![© 2002 Humana Press Inc.
999 Riverview Drive, Suite 208
Totowa, New Jersey 07512
www.humanapress.com
All rights reserved. No part of this book may be reproduced, stored in a retrieval system, or transmitted in
any form or by any means, electronic, mechanical, photocopying, microfilming, recording, or otherwise
without written permission from the Publisher.
The content and opinions expressed in this book are the sole work of the authors and editors, who have
warranted due diligence in the creation and issuance of their work. The publisher, editors, and authors are
not responsible for errors or omissions or for any consequences arising from the information or opinions
presented in this book and make no warranty, express or implied, with respect to its contents.
Due diligence has been taken by the publishers, editors, and authors of this book to assure the accuracy of
the information published and to describe generally accepted practices. The contributors herein have care-
fully checked to ensure that the drug selections and dosages set forth in this text are accurate and in accord
with the standards accepted at the time of publication. Notwithstanding, since new research, changes in
government regulations, and knowledge from clinical experience relating to drug therapy and drug reactions
constantly occur, the reader is advised to check the product information provided by the manufacturer of each
drug for any change in dosages or for additional warnings and contraindications. This is of utmost importance
when the recommended drug herein is a new or infrequently used drug. It is the responsibility of the treating
physician to determine dosages and treatment strategies for individual patients. Further, it is the responsi-
bility of the health care provider to ascertain the Food and Drug Administration status of each drug or device
used in their clinical practice. The publishers, editors, and authors are not responsible for errors or omissions
or for any consequences from the application of the information presented in this book and make no warranty,
express or implied, with respect to the contents in this publication.
This publication is printed on acid-free paper. '
ANSI Z39.48-1984 (American National Standards Institute) Permanence of Paper for Printed Library
Materials.
Cover design by Patricia F. Cleary.
Cover illustration: Oocyte injection in the mouse—embryo following injection of DNA (see pp. 63–64.)
For additional copies, pricing for bulk purchases, and/or information about other Humana titles,
contact Humana at the above address or at any of the following numbers: Tel: 973-256-1699;
Fax: 973-256-8341; E-mail: humana@humanapr.com or visit our website at http://humanapress.com
Photocopy Authorization Policy:
Authorization to photocopy items for internal or personal use, or the internal or personal use of specific
clients, is granted by Humana Press Inc., provided that the base fee of US $10.00 per copy, plus US $00.25
per page, is paid directly to the Copyright Clearance Center at 222 Rosewood Drive, Danvers, MA 01923.
For those organizations that have been granted a photocopy license from the CCC, a separate system of
payment has been arranged and is acceptable to Humana Press Inc. The fee code for users of the Transactional
Reporting Service is: [0-89603-696-0/02 $10.00 + $00.25].
Printed in the United States of America. 10 9 8 7 6 5 4 3 2 1
Library of Congress Cataloging-in-Publication Data
Transgenesis techniques: principles and protocols/edited by Alan R. Clarke.—2nd ed.
p. cm. — (Methods in molecular biology; v. 180)
ISBN 0-89603-696-0 (alk. paper)
1. Transgenic animals—Laboratory manual. 2. Animal genetic engineering—Laboratory manuals.
I. Clarke, Alan R. II. Methods in molecular biology (Totowa, NJ); v. 180.
QH442.6.T66 2002
576.5'07'24--dc21
2001024458](https://image.slidesharecdn.com/2142687-250526093034-92509160/85/Transgenesis-Techniques-Principles-And-Protocols-Jim-Mcwhir-Auth-8-320.jpg)
![6 McWhir
4. A Hierarchy of Complexity
Some transgenic applications in farm animals will involve complex target-
ing technology and yet in biologic terms may have quite humble goals (simple
loss of function mutation). In other instances, the technology may be relatively
crude (as with pronuclear injection of growth hormone [GH] genes) whereas
the biologic objective (to modify growth rate) is highly ambitious. In mouse
transgenic programs, the biologic objective is usually straightforward—to
observe and record the effects of ectopic gene expression or, in the case of
gene targeting, the effects of loss of gene function. Here, although the pheno-
typic consequences are often not predictable, the experiment will always be
informative.
In contrast to the mouse, many potential livestock applications (particu-
larly in agriculture) will involve intervention in complex metabolic pathways
in which the objective is to achieve a predetermined phenotypic change. Here,
there is an additional challenge: it is necessary to accurately predict the phe-
notypic consequences of a single genetic modification. Limited attempts to do
this in order to increase the growth rate of transgenic pigs have led to unfore-
seen consequences on animal health and fertility (31). The most straightfor-
ward applications of transgenesis in livestock are those in which the objective
is simply to harvest high levels of recombinant protein. In this case, there is no
requirement to modify endogenous metabolic pathways, a physiologic response
to transgene expression is neither required nor anticipated, and the risks of
adverse effects on animal health and welfare are minimized. It is not surpris-
ing, therefore, that the most successful transgenic applications to date have
involved expression of human therapeutic proteins in the milk of transgenic
sheep (32), pigs (33), cows (34), and goats (35).
5. Biopharming
Biopharming is the commercial production of pharmaceuticals from the
body fluids of transgenic animals. Although most attention has centered on
the mammary gland (for reviews see refs. 36–39), other body fluids may have
particular benefits for certain applications. For example, human GH has been
expressed in mouse bladder epithelium under the control of the mouse
uroplakin promoter (40). Advantages of bladder production might include the
ability to harvest from all animals at all stages of their lives and the small
number of other proteins from which the recombinant protein need to be puri-
fied. Transgenic swine have been generated that express human hemoglobin in
their blood as a potential cell-free substitute for human plasma (41,42). How-
ever, this blood-based approach has been hampered by difficulty in separating
human hemoglobin from its porcine counterpart.](https://image.slidesharecdn.com/2142687-250526093034-92509160/85/Transgenesis-Techniques-Principles-And-Protocols-Jim-Mcwhir-Auth-19-320.jpg)
![Gene Transfer in Drosophila 29
1. Leitz micromanipulator.
2. Nikon inverted phase-contrast microscope.
3. Vibration-free table, on which the microscope is mounted.
4. Loaded needle, containing the DNA to be injected.
5. Collar (Narishige, Tokyo) into which the needle is placed, which, in turn, is
attached to the micromanipulator.
Although the micromanipulator is used to position the needle, injection is
carried out by moving the microscope stage with the embryos on it. We use
an air-filled system to deliver the DNA into the embryos. This consists of a
60-mL glass syringe attached to the collar by a piece of rubber tubing
(Narishige Teflon™ tubing also may be used).
This system may appear very basic, but we find that the syringe imparts
adequate control of DNA delivery without producing the problems often
encountered when using a fluid-filled transmission system, and the system has
the advantage of being much cheaper. Injection needles are prepared from boro-
silicate capillaries (e.g., Clark Electromedical [Reading, UK] GC100TF-15
capillaries, which contain an internal filament) using a pipet puller. A rela-
tively inexpensive two-stage vertical needle puller can be used, such as the
PB-7 model from Narishige.
2.2. Fly Requirements
In general, a large number of embryos (in the region of 500–1000) need to
be injected for each construct in order to produce several independent
transformants. In our hands, between 25 and 75% of injected embryos will
hatch as larvae. Approximately 50% of the larvae will survive as adults, and
between 50 and 80% of the surviving adults will be fertile. Each surviving
adult will be individually mated, and approx 200 progeny from each mating
will be scored for the marker present on the transposon construct. Although the
frequency with which germline transformants are produced varies depending
on the construct injected (4), in general, on the order of 10% of the surviving
adults will produce at least one germline transformant among its progeny.
Therefore, it is reasonable to aim at obtaining about 100 adult survivors for
any given construct injected. We usually collect only one transformant
from the progeny derived from each surviving adult with which to estab-
lish stocks. This ensures that different transformants originated from inde-
pendent events.
