somewhat. Other light microscope–based methods for staining and imaging motor nerve terminals have included zinc-iodide/osmiumtetroxide, and methylene blue. However, all these methods were essentially superseded by immunostains of one kind or another, either using a peroxidase-based method to stain NMJ, or using fluorescent secondary antibodies to highlight proteins localized at these synapses (Gillingwater et al., 2002; Ribchester

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et al., 2004). Immunofluorescence is particulary well-suited to high-resolution analysis of preparations using confocal microscopy. However, several steps (and days of preparation) are required for making a good immunofluorescent specimen, so methods that are more rapid and reliable offer the potential for a higher throughput. Lichtman and colleagues showed that fluorescent vital staining of NMJ produces high contrast, effective results in some species, such as snakes (Lichtman et al., 1985). The same group also found that a styryl dye, 4-Di-2-Asp, produces sufficiently highcontrast passive staining of motor nerve terminals, enabling longitudinal study of motor nerve terminals over days, weeks, or months in living mice (Lichtman et al., 1987; Magrassi et al., 1987; Balice-Gordon and Lichtman, 1990). More widespread utilization of vital staining of NMJ was adopted when Betz and colleagues reported the activity-dependent staining and “destaining” of motor nerve terminals in diverse species using the aminostyryl dye “FM1-43” (Betz and Bewick, 1992; Betz et al., 1992). Other related molecules (such as FM2-10 or FM4-64, or RH414 from which FM1-43 was originally synthesized) also stain motor nerve terminals, although the concentrations, loading conditions, and excitation/emission wavelengths differ (Ribchester et al., 1994; Barry and Ribchester, 1995). Thin or small muscles give the best results with vital staining of motor nerve terminals using aminostyryl dyes, such as FM1-43 or its fixable analog, FM1-43fx (both from Invitrogen). The amphipathic molecular structure of styryl dyes like FM1-43 causes them to insinuate into, but not penetrate, exposed membranes. Thus, during exocytosis when the inner membranes of synaptic vesicles become exposed to the extracellular fluid, FMdye molecules diffuse into the open vesicle through the diffusion pore and become trapped therein, following endocytosis. In this way, styryl dyes stain “recycling” synaptic vesicles. In addition, the fluorescence of membranebound FM-dyes is an order of magnitude greater than when the molecules are free in aqueous solution. However, in order to achieve labeling of adequate numbers of vesicles and therefore to produce sufficient fluorescence to observe in the microscope requires intense stimulation, generated either physiologically via long periods of high-frequency nerve stimulation, or by using elevated, depolarizing concentrations of potassium ions. The protocol

described here is based on potassium-induced labeling of the terminals. For activity-independent coloration of axons and motor nerve terminals, nothing presently surpasses the transgenic expression of variants of enhanced green fluorescent proteins (eGFP), under control of a modified thy1 promoter (Caroni, 1997; Feng et al., 2000; Lu et al., 2009). Combinatorial, transgenic expression of GFP variants ultimately led to the generation of “brainbow” mice, with spectacular, aesthetically beautiful, as well as scientifically compelling, unique coloration of neuronal cell bodies, axons, and their terminals (Livet et al., 2007; Ribchester, 2009). While the original “thy1.2-XFP” transgenic lines, especially the “YFP-16” and “YFP-H” lines, give the best results for imaging motor axons and/or their neuromuscular arbors (Keller-Peck et al., 2001; Lu et al., 2009), it is impractical, for many purposes of routine characterization of NMJ structure, to crossbreed one of these YFP lines with another mouse line of interest, then nurture the offspring before appraising the neuromuscular phenotype morphologically (Wong et al., 2009). The utility of these fluorescent proteins combined with minimally invasive live imaging using confocal microendoscopy is described elsewhere in this volume (Ribchester, 2012).

Troubleshooting Presynaptic labeling with FM1-43 has not worked It is important to appreciate that the staining of nerve terminals depends on an adequate rate of recycling of synaptic vesicles in a living preparation. Thus, the method will not work if the preparation has been damaged or left standing for several hours, or if there are presynaptic neurotoxins present, or if the preparation has been fixed, or if the saline used to bathe the preparation is only a basic phosphate-buffered saline. A physiological saline with normal extracellular Ca concentration (at least 2 mM) is essential. In order to stimulate vesicle recycling and labeling, depolarization of the living terminals is essential. Check that the staining saline containing FM1-43 has been prepared with depolarizing saline and not normal saline. When exchanging the normal saline with staining saline, ensure that the dish containing the pinned muscle is properly drained. Try a longer incubation (30 to 40 min) period if the problem persists.

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Adequate washing is required to rinse passively bound FM1-43 from plasma membranes of the muscle fibers and connective tissue in order to provide adequate contrast. Washing of the preparation must be done with normal saline. If depolarizing saline is used for washing, then the labeled terminals will destain as a result of the maintained exocytosis in the absence of extracellular dye. The muscle is saturated with green fluorescence in the muscle fibers If substantial numbers of muscle fibers are damaged during the dissection, FM1-3 will enter the muscle fibers and label the extensive sarcoplasmic reticulum and other intracellular membranes, producing incandescent fluorescence that will most likely eclipse the fine detail of motor nerve terminal fluorescence. This internal fluorescence is frequently associated with “contraction clots” in the muscle fibers. This problem can only be avoided or averted by improved dissection technique. In a good preparation, all the muscle fibers adopt the appearance of smooth cylinders along their entire length (Fig. 1A). α-Bungarotoxin staining has not worked The method is extremely robust because αbungarotoxin binds to ACh receptors with very high affinity, so the only plausible reasons for failure of the stain are that a bad lot has been purchased (record the lot number of effective batches), or that some mistake has been made in making up or diluting the stock solution. Penetration of the labeled protein may limit staining to superficial muscle fibers in some muscles, but every endplate should become labeled swiftly in triangularis sterni muscle if the protocol here has been followed properly. Another possibility for failure is some pathological feature of the muscle/mouse that reduces the number or density of ACh receptors at endplates.

Anticipated Results Figure 1 shows a typical example of freshly dissected living preparation of triangularis sterni, together with fluorescence micrographs of intramuscular axons and motor nerve terminals stained with FM1-43 and postsynaptic ACh receptors stained with TRITCα-bungarotoxin. Note that in addition to active, depolarization-induced homogeneous or punctate staining of the motor nerve terminals by FM1-43, ghostly outlines of the intramuscular nerves are also visible. This is mainly passive staining due to the insinuation of the

FM1-43 dye molecules into the membranes of the myelin sheaths. Note also the almost perfect alignment between the motor nerve terminal arbor and post-synaptic ACh receptors.

Time Considerations

Preparing the solutions takes ∼15 min; dissection of triangularis sterni takes, with practice, about 15 to 30 min. Results can therefore readily be obtained within either a morning or an afternoon’s work.

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