Mouse Models of Human Cancer / Edition 1 available in Hardcover
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Mice have become the species of choice for modeling the complex interactions between tumor cells and the host environment. Mouse genetics are easily manipulated, and a growing array of technology exists for this purpose. Mouse models allow investigators to better understand causal relationships between specific genetic alterations and tumors, utilize new imaging techniques, and test novel therapies. Recent developments along these lines show great promise for the development of new anti-cancer treatments.
Mouse Models of Human Cancer provides researchers and students with a complete resource on the subject, systematically presenting the principles, methodologies, applications, and challenges associated with this exciting field. Offering a survey of the latest research and a description of future areas of interest, this text:
- Presents real experimental data
- Describes organ site-specific mouse models
- Clearly identifies suitable models for further drug testing
- Critically analyzes current methodologies and their limitations
- Features numerous recognizable expert contributors
- Lists key Web sites, reagents, and companies
From mouse handling and genetic engineering to preclinical trials, Mouse Models of Human Cancer is a comprehensive guide to using these models and relating them to human disease. Its uniform presentation describes organ-specific models in clinical, imaging, and molecular terms, and lays out the relevant genetics, experimental approaches, histological comparisons with human disease, and conclusions.
Combining stellar chapter authors, rich illustrations, and clear, up-to-date coverage, Mouse Models of Human Cancer is an invaluable resource for advanced students and cutting-edge researchers.
|Product dimensions:||8.75(w) x 11.30(h) x 1.15(d)|
About the Author
Eric Holland received his Ph.D. from the University of Chicago in 1985 and M.D. from Stanford University in 1990. He completed his residency in neurosurgery at UCLA and did postdoctoral work at Stanford and the NIH. He holds appointments at Memorial Sloan-Kettering Cancer Center and Cornell University. He is a clinically active neurosurgeon and heads a laboratory at the Sloan-Kettering Institute focused primarily on understanding the molecular mechanisms underlying the pathogenesis of CNS tumors and in modeling these cancers in the mouse. Dr. Holland is the recipient of the Searle Scholars Award, the American Brain Tumor Association Research Award, the Peter Steck Memorial Award, the Bressler Scholars Award and the Seroussi Award. He has served on the NIH Brain Tumor Process Review Group, is a member of the Mouse Models of Human Cancer Consortium and is the Principal Investigator of the Brain Tumor SPORE program at MSKCC
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Mouse Models of Cancer
John Wiley & SonsCopyright © 2004 John Wiley & Sons, Inc.
All right reserved.
Chapter OneAPPROACHES TO HANDLING, BREEDING, STRAIN PRESERVATION, GENOTYPING, AND DRUG ADMINISTRATION FOR MOUSE MODELS OF CANCER
Dawnalyn Boggess, Kathleen A. Silva, Carlisle P. Landel, Larry Mobraaten, and John P. Sundberg
The Jackson Laboratory, Bar Harbor, Maine
Mouse models are extremely valuable biomedical tools for cancer research. Their value to individual investigators depends upon the investigator's ability to manage a colony and manipulate the mice. Basic genotyping to maintain a colony of genetically engineered mice is a critical technique. Understanding the differences between inbred and outbred (nonsegregating vs. segregating backgrounds) determines the reproducibility of models. Preserving valuable colonies for future use through cryopreservation of embryos, sperm, or just the constructs used to create the models enables future work to be done. This chapter provides a broad overview of many of the methods commonly used to perform these tasks as well as how and where to find more specific information.
Inbred laboratory mice have long been the species of choice for biomedical research because (1) of the relatively low cost for maintaining a colony; (2) with inbreeding, mice become genetically identical to each other; (3) numerous reagents are available for veryspecialized testing; (4) there is a high degree of genetic homology between mice and humans; and (5) there are many sophisticated genetic tools available for working with mice. Mutations in laboratory mice do occur spontaneously. This may be due to inbreeding, where two alleles of a deleterious recessive allele become manifest. Alternatively, insertional mutagenesis may occur when a retrovirus integrates within the mouse genome. Genetic engineering has become technically accessible to many laboratories where overexpression (transgenesis), under-or nonexpression (targeted mutagenesis or the so-called knockout approaches), or selective temporal expression (so-called gene switch methods) of specific, known genes can now be done. Random point mutagenesis has come back into vogue using ethylnitrosourea (ENU) or ethylmethanesulfonate (EMS). All of these approaches provide an array of potential or real models for the study of many types of diseases, including cancer (Nakamura et al., 2002a, b).