Since the injections must be performed prior to pole cell formation, 1-h embryo
collections are used (see Subheading 3.3.). Therefore, the fly strain used for
embryo collections must be robust enough to provide sufficient eggs (at least
100) during a 1-h interval. One further consideration is that the presence of
defective P elements in the injected host strain can affect the frequency of](https://image.slidesharecdn.com/2142687-250526093034-92509160/85/Transgenesis-Techniques-Principles-And-Protocols-Jim-Mcwhir-Auth-40-320.jpg)
Transgenesis Techniques Principles And Protocols Jim Mcwhir Auth
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- 6. M E T H O D S I N M O L E C U L A R B I O L O G YTM John M. Walker, SERIES EDITOR 204. Molecular Cytogenetics: Methods and Protocols, edited by Yao-Shan Fan, 2002 203. In Situ Detection of DNA Damage: Methods and Protocols, edited by Vladimir V. Didenko, 2002 202. Thyroid Hormone Receptors: Methods and Protocols, edited by Aria Baniahmad, 2002 201. Combinatorial Library Methods and Protocols, edited by Lisa B. English, 2002 200. DNA Methylation Protocols, edited by Ken I. Mills and Bernie H, Ramsahoye, 2002 199. Liposome Methods and Protocols, edited by Subhash C. Basu and Manju Basu, 2002 198. Neural Stem Cells: Methods and Protocols, edited by Tanja Zigova, Juan R. Sanchez-Ramos, and Paul R. Sanberg, 2002 197. Mitochondrial DNA: Methods and Protocols, edited by William C. Copeland, 2002 196. Oxidants and Antioxidants: Ultrastructural and Molecular Biology Protocols, edited by Donald Armstrong, 2002 195. Quantitative Trait Loci: Methods and Protocols, edited by Nicola J. Camp and Angela Cox, 2002 194. Post-translational Modification Reactions, edited by Christoph Kannicht, 2002 193. RT-PCR Protocols, edited by Joseph O’Connell, 2002 192. PCR Cloning Protocols, 2nd ed., edited by Bing-Yuan Chen and Harry W. Janes, 2002 191. Telomeres and Telomerase: Methods and Protocols, edited by John A. Double and Michael J. Thompson, 2002 190. High Throughput Screening: Methods and Protocols, edited by William P. Janzen, 2002 189. GTPase Protocols: The RAS Superfamily, edited by Edward J. Manser and Thomas Leung, 2002 188. Epithelial Cell Culture Protocols, edited by Clare Wise, 2002 187. PCR Mutation Detection Protocols, edited by Bimal D. M. Theophilus and Ralph Rapley, 2002 186. Oxidative Stress and Antioxidant Protocols, edited by Donald Armstrong, 2002 185. Embryonic Stem Cells: Methods and Protocols, edited by Kursad Turksen, 2002 184. Biostatistical Methods, edited by Stephen W. Looney, 2002 183. Green Fluorescent Protein: Applications and Protocols, edited by Barry W. Hicks, 2002 182. In Vitro Mutagenesis Protocols, 2nd ed., edited by Jeff Braman, 2002 181. Genomic Imprinting: Methods and Protocols, edited by Andrew Ward, 2002 180. Transgenesis Techniques, 2nd ed.: Principles and Protocols, edited by Alan R. Clarke, 2002 179. Gene Probes: Principles and Protocols, edited by Marilena Aquino de Muro and Ralph Rapley, 2002 178.`Antibody Phage Display: Methods and Protocols, edited by Philippa M. O’Brien and Robert Aitken, 2001 177. Two-Hybrid Systems: Methods and Protocols, edited by Paul N. MacDonald, 2001 176. Steroid Receptor Methods: Protocols and Assays, edited by Benjamin A. Lieberman, 2001 175. Genomics Protocols, edited by Michael P. Starkey and Ramnath Elaswarapu, 2001 174. Epstein-Barr Virus Protocols, edited by Joanna B. Wilson and Gerhard H. W. May, 2001 173. Calcium-Binding Protein Protocols, Volume 2: Methods and Techniques, edited by Hans J. Vogel, 2001 172. Calcium-Binding Protein Protocols, Volume 1: Reviews and Case Histories, edited by Hans J. Vogel, 2001 171. Proteoglycan Protocols, edited by Renato V. Iozzo, 2001 170. DNA Arrays: Methods and Protocols, edited by Jang B. Rampal, 2001 169. Neurotrophin Protocols, edited by Robert A. Rush, 2001 168. Protein Structure, Stability, and Folding, edited by Kenneth P. Murphy, 2001 167. DNA Sequencing Protocols, Second Edition, edited by Colin A. Graham and Alison J. M. Hill, 2001 166. Immunotoxin Methods and Protocols, edited by Walter A. Hall, 2001 165. SV40 Protocols, edited by Leda Raptis, 2001 164. Kinesin Protocols, edited by Isabelle Vernos, 2001 163. Capillary Electrophoresis of Nucleic Acids, Volume 2: Practical Applications of Capillary Ele`¤ ophoresis, edited by Keith R. Mitchelson and Jing Cheng, 2001 162. Capillary Electrophoresis of Nucleic Acids, Volume 1: Introduction to the Capillary Electrophoresis of Nucleic Acids, edited by Keith R. Mitchelson and Jing Cheng, 2001 161. CytoskeletonMethodsandProtocols,editedbyRayH.Gavin,2001 160. Nuclease Methods and Protocols, edited by Catherine H. Schein, 2001 159. Amino Acid Analysis Protocols, edited by Catherine Cooper, Nicole Packer, and Keith Williams, 2001 158. Gene Knockoout Protocols, edited by Martin J. Tymms and Ismail Kola, 2001 157. Mycotoxin Protocols, edited by Mary W. Trucksess and Albert E. Pohland, 2001 156. Antigen Processing and Presentation Protocols, edited by Joyce C. Solheim, 2001 155. Adipose Tissue Protocols, edited by Gérard Ailhaud, 2000 154. Connexin Methods and Protocols, edited by Roberto Bruzzone and Christian Giaume, 2001 153. Neuropeptide Y Protocols, edited by Ambikaipakan Balasubramaniam, 2000 152. DNA Repair Protocols: Prokaryotic Systems, edited by Patrick Vaughan, 2000 151. MatrixMetalloproteinaseProtocols, editedby IanM.Clark,2001 150. Complement Methods and Protocols, edited by B. Paul Mor- gan, 2000 149. The ELISA Guidebook, edited by John R. Crowther, 2000 148. DNA–Protein Interactions: Principles and Protocols (2nd ed.), edited by Tom Moss, 2001 147. Affinity Chromatography: Methods and Protocols, edited by Pascal Bailon, George K. Ehrlich, Wen-Jian Fung, and Wolfgang Berthold, 2000 146. Mass Spectrometry of Proteins and Peptides, edited by John R. Chapman, 2000.
- 7. Humana Press Totowa, New Jersey M E T H O D S I N M O L E C U L A R M E T H O D S I N M O L E C U L A R B I O L O G YTM Transgenesis Techniques Principles and Protocols SECOND EDITION Edited by Alan R. Clarke School of Biosciences, Cardiff University Cardiff, UK
- 8. © 2002 Humana Press Inc. 999 Riverview Drive, Suite 208 Totowa, New Jersey 07512 www.humanapress.com All rights reserved. No part of this book may be reproduced, stored in a retrieval system, or transmitted in any form or by any means, electronic, mechanical, photocopying, microfilming, recording, or otherwise without written permission from the Publisher. The content and opinions expressed in this book are the sole work of the authors and editors, who have warranted due diligence in the creation and issuance of their work. The publisher, editors, and authors are not responsible for errors or omissions or for any consequences arising from the information or opinions presented in this book and make no warranty, express or implied, with respect to its contents. Due diligence has been taken by the publishers, editors, and authors of this book to assure the accuracy of the information published and to describe generally accepted practices. The contributors herein have care- fully checked to ensure that the drug selections and dosages set forth in this text are accurate and in accord with the standards accepted at the time of publication. Notwithstanding, since new research, changes in government regulations, and knowledge from clinical experience relating to drug therapy and drug reactions constantly occur, the reader is advised to check the product information provided by the manufacturer of each drug for any change in dosages or for additional warnings and contraindications. This is of utmost importance when the recommended drug herein is a new or infrequently used drug. It is the responsibility of the treating physician to determine dosages and treatment strategies for individual patients. Further, it is the responsi- bility of the health care provider to ascertain the Food and Drug Administration status of each drug or device used in their clinical practice. The publishers, editors, and authors are not responsible for errors or omissions or for any consequences from the application of the information presented in this book and make no warranty, express or implied, with respect to the contents in this publication. This publication is printed on acid-free paper. ' ANSI Z39.48-1984 (American National Standards Institute) Permanence of Paper for Printed Library Materials. Cover design by Patricia F. Cleary. Cover illustration: Oocyte injection in the mouse—embryo following injection of DNA (see pp. 63–64.) For additional copies, pricing for bulk purchases, and/or information about other Humana titles, contact Humana at the above address or at any of the following numbers: Tel: 973-256-1699; Fax: 973-256-8341; E-mail: humana@humanapr.com or visit our website at http://humanapress.com Photocopy Authorization Policy: Authorization to photocopy items for internal or personal use, or the internal or personal use of specific clients, is granted by Humana Press Inc., provided that the base fee of US $10.00 per copy, plus US $00.25 per page, is paid directly to the Copyright Clearance Center at 222 Rosewood Drive, Danvers, MA 01923. For those organizations that have been granted a photocopy license from the CCC, a separate system of payment has been arranged and is acceptable to Humana Press Inc. The fee code for users of the Transactional Reporting Service is: [0-89603-696-0/02 $10.00 + $00.25]. Printed in the United States of America. 10 9 8 7 6 5 4 3 2 1 Library of Congress Cataloging-in-Publication Data Transgenesis techniques: principles and protocols/edited by Alan R. Clarke.—2nd ed. p. cm. — (Methods in molecular biology; v. 180) ISBN 0-89603-696-0 (alk. paper) 1. Transgenic animals—Laboratory manual. 2. Animal genetic engineering—Laboratory manuals. I. Clarke, Alan R. II. Methods in molecular biology (Totowa, NJ); v. 180. QH442.6.T66 2002 576.5'07'24--dc21 2001024458
- 9. Preface v The past decade has witnessed a spectacular explosion in both the develop- mentanduseoftransgenictechnologies.Notonlyhavethesebeenusedtoaidour fundamental understanding of biologic mechanisms, but they have also facili- tated the development of a range of disease models that are now truly beginning to impact upon our approach to human disease. Some of the most exciting model systems relate to neurodegenerative disease and cancer, where the availability of appropriate models is at last allowing radically new therapies to be developed and tested. This latter point is of particular significance given the current concerns of the wider public over both the use of animal models and the merits of using genetically modified organisms. Arguably, advances of the greatest significance have been made using mammalian systems—driven by the advent of embryonic stem-cell–based strategies and, more recently, by cloning through nuclear transfer. For this reason, this new edition of Transgenesis Techniques focuses much more heavily on manipulation of the mammalian genome, both in the general discussions and in the provision of specific protocols. Of all mammalian experimental systems, the laboratory mouse is probably the most widely used, a situation that almost certainly derives from the fact that it is genetically the most tractable. This second edition, therefore, devotes much space to methodologies required for the creation and maintenance of genetically modified murine strains. In addition to protocols for conventional pronuclear injection, chapters have been included covering alternative routes to the germline, by either retroviral or adenoviral infection. Extensive cover- age is also given to the generation, maintenance, and manipulation of embry- onic stem cell lineages, since this is now widely recognized as an indispensable approach to genotype–phenotypeanalysis.PartVcontainsprotocolstofacilitate gene targeting and so permit both constitutive and conditional gene targeting. The latter approach, reliant on either the Cre-lox or the Flp-frt system, is rapidly gaining favor as a method of choice for the analysis of null mutations because it solves the twin difficulties of embryonic lethality and developmental compen- sation—two problems that have hampered the analysis of simple “knock-out strains.” The proliferation of newly engineered murine strains has given rise to one problem within the field, namely, that of the long-term storage of lines for which
- 10. vi Preface there might be no immediate requirement. Within many laboratories, this is now far from a trivial problem, and, therefore, methodologies are included that detail the cryopreservation of both male and female germlines. Although the mouse is currently the most genetically tractable system, it is not without its limitations and clearly cannot deliver all appropriate experimental or commercial systems. Transgenic manipulation of the rat germline is now delivering valuable models across a range of fields, perhaps most notably in neurobiology and in the study of vascular diseases. This edition, therefore, also focuses on the generation, maintenance, and cryopreservation of rat transgenic lines. The mouse and the rat remain essentially laboratory models. However, perhaps the most radical change to occur within the field relates to our emerging ability to genetically engineer livestock. In particular, the advent of cloning as a viable technology has wide ramifications for the scientific and industrial communities as well as for the wider public. Protocols are given for the generation of transgenic sheep by nuclear transfer, and, furthermore, the potential implications and future directions of large animal transgenesis are discussed in some detail. Finally, this second edition carries a very detailed part relating to the basic analysis of transgenic organisms. Although many of the techniques included are widely used throughout molecular biology, those pertinent to transgenic analysis have been brought together to facilitate the rapid analysis of phenotype. Used in conjunction with the plethora of techniques relating to the generation and mainte- nance of transgenic strains, the contributors and I anticipate that this new edition of Transgenic Techniques will prove an invaluable asset to any laboratory either already engaged in transgenic manipulation or setting out along this route. Alan R. Clarke
- 11. Preface ................................................................................................. v Contributors ......................................................................................... ix PART I. TOPICAL REVIEWS IN TRANSGENESIS 1 Biomedical and Agricultural Applications of Animal Transgenesis Jim McWhir................................................................................... 3 PART II. TRANSGENESIS IN INVERTEBRATE AND LOWER VERTEBRATE SPECIES 2 Gene Transfer in Drosophila Mark J. O'Connor and William Chia ........................................ 27 PART III. TRANSGENESIS IN THE MOUSE: OOCYTE INJECTION 3 Oocyte Injection in the Mouse Gary A. J. Brown and Timothy J. Corbin ............................... 39 PART IV. ALTERNATIVE ROUTES TO THE GERMLINE 4 Adenoviral Infection Tohru Tsukui and Yutaka Toyoda ........................................... 73 5 Retroviral Infection Richard A. Bowen ...................................................................... 83 PART V. TRANSGENESIS IN THE MOUSE: THE ES CELL ROUTE 6 In Vitro Isolation of Murine Embryonic Stem Cells David Wells................................................................................. 93 7 Production of Chimeras Derived from Murine Embryonic Stem Cells David Wells............................................................................... 127 8 Gene Targeting Strategies David W. Melton ....................................................................... 151 9 Cre/loxP Recombination System and Gene Targeting Ralf Kühn and Raul M. Torres................................................ 175 vii Contents
- 12. viii Contents PART VI. CRYOPRESERVATION OF MOUSE LINES 10 Cryopreservation of Transgenic Mouse Lines Jillian M. Shaw and Naomi Nakagata.................................... 207 11 Ovarian Tissue Transplantation and Cryopreservation: Application to Maintenance and Recovery of Transgenic and Inbred Mouse Lines Jillian M. Shaw and A. O. Trounson...................................... 229 PART VII. TRANSGENESIS IN THE RAT 12 Transgenesis in the Rat Linda J. Mullins, Gillian Brooker, and John J. Mullins ...... 255 PART VIII. TRANSGENESIS IN DOMESTIC SPECIES 13 Generation of Transgenic Livestock by Pronuclear Injection A. John Clark............................................................................ 273 14 Transgenic Sheep from Cultured Cells Keith H. S. Campbell ............................................................... 289 PART IX. CHARACTERIZATION AND ANALYSIS OF TRANSGENIC STRAINS 15 Analysis of Transgenic Mice Stefan Selbert and Dominic Rannie ...................................... 305 Index ................................................................................................. 343
- 13. RICHARD A. BOWEN • Animal Reproduction and Biotechnology Laboratory, Colorado State University, Fort Collins, CO GILLIAN BROOKER • Molecular Physiology Laboratory, University of Edinburgh Medical School, Edinburgh, UK GARY A. J. BROWN • Transgenic Mouse Core Facility, Shands Cancer Center, University of Florida, Gainesville, FL KEITH H. S. CAMPBELL • Division of Animal Physiology, University of Nottingham, Nr. Loughborough, Leicestershire, UK WILLIAM CHIA • Institute of Molecular and Cell Biology, Singapore A. JOHN CLARK • Division of Gene Expression and Development, Roslin Institute, Roslin, Midlothian, UK TIMOTHY J. CORBIN • Amgen, Inc., Thousand Oaks, CA RALF KÜHN • Institute for Genetics, Cologne, Germany JIM MCWHIR • Division of Molecular Biology, Roslin Institute, Roslin, Midlothian, Scotland DAVID W. MELTON • Molecular Medicine Centre, University of Edinburgh, Edinburgh, UK JOHN J. MULLINS • Molecular Physiology Laboratory, University of Edinburgh Medical School, Edinburgh, UK LINDA J. MULLINS • Molecular Physiology Laboratory, University of Edinburgh Medical School, Edinburgh, UK NAOMI NAKAGATA • Division of Reproductive Engineering, Center for Animal Resources and Development (CARD), Kumamoto University, Kumamoto, Japan MARK J. O’CONNOR • Institute of Molecular and Cell Biology, Singapore DOMINIC RANNIE • Department of Pathology, University of Edinburgh Medical School, Edinburgh, UK STEFAN SELBERT • Mice and More GmbH and Co. KG, Hamburg, Germany JILLIAN M. SHAW • Monash Institute of Reproduction and Development, Monash University, Clayton, Victoria, Australia ix Contributors
- 14. x Contributors RAUL M. TORRES • Department of Immunology, University of Colorado Health Sciences Center; National Jewish Medical and Research Center, Denver, CO YUTAKA TOYODA • Department of Reproductive and Developmental Biology, Institute of Medical Science, University of Tokyo, Tokyo, Japan A. O. TROUNSON • Monash Institute of Reproduction and Development, Monash University, Clayton, Victoria, Australia TOHRU TSUKUI • Department of Reproductive and Developmental Biology, Institute of Medical Science, University of Tokyo, Tokyo, Japan DAVID WELLS • Reproductive Technologies Group, AgResearch, Ruakura Research Center, Hamilton, New Zealand
- 15. Applications of Animal Transgenesis 1 I TOPICAL REVIEWS IN TRANSGENESIS