Maintaining and working with all of these types of mice are challenging, even for those scientists familiar with conventional inbred strains. Detailed texts are available on systematic workup and characterization methods for mutant mice (Sundberg and Boggess, 2000), general background diseases, including cancer, in commonly used strains (Maronpot et al., 1999; Mikaelian et al., 2004; Mohr et al., 1996), or details on specific organ systems (Smith et al., 2002; Sundberg, 1994b; Ward et al., 2000) which are beyond the scope of this book. Much of this information, including color images, is coming on-line through sources such as the Mouse Tumor Biology Database (tumor.informatics.jax.org/FMPro?-db =TumorInstance&-format=mtdp.html&-view), Mouse-PhenomeDatabase (aretha.jax.org/pubcgi/phenome/ mpdcgi?rtn=docs/home), and European Mouse Pathology Database (pathbase.net). Details on mouse genetics and gene mapping (Silver, 1995; Sundberg et al., 1998) and access to mouse strains and mutations are readily available through the Mouse Genome Informatics Database (informatics.jax.org). These and many other resources are listed in Table 1.1.
This chapter provides an overview of methodologies commonly used to maintain, manipulate, and preserve genetic-based mouse models of human diseases, especially cancer models.
METHODS FOR HANDLING LABORATORY MICE
Each institution has its own regulations dealing with maintaining a rodent colony, which includes various types of housing, bedding, feed, and water. Regardless, it is critical to optimize these to maintain a pathogenfree colony. This includes autoclaving feed, acidifying or treating water to account for specific issues in each locality, and avoiding bedding that releases aromatic hydrocarbons that might complicate cancer studies. It is beyond the scope of this chapter to describe all the variations, but these are described elsewhere (Clark et al., 1996). Maintaining a pathogen-free colony, or at least a specific pathogen-free colony whereby the pathogen status is known for each colony, is critical as well. Controversy over complications of phenotypes due to pathogen status of colonies has dictated that careful and continual microbiological monitoring be incorporated into colony maintenance protocols. Again, it is beyond the scope of this chapter to describe disease-monitoring selection and detection methods, but these are available (Lindsey et al., 1991a, b). This section will summarize some commonly used handling methods for mice to identify individuals as well as to collect bodily fluids and excretions.
Methods for individual identification of mice vary between institutions and laboratories. Some of the commonly used methods are summarized below (Seymour et al., 2004). All manipulations of mice require prior approval of the Institutional Animal Care and Use Committee (IACUC). Developing a good working relationship with the institutional attending veterinarian will be a great help in planning experiments, preparing documentation for IACUC protocol approval, and obtaining training in many of these procedures to optimize results.
Identifying Individual Mice
There are a number of methods that can be used for identification of individual mice in a colony or study. These range from ear notching using a small punch (National Band and TAg Company, Newport, KY) to making a hole or notch in the mouse's ear (Figs. 1.1 and 1.2), ear tags (National Band and TAg Company) attached to the ear, digit amputation using scissors (Fig. 1.3), foot (Fig. 1.4) or tail tattooing using various tattooing machines (Animal Identification and Marking Systems, AIMS Inc., Ottawa, Canada), and implanting electronic chips under the skin of the mice (Bio Medic Data Systems, Inc., Seaford, DE, or American Veterinary Identification Devices, Norco, CA). Wounding mice can result in abnormal healing such as keloids or development of specific types of cancer in some models. Therefore, attention to methods that might complicate procedures is important.
Collecting Blood and Body Fluids
Blood is collected by retro-orbital bleeding, tail tip amputation, cardiac puncture, or decapitation. The reasons to use each method varies with age, purpose of the study, volume needed, and methods approved at each institution. Methods are summarized below as modified from Seymour et al. (2004).
Retro-Orbital Bleeding This method is a valuable, nonterminal method for blood collection to obtain small volumes of blood. Retro-orbital bleeding is performed using manual restraint. Mice are grasped with thumb and index finger behind the ears placing the tail secured between the handlers' last finger and palm to immobilize them. A variety of anesthetics may be required by the IACUC at some institutions to perform this procedure.
The tip of an hematocrit tube is inserted into the medial canthus of the eye at a slight angle. Using a twisting motion with slight pressure on the tube, the retro-orbital venous plexus is entered. Once the blood begins flowing, the hematocrit tube should be withdrawn slightly to allow collection by capillary action. The hematocrit tube may be heparinized, contain ethylenediaminetetraacetic acid (EDTA), or other anticoagulants for plasma collection or contain no anticoagulant for serum collection.