- 16. Applications of Animal Transgenesis 3 3 From: Methods in Molecular Biology, vol. 180: Transgenesis Techniques, 2nd ed.: Principles and Protocols Edited by: A. R. Clarke © Humana Press Inc., Totowa, NJ 1 Biomedical and Agricultural Applications of Animal Transgenesis Jim McWhir 1. Introduction In 1980, Gordon et al. (1) showed that DNA injected into the pronuclei of single-cell embryos could be incorporated, expressed, and transmitted to the offspring of transgenic mice. Since then, pronuclear injection has become a widely used and invaluable tool for the study of mammalian gene function. The same technique has also been used to generate transgenic livestock (2); however, the proportion of injected and transferred embryos giving rise to transgenic animals is greatly reduced relative to mice (1 to 2% vs 10–25%). Two general disadvantages of pronuclear injection apply equally to all species: unpredictable effects of site of incorporation and transgene copy number on gene expression lead to a requirement for testing multiple lines to ensure appropriate transgene expression, and the technique is restricted to the addi- tion of genetic material. The disadvantages of pronuclear injection have been partially circumvented in mice with the development of an alternate route to transgenesis through murine embryonic stem (ES) cells (3,4). ES cell lines are isolated from undif- ferentiated cells of the early embryo and retain in culture their capacity to dif- ferentiate into the full range of embryonic tissues. Hence, ES cells can be genetically modified in vitro and returned to the early embryo, where they resume their normal program of development. This procedure leads to the gen- eration of chimeric animals whose tissues, including germ cells, are frequently derived from both host embryo and ES genotypes, and a proportion of chime- ras will transmit the ES-derived genetic modification to their offspring. Pre- cise genetic modification can be achieved in ES cells by taking advantage of homologous recombination to target single-copy transgenes to specific sites or
- 17. 4 McWhir to modify existing genes in situ. A major limitation of this technology is that, at present, germline-competent ES cells are available only in the mouse. As a consequence of the inefficiency of pronuclear injection in farm ani- mals, the absence of proven ES cells in these species, and the high cost of animal maintenance, the literature describing transgenic livestock has been better served by reviews than by concrete example. Perhaps the single excep- tion has been the use of transgenic livestock to produce a small number of pharmaceutical proteins. This situation may be about to change; the develop- ment of techniques for cloning livestock from cultured cells (5,6), of cell- based transgenesis in sheep (7), and the imminent possibility of gene targeting in livestock have dramatically altered the logistic and biologic constraints. 2. What Has Changed? Cell-based methods of transgenesis by nuclear transfer or by ES chimerism have the critical advantage that genetic modification is carried out on cycling cell populations rather than directly on embryos. Hence, mass transfection is followed by selection for expression of a marker transgene in cultured cells and gives rise to hundreds of primary transfectants. These in turn give rise to limitless numbers of clonally derived cells, each with the potential to give rise to a transgenic founder animal. Significantly, DNA from modified cells can be prepared and characterized prior to their use in animal experiments. Subse- quent nuclear transfer uses only those cells carrying the desired modification, and 100% of resulting animals will be transgenic. This cell-based approach to transgenesis has recently been exemplified in livestock by the arrival of Polly (7), a transgenic sheep carrying a gene encoding the human blood-clotting factor IX. In principle, the ability to clone from cultured cells following genetic modification has provided the means to identify rare cells in which DNA has integrated into homologous sequences already present in the genome (gene targeting). Although this has not yet been exemplified in the cell populations proven in nuclear transfer, human somatic cells have been successfully targeted using those same techniques that are now routine for murine ES cells (8–13). It only remains to couple targeting in livestock- derived cells with nuclear transfer. A major application of cloning technology, therefore, will be to generate animals that carry subtle gene modifications generated by gene targeting in cultured somatic cells. The specific advan- tages to gene targeting that accrue are discussed in later sections. The only caveat at the time of this writing is the formal possibility that the properties of cells necessary to support targeting may be incompatible with those required for nuclear transfer.
- 18. Applications of Animal Transgenesis 5 3. ES Cells in Livestock Cloning from genetically modified somatic cells may be thought to render ES cells redundant for most applications in livestock. Murine ES cells, how- ever, are particularly well adapted to gene targeting and also provide an in vitro model of differentiation that may offer novel biomedical opportunities in transplantation therapy. It seems likely that interest in livestock ES cells will persist. There are numerous reports of ES-like cells in several species: hamster (14), mink (15), rat (16), chicken (17), sheep (18–20), cattle (21–25), pig (18,20,26,27), rhesus monkey (28), and human (29). None of these reports, however, has yet met the definitive test of germline transmission (there is no intended suggestion that this test should be applied in the special case of human cells); in spite of intensive effort, germline ES cell technology remains restricted to mice. Even were proven ES cells available, the ES route to transgenesis in farm animals would have the serious disadvantage that it requires an extra chimeric generation to establish transgenic founder animals. By contrast, the cloning option would generate transgenic individuals in the first generation. Perhaps the greatest disadvantage of the chimeric route is that it requires test breeding of all animals generated, including an unknown proportion (possibly 100%) that will be incapable of germline transmission. This contrasts with a cloning experiment in which failure is self-evident at an early stage by the absence of pregnancies. The aforementioned considerations raise the possibility that one might enjoy the best of both the cloning and ES options by employing targeted ES or ES-like cells in nuclear transfer. Here, there are several unresolved issues. Unlike murine ES cells, the livestock-derived ES-like lines reported to date are poorly adapted to single-cell cloning—a problem that will need to be overcome if these cells are to be used in gene targeting (reviewed in ref. 30). In addition, the early results of somatic nuclear transfer suggest that an important ingredient is that the nuclear donor be in a state of quiescence or Go (3). It seems likely that in addition to issues of cell-cycle compatibility between nucleus and ooplasm, the quiescent nucleus be configured in such a way as to favor reprogramming. Alternatively, it may be important that the somatic cell program of gene expression be shut down before the full devel- opmental program is reinitiated. In either event, ES cells (unlike fibroblasts) do not readily enter quiescence on serum starvation. Several important ques- tions remain unanswered: Can ES cells be entered into quiescence in some novel way? Can the differentiated derivatives of targeted ES cells be entered into quiescence? Do the ES-like cells currently available from livestock sup- port gene targeting?
- 19. 6 McWhir 4. A Hierarchy of Complexity Some transgenic applications in farm animals will involve complex target- ing technology and yet in biologic terms may have quite humble goals (simple loss of function mutation). In other instances, the technology may be relatively crude (as with pronuclear injection of growth hormone [GH] genes) whereas the biologic objective (to modify growth rate) is highly ambitious. In mouse transgenic programs, the biologic objective is usually straightforward—to observe and record the effects of ectopic gene expression or, in the case of gene targeting, the effects of loss of gene function. Here, although the pheno- typic consequences are often not predictable, the experiment will always be informative. In contrast to the mouse, many potential livestock applications (particu- larly in agriculture) will involve intervention in complex metabolic pathways in which the objective is to achieve a predetermined phenotypic change. Here, there is an additional challenge: it is necessary to accurately predict the phe- notypic consequences of a single genetic modification. Limited attempts to do this in order to increase the growth rate of transgenic pigs have led to unfore- seen consequences on animal health and fertility (31). The most straightfor- ward applications of transgenesis in livestock are those in which the objective is simply to harvest high levels of recombinant protein. In this case, there is no requirement to modify endogenous metabolic pathways, a physiologic response to transgene expression is neither required nor anticipated, and the risks of adverse effects on animal health and welfare are minimized. It is not surpris- ing, therefore, that the most successful transgenic applications to date have involved expression of human therapeutic proteins in the milk of transgenic sheep (32), pigs (33), cows (34), and goats (35). 5. Biopharming Biopharming is the commercial production of pharmaceuticals from the body fluids of transgenic animals. Although most attention has centered on the mammary gland (for reviews see refs. 36–39), other body fluids may have particular benefits for certain applications. For example, human GH has been expressed in mouse bladder epithelium under the control of the mouse uroplakin promoter (40). Advantages of bladder production might include the ability to harvest from all animals at all stages of their lives and the small number of other proteins from which the recombinant protein need to be puri- fied. Transgenic swine have been generated that express human hemoglobin in their blood as a potential cell-free substitute for human plasma (41,42). How- ever, this blood-based approach has been hampered by difficulty in separating human hemoglobin from its porcine counterpart.
- 20. Applications of Animal Transgenesis 7 By far the most readily harvestable source of recombinant protein is milk. While fermentation technologies and transgenic plant alternatives may be favored for some applications, the mammary gland provides several general advantages. Milk is a less complex fluid than blood, thus enhancing the pros- pects for rapid purification of recombinant protein. In addition, milk proteins are present in the circulatory system at undetectable or very low levels, thus minimizing potential animal health problems associated with high circulating levels of metabolically active proteins. Unlike fermentation-derived products, recombinant proteins produced in the mammary gland are posttranslationally modified in a manner that closely mimics their modification in humans (43), and are more likely to be stable, have high biologic activity, and be non- immunogenic in patients. While the mammary gland may be the preferred option in this regard, there is still scope for improvement. For example, some proteins purified from milk have a lower than expected molecular mass (44– 46); the ovine or bovine mammary gland product does not exactly mimic the human-derived protein. At least one group has addressed this issue by the coinjection of a furin transgene designed to increase the level of posttransla- tional modification (47). Production of pharmaceutical proteins in the transgenic mammary gland is rapidly being commercialized to produce products such as _-1 antitrypsin for treatment of emphysema and cystic fibrosis (48); the blood-clotting factors antithrombin III (35), factor VIII (49,50), factor IX (5,51), and fibrinogen (52) for treatment of bleeding disorders; and protein C (46,53) for treatment of blood clots. Recombinant antithrombin III and _-1 antitrypsin from transgenic live- stock are now undergoing phase III and phase II clinical trials, respectively, and this first generation of transgenic livestock has already spawned a signifi- cant biopharming industry. Most of the achievements in biopharming to date have employed pro- nuclear injection. How might cell-based techniques and gene targeting be used to improve the rate and direction of progress? As with any biologic system, there are upper limits to the synthetic potential of the mammary gland. One way to boost the production of therapeutic proteins would be to delete nonessential milk protein genes and simultaneously replace them with the desired transgene by gene targeting. This approach would not only intro- duce the transgene into an active site in the genome, but would simulta- neously provide excess synthetic capacity by knocking out the gene for a competing high-volume protein. This method may be essential to providing proteins that are required in very large quantities such as human serum albu- min (potentially useful in the treatment of burns). Gene targeting also can be used to improve the level and repeatability of transgene expression. Microinjection of DNA into the pronucleus usually
- 21. 8 McWhir results in multiple copies of the transgene being integrated in large arrays. In most cases, the level of expression of the transgene is not correlated with the number of copies and is subject to random effects of elements at the site of incorporation. In practice, up to 10 transgenic lines may have to be analyzed to obtain a single line in which the transgene is expressed in the desired temporal and spatial manner. A potential solution to this problem is to introduce the transgene into chosen sites in the genome by homologous recombination in ES cells. In mice this strategy has been used successfully to introduce a lacZ cas- sette into the hypoxanthine phosphoribosyl transferase (HPRT) locus (54), to replace the `-globin (55) and _-lactalbumin genes (56), and to introduce a bcl-2 minigene into the HPRT locus in ES cells (57). A variation on this theme involves an in vitro prescreen of marked, random sites for appropriate transgene expression followed by targeted replacement to introduce a transgene into the same site (58). Cloning not only facilitates cell-based transgenesis but also carries innate advantages. Based on averages of the data reported in the “Dolly” article (5), the proportion of nuclear transfer embryos that develop to term is low (approx 1.0%). Fortunately, much of this cost is owing to embryos that fail prior to reimplantation into foster females and is borne in the laboratory rather than on the farm. The proportion of embryos transferred into final recipients that give rise to lambs rises to 6.0%, and since two embryos are generally implanted per recipient, the proportion of recipients that give rise to nuclear transfer lambs rises to about 12%. Although there is still a large requirement for ani- mals to act as embryo donors, cloning compares favorably in efficiency with pronuclear injection. PPL Therapeutics, in collaboration with Roslin Institute, have recently generated the first additive transgenic sheep, Polly, using cell- based transgenesis (7). Even with the present rates of nuclear transfer success, it was estimated that 2.5 times as many sheep would have been required to create Polly by pronuclear injection. The practical consequence of this effi- ciency gain is that more potential therapeutic products will be tested. Another barrier to the testing of novel milk-derived recombinant proteins is difficulty in obtaining sufficient quantities of purified protein from single founder ani- mals (often male) for preliminary trials. This alone can be a sufficient commer- cial risk to rule out many potential applications. The cloning option provides a means with which to generate multiple female founders in a single generation. 6. Nutraceuticals Closely related to biopharming is the idea that genetic modification of milk proteins could be used to improve the nutritional or industrial properties of milk. A major nutritional objective is the humanization of bovine milk for the infant formula market. Transgenic cattle have been generated that carry the
- 22. Applications of Animal Transgenesis 9 cDNA for human lactoferrin (58), the major whey protein in human milk (although of low abundance in bovine milk). Lactoferrin may also play roles in iron transport and in protecting against bacteria. Many other strategies toward the humanization of milk have been widely discussed (60–63). In one example, human _-lactalbumin was introduced into mice in order to mimic the balance of whey to casein characteristic of human milk (54). Although several such ideas have been modeled in transgenic mice (for reviews, see refs. 60–63), only the lactoferrin approach has been attempted in livestock. Potential applications of transgenesis to alter the industrial properties of milk include modifying the casein content to alter milk-clotting properties, altering the proportion of hydrophobic residues in `-casein to improve its emulsify- ing properties (63), and improving the rate of maturation of cheese by intro- ducing an altered _s1-casein transgene (63). According to a 1990 estimate, a 20% increase in the content of _s1-casein would be worth almost $200 million annually in the United States alone. Again, none of these strategies have yet been exemplified in livestock. A striking aspect of nutraceuticals is that in spite of the broad range of iden- tified opportunity, there are few examples of reduction to practice. This has been attributed in part to the relatively low value of agricultural vs biomedical products, and in part to the inherent conservatism of agricultural industries (58). Consideration of the time and money required to generate sufficient num- bers of genetically modified animals to support, e.g., the cheese industry sug- gests that a certain amount of conservatism may be appropriate. In common with other agricultural applications, the greatest barrier to progress lies in the fact that engineered milks destined for human consumption are still not broadly acceptable to the consumer and in many countries are proscribed by law. 7. Animal Models In mice, the ES system allows us to re-create precisely, genetic lesions that are associated with human genetic disease (64). The production of gene-tar- geted mouse models has become routine (for a detailed description of target- ing, see ref. 65). In livestock, the practicality of engineered animal models will be sensitive to the added value of the livestock model over the corresponding mouse model and for broad application will require the development of gene targeting in livestock cell lines. With animal models in general, the resulting phenotype is usually anticipated, although species differences frequently con- found this expectation. Mouse models of cystic fibrosis, e.g., fail to present the same lung pathology characteristic of the human disease (66), and mice lack- ing the gene whose dysfunction in humans is associated with Lesch Nyhan syndrome (HPRT) are overtly normal (67,68). While it is clear that the major- ity of mouse models have been invaluable in the study of human disease,
- 23. 10 McWhir it is equally true that for certain diseases, the mouse models have serious limitations. Livestock species share similarities with humans in anatomy, size, physiol- ogy, and life span, which often renders them better models than rodents. The pig has been particularly useful in the past as a model of kidney dysfunction, ischemic heart disease, hypercholesteremia, and atherosclerosis (reviewed in refs. 69). The sheep has been proposed as a potential animal model of the human condition cystic fibrosis, which results from defects in the cystic fibro- sis transmembrane conductance regulator (CFTR) gene (70). Ovine CFTR pro- tein is 95.3% similar to the human amino acid sequence and has a very similar expression pattern. In addition, the sheep lung epithelium shares anatomic, functional, and electrophysiologic similarities with the human (68). At least one group has embarked on a search for a spontaneous CFTR mutant among commercial flocks in New Zealand (71); however, the success of such a large- scale screening program cannot be taken for granted. To date, livestock models have been restricted to spontaneous mutants and to pharmacologic models in which wild-type animals are challenged with disease-causing agents. A single but fortuitous exception to this generalization may be the GH pigs that were originally generated in an attempt to enhance growth rate (30). Although these animals have not proven useful in agriculture, it has been suggested that they may provide a model of the human growth disorder acromegaly (69). Cell-based transgenesis in livestock and the possibility of gene targeting in these species open new opportunities for engineering large-animal mod- els. The ovine CFTR gene, e.g., may be a prime candidate for knockout by gene targeting. Even were a spontaneous ovine mutant available, a significant advantage to the engineered model is that one or two of the commonly occur- ring point mutations in cystic fibrosis patients could be precisely mimicked. Other candidates include the prp genes of sheep and cattle. Misfolding of the prp gene product (the prion protein) is associated with the spongiform encepha- lopathies: scrapie in sheep; BSE in cattle; and CJD, GSS, and Kuru in humans. Although mice carrying inactive prp genes show certain subtle alterations in circadian rhythms (10), they are fully viable, developmentally and behavior- ally normal, and resistant to scrapie (73). To confirm this circumstantial evi- dence for the control of scrapie by PrP, it would be invaluable to determine whether sheep carrying inactive prp are similarly resistant and to establish whether they can carry and transmit the infective agent. One of the general limitations of generating knockout animals in livestock is that in most instances only the homozygous knockout is useful. This pre- sents particular problems in disseminating loss-of-function genotypes into commercial populations. As a consequence, PrP-deficient animals are most likely to be restricted to the small numbers required for fundamental research,