Retro-orbital bleeding can also be used to collect larger volumes of blood if the mouse is going to be euthanized for necropsy. The mouse is anesthetized; then blood is collected in the same manner as for smaller volumes described above using heparinized hematocrit tubes. It is possible to fill 10-12 tubes using this method depending upon the size, age, and health status of the mouse. Each tube holds approximately 75 µL of blood. These tubes can be expressed into the appropriate collection tubes for the types of testing being done (see section on blood handling). This method decreases the variability in test results that may be caused by stress the mouse suffers during other methods of blood collection. This method also allows for more uniformity in sample volume when working with many mice.
Tail Bleeding Some technicians hold the mouse as described for the eye bleed for a manual tail bleed procedure. However, tail bleeding is often done when the mouse is placed in a restraint device with the tail protruding with an Eppendorf or glass tube for blood collection positioned below it (Fig. 1.5). Care must be taken not to contaminate blood with fecal matter or urine when using the tail-bleeding method. A tail bleed is accomplished by severing the ventral coccygeal vein on the ventral side of the base of the tail with a razor blade. Warming the mouse or its tail prior to bleeding will increase the blood flow.
Cardiac Puncture Cardiac puncture is a clean and simple way to get as much blood as possible from a mouse at the time of necropsy. It must be performed immediately after the mouse is euthanized. Many people perform cardiac puncture without opening up the thorax by inserting a 25- to 27-gauge needle through the skin and intercostal muscles and into the heart, then withdrawing blood into the syringe (a 1-mL syringe is large enough for most mice). This approach requires some technical skill but often results in sufficient amounts of blood being obtained. An alternative approach is to open the thoracic cavity immediately after euthanizing the mouse. This will expose the heart, making it easy to correctly insert a needle, obtaining a cleaner sample of larger volume. This is achieved by following normal necropsy procedures detailed elsewhere (Relyea et al., 2000; Seymour et al., 2004), being careful not to sever major blood vessels. Once the abdominal muscles have been incised, a pair of sharp/sharp scissors is used to cut the ribs in a triangular shape, starting equidistant from the xiphoid process on both sides and angling toward the clavicle. The sternum should not be severed since this may rupture major blood vessels near the thoracic inlet, causing rapid and excessive blood loss. The ribs should be retracted to reveal the heart and lungs. Using this approach, the right ventricle can be visualized and cleanly punctured. Blood is withdrawn slowly into the syringe. Gentle pressure applied to the visceral organs, working toward the heart, pushes abdominal blood into the right ventricle to optimize collection. Blood should be promptly and gently expressed into an appropriate collection tube before clotting occurs. The syringe plunger should be depressed slowly since high-velocity ejection may damage red blood cells, resulting in hemolysis.
Decapitation This method is seldom approved by animal-care-and-use committees unless there is sound scientific justification. This is the method of choice for some endocrine assays when hormones may be released during handling, altering results, or for mice less than 10 days old. A number of commercial guillotine-type instruments are available (Harvard Apparatus Inc., Holliston, MA). Alternatively, mice can have their heads disarticulated with a large pair of scissors (6.5-7 in., blunt/sharp). The mouse is firmly grasped by the skin at the back of the neck and, holding it over a blood collection tube, the head is swiftly and cleanly severed.
For serum collection, blood should be collected into small-volume serum separator tubes (Microtainer Brand, stock number 365956, Becton Dickinson, Franklin, NJ). These can then be held at room temperature for 15 min to 1 h and then centrifuged. The serum should be decanted and stored in plastic tubes (Eppendorf tubes, Brinkmann Instruments, Inc., Westbury, NY; Nunc Tubes, Nalge Nunc International, Fisher Scientific, Pittsburgh, PA) for storage and frozen at -80°C until used. Plasma is obtained by collecting blood in tubes containing EDTA or heparin to prevent clotting. The blood is then centrifuged, the plasma is decanted into tubes, and it is frozen for future use. Whole blood should be handled according to the instructions provided by the diagnostic laboratory that will be doing the blood chemistry workup.
Feces are collected at the time of necropsy and can be frozen in plastic tubes for a variety of assays. Most mice defecate upon handling so a few fresh samples can be obtained from a defined individual. This is a simple resource for Helicobacter spp. surveillance using polymerase chain reaction (PCR) methods (Mahler et al., 1998) or for fecal IgA quantification (Bristol et al., 2000).
Urine is usually expelled when a mouse is handled. A person can pick up the mouse as described for the retro-orbital bleeding procedure, holding the tail back with the little finger. In this position most mice micturate a few drops of urine. If needed, the mouse can be manually restrained and the caudal abdomen gently massaged to induce urination. Urine can be collected in a clean plastic tube or tests done directly with a variety of chemically impregnated strips. Chemstrip (Boehringer Mannheim Diagnostics, Indianapolis, IN) and Ames Multistix (Miles Inc. Diagnostic Division, Tarrytown, NY) are two urine analysis reagent strips that test for numerous urine components, including glucose, ketones, protein content, and pH. For more specific tests that require larger volumes of urine or urine collected over defined intervals, metabolic cages are commercially available (Columbus Instruments, Columbus, OH). Urine specific gravity is measured using a hand-held refractometer. Some companies offer refractometers especially made for urine testing, such as the Fisherbrand UriSystem Refractometer (Fisher, Pittsburgh, PA).