- 24. Applications of Animal Transgenesis 11 and possibly to the generation of new cell lines for use as nuclear transfer donors for biomedical applications. If PrP-deficient animals can be shown to be incapable of carrying the infective agent or agents, then animals cloned from such prp-deficient cell lines could be declared scrapie/BSE-free. 8. Xenotransplantation At any one time, some 5000 persons await suitable organs for heart trans- plants in the United Kingdom and about 50,000 in the United States. Many of these patients will die before a suitable donor is available. According to one estimate, only 10% of those patients who could benefit from a heart transplant actually receive one (74). There is, therefore, considerable interest in geneti- cally engineering pigs so that their organs will be acceptable to the human immune system (xenografting). The major epitope leading to hyperacute rejection of xenografts in humans is a sugar residue produced by the action of the enzyme _1,3 galactosyl trans- ferase. This enzyme is inactive in humans and Old World primates but is func- tional in all other mammalian species. The binding of xenoreactive antibodies following xenotransplantation activates the classic complement pathway lead- ing to rapid (within minutes) cell lysis (reviewed in ref. 75). Hence, two poten- tial transgenic strategies to address the problem of hyperacute rejection are either to block the complement pathway or to reduce levels of the major xenoreactive epitope, Gal _ 1,3 Gal. Two lines of transgenic pig have been produced that carry transgenes encoding two of the three main regulators of human complement activation: human decay accelerating factor (hDAF) (76), and human CD59, respectively (77). In perfusion tests, the genetically modified hearts are protected from the action of human complement (76,77) and following transplantation to cyno- molgus monkeys, hDAF hearts lead to a significant increase in survival (75). Mice rendered dysfunctional at the _1,3 galactosyl transferase locus by gene targeting are fully viable, and several attempts have been made to reduce gal transferase activity in pigs by additive transgenesis. Transgenic mice and pigs have been generated that express human fucosyl transferase (78,79). This gene is not normally expressed in pigs and mice, and its transgenic expression leads to reduced levels of the Gal _ 1,3 Gal epitope. Further reduction in Gal _ 1,3 Gal levels was obtained by combined expression of _-galactosidase and _1,3 fucosyltransferase (80). Perhaps the optimal transgenic strategy would be to use gene targeting to inactivate the _1,3 galactosyl transferase gene, although this awaits the development of gene targeting and of somatic cloning in pigs. Regardless of the method employed, controlling the hyperacute response will not prevent eventual T-cell rejection, and successful xenotransplantation must deal with this downstream problem either by improvements to immuno-
- 25. 12 McWhir suppression regimes or by further engineering strategies. If we look to the future and make certain optimistic assumptions, it is possible to imagine the eventual humanization of the porcine major histocompatibility complex, although this sort of strategy would depend greatly on further advances in chro- mosome engineering. Safety issues surrounding xenotransplantation have led to intense public debate. Of particular concern is the risk of zoonoses following the demonstra- tion that human cells in vitro can be infected by an endogenous porcine retrovirus (81), although it remains unclear if infection can also occur in the normal in vivo situation. Although porcine pancreatic islets have been trans- ferred to human patients for some time with no evidence of viral infection, xenotransplantation involves extra factors associated with viral activation such as heavy immunosuppression. At present time, most countries have imposed a moratorium on human transplantation of xenografts. The heavy demand for organs may nonetheless make it likely that clinical trials will proceed in the near future. Guidelines to minimize risk are presently being prepared by the appropriate regulatory bodies. 9. Agriculture The first application of additive transgenic technology to improving the per- formance of livestock was the introduction of extra genes for GH in an attempt to improve the growth rate and feed efficiency of pigs (30). The resulting pigs did show improvements in feed efficiency and fat content, but they also suf- fered from a variety of debilitating defects associated with poor control of transgene expression. Since then, potential applications of transgenesis in agri- culture have been widely reviewed (56,82,83), but have seldom been reduced to practice. Notable exceptions include sheep engineered for improved wool production either by transfer of bacterial genes for cysteine synthesis (84); by addition of genes for wool keratin proteins, which improve wool fiber ultra- structure (85); or by expression of insulin-like growth factor-1 in hair follicles (86). A second promising area for agricultural transgenesis is disease resis- tance. Pigs have been generated that express low levels of the murine Mx1 gene associated with resistance to influenza (86). It has been suggested that high expression of Mx1 may be developmentally lethal (87). Hence, as with GH pigs, it is again apparent that successful transgenic programs often require extremely tight control of transgene regulation. In general, agricul- tural applications of transgenesis have been hindered by inefficiencies in the production of transgenic founders, in the proportion of founders with appro- priate transgene expression, and in dissemination of transgenic stock to com- mercial populations.
- 26. Applications of Animal Transgenesis 13 Two major sources of inefficiency in the production of transgenic founders have been the large numbers of embryos required for injection and the large requirement for recipient females to bring nontransgenic embryos to term. The production of in vitro matured and fertilized oocytes taken from ovarian fol- licles has dramatically reduced the cost of transgenic programs in cattle (88,89). Attempts to reduce the number of transfers of nontransgenic embryos by poly- merase chain reaction screening prior to transfer have been hampered by diffi- culty in distinguishing between integrated and nonintegrated transgenes; however, such techniques have proven useful in increasing the percentage of transmission from transgenic founders (89). Identification of transgenic embryos immediately following microinjection has been achieved in mice by inclusion of a fluorescent marker, green fluorescent protein, whose product can be visualized prior to transfer without harming the embryo (90). It remains to be seen if this technique can be adapted to large-animal transgenesis. An alternative technique based on in situ hybridization of metaphase spreads obtained from biopsied material also shows promise (91). Our understanding of gene expression has improved in recent years, and it now seems possible that in future applications many of the problems associ- ated with inappropriate transgene expression can be avoided. Tissue-specific and copy number–independent expression of transgenes can be improved by the inclusion of a locus control region in the transgenic construct (92,93). The complementary approach is to introduce the transgene into chosen sites in the genome by homologous recombination. This latter strategy has been used suc- cessfully in mice (51–55). Other approaches are the cointroduction of the transgene with fragments that “rescue” genes from positional silencing (94) and the flanking of transgenes with inverted terminal repeat sequences from adenoassociated virus (95); however, the latter method is not yet exemplified in mammalian cells. ES cells also can be modified by the introduction of a loxP site that is recognized by the bacterial site-specific recombinase Cre (96,97). In a subsequent step, and in the presence of Cre recombinase, this allows site- specific insertion of transgenes that carry flanking loxP sites (97–99). Improvements in transgenic technology per se will not affect the down- stream economics of animal breeding. Incorporating transgenes into commer- cial populations by conventional breeding will take many years and will reduce the rate of genetic progress by conventional selection. In one study simulating the effect of transgenic strategies to increase the male:female sex ratio in beef cattle, it was concluded that reduction in genetic progress in other characters could offset any gains in average growth rate (100). Clearly, any transgenic program in agriculture that required the establishment of a nucleus herd would need to offer a very large potential benefit (reviewed in ref. 101). However, if we assume substantial improvements in the efficiency of embryo cloning and
- 27. 14 McWhir cell-based transgenesis, then it is possible to imagine the provision of cloned, transgenic embryos directly to the producer. In this scenario, transgenic strate- gies offering even modest improvements could be introduced to the most pro- ductive genotypes and disseminated to commercial herds in a single generation. Even without transgenesis, producers could raise the average performance of their herd to that of the best within a single generation by purchasing cloned embryos. An added advantage is that all embryos could be of chosen sex. The development of coherent strategies for the application of cloning and transgenesis in agriculture will require careful consideration of selection schemes to accommodate continued genetic improvement by traditional breeding. In addition to the efficiency advantages of cell-based transgenesis, this tech- nique opens the possibility of applying gene-targeting techniques to livestock to generate novel phenotypes. One example of a candidate “knockout” gene is myostatin, which has been identified as the gene whose dysfunction results in the double muscling condition in cattle (102). Other candidate genes may well be identified in the search for quantitative trait loci (QTL) in livestock. 10. Impact of Gene Mapping and Functional Genomics Genetic linkage maps are now being developed for many species. In cattle, sheep, and pigs, this has allowed investigators to map genes that have a large effect on quantitative traits—the so-called QTL. Many genes on the human and mouse maps have resulted from mapping expressed sequence tags (ESTs), short sequences obtained from cDNA libraries that are often highly conserved across species (for reviews, see refs. 103 and 104). How the information obtained from mapping programs can be used to establish gene function is the new and still poorly defined area of “functional genomics.” At least one com- pany, Lexicon, now offers a database of sequence tags obtained from randomly “knocked out” genes in murine ES cells. Hence, in principal, candidate QTL genes obtained from EST databases can be cross-referenced to an ES “knock- out” in the Lexicon database and the corresponding mouse generated and ana- lyzed for phenotype. So far only a few QTL genes have been identified. Two of these are the melatonin receptor (105) and the KIT gene (106), which are associated with coat color. Arguably, these are not strictly QTLs because coat color is not a quantitative trait. However, their identification has arisen directly from the QTL effort, so it is perhaps appropriate in this case to stretch the definition. Others include genes associated with stress susceptibility in pigs (107) and hyperplasia in cattle (101). Transgenesis, both in model species and in live- stock, is likely to play a key role in functional genomics by providing the information that either proves or disproves an association with phenotype.
- 28. Applications of Animal Transgenesis 15 Transgenesis also could be used to introgress transgenes into commercial popu- lations. Although a discussion of the economics of transgenesis vs traditional allele introgression is beyond the scope of this review, it seems likely that the key advantage to the transgenic route would be the opportunity to introduce the transgene on a variety of genetic backgrounds. 11. The Future: Gene and Cell Therapies ES cells can be induced to differentiate in vitro into a variety of lineages with potential therapeutic value such as hematopoietic stem cells (108), neural stem cells (109), and myoblasts (110). In principle, one can imagine the appli- cation of nuclear transfer to generate cloned embryos for the subsequent isola- tion of isogenic ES cell lines tailored to individual patients. Isogenic cell lines could then be genetically modified to repair oncogenes or to replace defective alleles and returned to the patient following in vitro differentiation. The sim- plest example of such an approach would be the complete replacement of the patient’s hematopoietic system following oncogene repair by gene targeting. Clearly, applying such a strategy in medicine raises serious ethical issues. The most troublesome of these arises from the requirement for recipient oocytes. So, is the oocyte really necessary? That nuclear transfer in embryos may be simply a special instance of a more general nuclear reprogramming phenomenon is an exciting prospect. As we learn more about the processes involved in nuclear reprogramming in the embryo, it seems at least possible that we may identify factors that can be used to achieve the direct transformation of somatic cells to ES cells without the requirement for an embryo’s intermediate. One possibility is that the ES cytoplasm itself may have reprogramming activity. It is intriguing that in fusions of embryonic germ cells with thymocytes, the ES phenotype is dominant (111). 12. Conclusion This discussion has been rather broad in scope and, of necessity, has been less than exhaustive in its treatment of individual areas of application. For more comprehensive information, the reader is referred to the many excellent reviews cited throughout. In addition to the areas of application discussed, it seems likely that unusual uses will arise that are neither biomedical nor agricultural. For example, Nexia Pharmaceuticals (Canada) has generated transgenic goats expressing spider silk proteins in their milk and anticipate wide use of the recombinant product in the manufacture of highly resilient industrial materi- als. Other omissions in this limited review include avian transgenesis and detailed discussion of the ethical, public acceptability of and economic con- straints to the further uptake of transgenic technology.
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- 37. Gene Transfer in Drosophila 25 II TRANSGENESIS IN INVERTEBRATE AND LOWER VERTEBRATE SPECIES
- 38. Gene Transfer in Drosophila 27 27 From: Methods in Molecular Biology, vol. 180: Transgenesis Techniques, 2nd ed.: Principles and Protocols Edited by: A. R. Clarke © Humana Press Inc., Totowa, NJ 2 Gene Transfer in Drosophila Mark J. O’Connor and William Chia 1. Introduction The generation of germline transformants in Drosophila melanogaster has relied on the utilization of transposable elements to effect the chromosomal integration of injected DNA (1,2). The success of this approach has depended largely on our understanding of the biology of P elements and the syncytial nature of the early Drosophila embryo. The first 13 embryonic divisions fol- lowing fertilization are nuclear, resulting in the formation of a syncytium. Con- sequently, if microinjection into the posterior end of the embryo is carried out prior to cellularization, a proportion of the microinjected DNA will be present in the cytoplasm of the pole cells, the progenitor cells of the germline. In practice, the DNA to be injected comprises two components. The first con- sists of a helper plasmid containing a defective P element that, although capable of producing the P transposase, which can act in trans to mobilize P transposons, is itself immobile (see Note 1). The second component consists of a transposon construct in which the sequence to be integrated as a transgene is situated between the 31-bp P element inverted terminal repeats along with a suitable marker (see Note 2). The transposase produced by the helper plasmid will act on the inverted repeats of the transposon construct and facilitate the integration of the transposon into essentially random chromosomal sites of the recipient’s germline. Both P element biology and the characteristics of P element–mediated transformation have been reviewed extensively (e.g., see ref. 3). In this chapter, we deal prima- rily with the technical details necessary for obtaining germline transformants. 1.1. Outline of Events Involved in Generation of Germline Transformants 1. Construct the desired plasmid containing the transgene, marker, and necessary P element sequences for transposition.
- 39. 28 O’Connor and Chia 2. Coinject the transposon along with a defective helper plasmid supplying the P element transposase. 3. Mate the survivors (Go) to an appropriate strain that will allow for the scoring of the marker carried on the transposon construct. 4. Select for transformed progeny that have acquired the marker carried on the transposon and balance the transformants. 5. Test the structure and copy number of the transgene(s) in the transformant lines. 6. Choose unrearranged single insert lines for phenotypic analysis. 2. Materials 2.1. Microinjection System Figure 1 shows the injection apparatus we use. This system consists of the following: Fig. 1. Typical arrangement of the apparatus used for injection of Drosophila embryos.
- 40. Gene Transfer in Drosophila 29 1. Leitz micromanipulator. 2. Nikon inverted phase-contrast microscope. 3. Vibration-free table, on which the microscope is mounted. 4. Loaded needle, containing the DNA to be injected. 5. Collar (Narishige, Tokyo) into which the needle is placed, which, in turn, is attached to the micromanipulator. Although the micromanipulator is used to position the needle, injection is carried out by moving the microscope stage with the embryos on it. We use an air-filled system to deliver the DNA into the embryos. This consists of a 60-mL glass syringe attached to the collar by a piece of rubber tubing (Narishige Teflon™ tubing also may be used). This system may appear very basic, but we find that the syringe imparts adequate control of DNA delivery without producing the problems often encountered when using a fluid-filled transmission system, and the system has the advantage of being much cheaper. Injection needles are prepared from boro- silicate capillaries (e.g., Clark Electromedical [Reading, UK] GC100TF-15 capillaries, which contain an internal filament) using a pipet puller. A rela- tively inexpensive two-stage vertical needle puller can be used, such as the PB-7 model from Narishige. 2.2. Fly Requirements In general, a large number of embryos (in the region of 500–1000) need to be injected for each construct in order to produce several independent transformants. In our hands, between 25 and 75% of injected embryos will hatch as larvae. Approximately 50% of the larvae will survive as adults, and between 50 and 80% of the surviving adults will be fertile. Each surviving adult will be individually mated, and approx 200 progeny from each mating will be scored for the marker present on the transposon construct. Although the frequency with which germline transformants are produced varies depending on the construct injected (4), in general, on the order of 10% of the surviving adults will produce at least one germline transformant among its progeny. Therefore, it is reasonable to aim at obtaining about 100 adult survivors for any given construct injected. We usually collect only one transformant from the progeny derived from each surviving adult with which to estab- lish stocks. This ensures that different transformants originated from inde- pendent events. Since the injections must be performed prior to pole cell formation, 1-h embryo collections are used (see Subheading 3.3.). Therefore, the fly strain used for embryo collections must be robust enough to provide sufficient eggs (at least 100) during a 1-h interval. One further consideration is that the presence of defective P elements in the injected host strain can affect the frequency of
- 41. 30 O’Connor and Chia transformation. Consequently, care should be taken to ensure that such ele- ments are not present in the chosen host strain. 2.3. Miscellaneous 2.3.1. Preparation of DNA 1. Qiagen anion-exchange columns. 2. Injection buffer: 5 mM KCl, 0.1 mM Na phosphate, pH 7.8. 3. Millipore filters (0.45-μm). 2.3.2. Egg Collection and Egg Processing 1. Egg collection chamber. This can be made from open-ended plastic cylinders of any sort large enough to contain a few hundred flies. The chambers should have fine gauze placed over one end for ventilation, and once the flies have been placed into the chambers, small Petri dishes containing yeast-glucose food and smeared with moist, live yeast are taped to the other end. 2. Glass or plastic tube with a nitex gauze over one end. 3. Freshly diluted 50% household bleach. 4. 0.02% (v/v) Triton X-100. 5. Black nitrocellulose filters. 6. Fine paint brush. 7. Cover slips (22 × 40 mm). 8. Solution of Sellotape in n-heptane. 9. Voltalef oil. 3. Methods 3.1. Preparation of DNA Plasmid DNA for microinjection may be prepared either by the cesium chlo- ride–ethidium bromide centrifugation method, or by the more convenient Qiagen anion-exchange columns produced by Qiagen (Chatsworth, CA). The latter method produces clean DNA and is not only quicker but also avoids the use of ethidium bromide and organic solvents, such as phenol and chloroform, which could potentially reduce embryonic survival rates. The concentration of DNA for microinjection needs to be quite high (between 400 and 600 μg/mL) with “helper” plasmid, if used, at a concentration of 200 μg/mL. The DNA to be injected should be ethanol precipitated and given an 80% ethanol wash before being redissolved in injection buffer. Aliquots of 20 μL can then be stored at –20°C. Prior to loading the DNA into injection needles, the aliquots should be heated to 65°C for 10 min to ensure that the DNA is fully dissolved and then spun through 0.45-μm Millipore filters for a couple of minutes to remove any dust or particles, which could potentially block the needle.