GENERAL BREEDING STRATEGIES FOR MAINTAINING COLONIES OF GENETICALLY ENGINEERED MICE
When establishing a colony of potentially mutant mice, it is important to examine all mice as soon after birth as possible and at regular intervals (at least every other day during the first 2 weeks) thereafter until the phenotype (clinical features) are well defined. Defects may be observable at birth, as is seen in flaky skin (sn) mice, which have a mild anemia at birth, that make them easy to identify (Sundberg et al., 1997b) or juvenile alopecia (jal) mice, which have easily identifiable abnormalities of the vibrissae as early as 2-3 days of age (McElwee et al., 1999). Other defects may develop as the mouse ages, as in harlequin ichthyosis (ichq), in which mice are normal until 5 days of age, when they develop thick scaling skin, then die by 10-12 days of age (Sundberg et al.,
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Table of Contents
PART I: MOUSE HANDLING AND ENGINEERING.
1. Approaches to Handling, Breeding, Strain Preservation, Genotyping, and Drug Administration for Mouse Models of Cancer (Dawnalyn Boggess, Kathleen A. Silva, Carlisle P. Landel, Larry Mobraaten, and John P. Sundberg).
2. Pathology of Mouse Models of Cancer (Robert D. Cardiff).
3. Intercurrent Infections in Genetically Engineered Mice (Stephen W. Barthold).
4. Germline Modification Strategies (Silvia Grisendi and Pier Paolo Pandolfi).
5. Somatic Cell Gene Transfer (Lene Uhrbom and Eric C. Holland).
PART II: ORGAN SITE–SPECIFIC MOUSE MODELS OF HUMAN CANCER.
6. Lung Cancer (Jonathan M. Kurie, Parviz Minoo, Stephen S. Hecht, Alexander Nikitin, and Franco J. DeMayo).
7. Mammary Gland Cancer (Dalit Barkan, Cristina Montagna, Thomas Ried, and Jeffrey E. Green).
8. Prostate Cancer (Katharine Ellwood-Yen and Charles Sawyers).
9. Skin Cancer (Marcus Bosenberg).
10. Ovarian Cancer (Sandra Orsulic).
11. Peripheral Nervous System Tumors (David H. Gutmann, Arie Perry, Reshma Rangwala, and Larry S. Sherman).
12. Central Nervous System Tumors (Andrew B. Lassman and Eric C. Holland).
13. Myeloid Malignancies (Scott C. Kogan).
14. Lymphoid Malignancies (Michael Teitell and Pier Paolo Pandolfi).
PART III: GENERAL ISSUES IN CANCER BIOLOGY.
15. Genetic Modifiers (David W. Threadgill, Kent W. Hunter, Fei Zou, and Kenneth F. Manly).
16. Modeling Blood Vessel Formation in the Mouse (Robert Benezra, Shahin Rafii, and David Lyden).
17. Metastasis (Dawn S. Chandler and Guillermina Lozano).
18. Immunologic Study of Tumors in Mouse Models (Taha Merghoub and Alan N. Houghton).
PART IV: IMAGING TECHNOLOGIES.
19. Micro–Computed Tomography of Mouse Cancer Models (Jamey P. Weichert).
20 .Magnetic Resonance Imaging in Mouse Cancer Models (Manickam Muruganandham and Jason A. Koutcher).
21. Bioluminescent Imaging of Mouse Cancer Models (Christopher H. Contag).
PART V: PRECLINICAL TRIALS.
22 .Use of Magnetic Resonance Imaging for Evaluation of Treatment Response (Brian D. Ross, Thomas L. Chenevert, Bradford A. Moffat, Alnawaz Rehemtulla, Daniel E. Hall, Patrick McConville, and Jonathan Moody).
23. Physiologically Based Pharmacokinetic and Pharmacodynamic Modeling (Raymond S. H. Yang, James E. Dennison, Melvin E. Andersen, Ying C. Ou, Kai H. Liao, and Brad Reisfeld).
24. Trial Design and Biostatistics (Michael F. W. Festing).
25. Cancer Drug Development in the Modern Era (William R. Sellers and Alex Matter).
26. Preclinical Trials in Mouse Cancer Models (Brian Weiss and Kevin Shannon).