- 42. Gene Transfer in Drosophila 31 3.2. Preparation of Needle To obtain a needle that possesses the appropriate shape, the first-stage pull should generate a stretch with a length of about 8 mm and a diameter of approx 200 μm. The heating filament should then be moved to the center of this stretch so that the second pull produces a very fine tip of approx 2 mm in length with an end of between 1 and 5 μm in diameter. The heater settings for the first and second pull will need to be determined empirically in order to produce a good- quality needle. Once a needle has been prepared, the simplest way to load it with the DNA solution is to add 1 to 2 μL of the injection DNA at the back of the capillary with a micropipet. The internal filament that runs along the length of the capil- lary draws the DNA solution to the front of the needle, which can then be placed into the collar of the microinjection system. The survival of injected embryos is affected to a large extent by the sharp- ness of the needle. To obtain a sharp point, the needle can be broken at an angle against a cover slip mounted onto a glass slide. This process is visual- ized using the inverted-phase microscope and is made easier by placing a drop of Voltalef halocarbon oil on the junction between the slide and the cover slip where the needle is to be broken. When the needle breaks, a small amount of the oil can usually be seen to enter the tip. The flow of DNA can then be tested by applying a little pressure to the syringe. The needle is now ready to use for microinjection. In between injecting embryos, the needle can be lowered into a small (5-cm) Petri dish lid containing Voltalef halocarbon oil. This helps prevent evapora- tion of the DNA solution and the concomitant clogging of the needle that can otherwise occur. 3.3. Egg Collection Synchronous and abundant batches of eggs are required for injections. In gen- eral, 300–600 adults will produce enough eggs for a few days of microinjec- tions. The flies should be transferred into collection chambers. To optimize egg laying, the flies should be kept at 25°C for a further 2 d in the chambers before starting egg collections for injection, and the Petri dishes containing the food should be changed every day. At the end of the second day, and every subsequent day, the flies should be transferred to 18°C overnight and then returned to 25°C on the morning of collection. The first hour’s collection should be discarded because female flies tend to retain eggs until fresh food is sup- plied. Thereafter, at 60-min intervals, the collection plates can be removed and replaced with new ones.
- 43. 32 O’Connor and Chia The eggs to be injected are washed off the collection plates with distilled water and passed down a glass or plastic tube containing a nitex gauze over one end to retain the embryos. The eggs are then ready for dechorionation. 3.4. Preparation of Embryos for Microinjection 1. The first step in preparing the eggs for microinjection requires the removal of the tough outer chorion (see Note 3). To achieve chemical dechorionation, place the tube with nitex gauze and embryos into a beaker containing 10 mL of a 50% solution of household bleach. Gently shake the beaker and tube and, after 2–2.5 min of dechorionation, dilute the bleach by adding an equal volume of a 0.02% Triton X-100 solution. Then remove the tube from the beaker and wash the eggs thoroughly with distilled water. 2. Transfer the embryos onto a black nitrocellulose filter with a fine paint brush and line up along one of the ruled lines on the filter in such a way that the micropile is nearest to you. It is important to keep the filter damp to prevent the eggs from drying out. 3. When 50–60 embryos have been lined up, transfer them to a 22 × 40 mm cover slip; the cover slip can be made adhesive by the prior application of a solution of Sellotape in n-heptane. Stick the cover slip with attached embryos onto a micro- scope slide using a small drop of Voltalef oil and a little pressure. Place the whole slide inside an airtight box containing silica gel in order to desiccate the embryos (see Note 4). 4. At the end of the desiccation period, take the eggs out of the box containing the silica gel and cover with a layer of Voltalef oil. This oil, although being oxygen permeable, is water impermeable and therefore prevents any further desiccation of the embryos. The embryos are now ready to be injected. 3.5. Microinjection of Drosophila Embryos 1. Once the needle is lifted safely out of the way, place the slide containing the embryos on the microscope stage so that the eggs have their posterior facing the needle. Use the micromanipulator to bring the needle into the same plane as the line of eggs. 2. Bring the tip of the needle level with the center of the first egg; this is gaged by running the very end of the needle up and down the edge of the embryo. This method ensures that the needle will not slide over the surface of the egg and will also help decrease the amount of damage to the embryo. Then move the embryo toward the needle with a purposeful motion so that the vitelline membrane is just penetrated. Draw back the needle so that the tip is only just within the cytoplasm. Most of the embryos to be injected will be in the early cleavage stage (15 min to 1 h 20 min) and will have a space between the posterior pole and the vitelline membrane. It is important that the needle be inserted through the space and that the DNA be deposited in the posterior pole of the embryo proper. It is here, at the posterior pole, that the germline will be formed. Next, inject the embryo with a quantity of DNA solution equivalent to approx 1% of the egg’s total volume and
- 44. Gene Transfer in Drosophila 33 remove the needle. Repeat the procedure until all the embryos have been injected (see Note 5). 3. Kill any embryos in which pole cell formation has already taken place running them through with the needle. Do not count these among those eggs that have been successfully injected. 4. Remove the cover slip containing the injected embryos from the slide and place onto a flat yeast-glucose-charcoal plate. Apply a further thin layer of Voltalef oil to the line of embryos and place the plate into a box kept humid by damp tissues. Then place the box on a level surface in an 18°C incubator for 48 h. If the plate is not kept level, the Voltalef oil will run off, and the embryos will overdesiccate and die. 5. After this time, count the hatched larvae, transfer into vials containing fly food, and return to the 18°C incubator to develop. The percentage survival to first instar larvae can be determined by dividing the number of survivors by the number of successfully injected embryos. 4. Notes 1. There exist a number of plasmids that, when injected, can provide the P element transposase necessary to mobilize the coinjected transposon. Two of the most widely used sources are pp25.7wc (wings clipped; ref. 5) and pUChs62-3 (6). The wings-clipped transposase source contains a complete 2.9-kb P element in which the last 22 bp has been deleted so that the element is no longer mobile. The pUChs62-3 transposase source comprises the engineered transposase gene (62-3) in which the intron separating the second and third exons (normally only spliced in the germline) has been removed (6,7). This modified transposase gene is placed under the control of the HSP70 promoter, although the constitutive expression of this promoter is of a sufficiently high level such that heat shock is not necessary. Injecting this construct will result in the transient expression of a functional transposase in both germline and somatic tissues. An alternative approach to coinjecting a plasmid that provides a transposase source is to inject embryos that possess a chromosomal source of the 62-3 transposase (3). 2. Many vectors suitable for constructing transposons have been described. We con- sider here three of the more widely used ones. The transformation vectors based on rosy (ry) as a scorable marker were the first to be used. One of the most versatile versions of the ry-based vectors is pDM30 (8). The major advantage of using ry-based vectors is that since 1% of wild-type ry expression is sufficient to yield ry+ eye color, insertions into positions that result in a low level of expres- sion can still be recovered. However, the ry gene is large (usually a 7.2-kb HindIII fragment carrying ry is used), and this results in a less-than-optimal vec- tor size. For example, the largeness of ry-based vectors can make the construction of transposons more difficult and can also contribute to a decreased transforma- tion frequency. Another popular series of transformation vectors use the white (w) gene as a marker (9). In the most widely used w vectors, a mini-white gene (10) with a
- 45. 34 O’Connor and Chia subthreshold of w+ activity is used. There are several advantages associated with these mini-w-based vectors. First, the gene is small, ~4 kb, compared with ry. Second, since mini-w has subthreshold activity, for most insertions, flies that are heterozygous for mini-w can be distinguished from flies that are homozygous on the basis of eye color. Finally, w is easier to score than ry when large numbers of flies are involved. The latest versions of these vectors (the Casper series) may be requested from the Thummel or Pirrotta laboratories. A third series of vectors are those based on G418 antibiotic selection (11). In these vectors, the bacterial neomycin resistance gene is used as a selectable marker in place of visible markers such as ry and w. The advantage of using such vectors is that transformants can be selected on Drosophila food containing G418 (usually 500–1000 mg/mL), eliminating the chore of screening many flies for a visible marker. However, the major disadvantage is that the window of G418 concentration that will allow true transformants to survive, but that will reduce the leakage of nonresistant animals to an acceptable level, is narrow. Conse- quently, transformants owing to insertions into chromosomal sites resulting in a low level of expression will not be recovered. Other transformation vectors, such as those based on Adh, which allow for selection on media containing alcohol, have also been described. In addition, a transformation vector (pCaWc) in which both the transposon and the transposase are carried on the same plasmid molecule (with the transposase located outside the P element 31-bp repeats) has been successfully employed for obtaining transformants (12). There are also “shuttle vectors” that greatly facilitate the construction of complex transposons. These vectors (e.g., pHSX, referred to in ref. 12) contain large polylinkers flanked by restriction enzyme sites such as NotI (which occurs only very rarely) and enable several DNA fragments to be assembled and then excised as one contiguous piece. The construct can then be inserted into the single NotI site of transformation vectors such as pDM30 or the Casper series. Finally, transformation vectors designed for placing genes under the control of HSP70 and actin promoters have been described (13), as have transformation vectors designed to facilitate the insertion of desired sequences upstream of a LacZ reporter gene to drive its expression (10,13). 3. Two methods of dechorionation can be employed: chemical and mechanical. However, we favor the chemical method because it is far easier and less time- consuming. 4. This stage is of vital importance if the embryos are to withstand being punctured and accommodate the volume of DNA being introduced. Moreover, this step of the procedure is probably the most crucial, in terms of survival rates, because there is only a narrow margin between a sufficient reduction in egg turgor and excessive drying, which kills the embryos. If possible, embryos should be pre- pared in an environment with constant temperature and humidity conditions, because this will facilitate the determination of the optimum desiccation time. However, if this is not possible, the experimenter will have to determine the des- iccation time empirically, since this will tend to fluctuate depending on the cli-
- 46. Gene Transfer in Drosophila 35 matic conditions. As a starting point, we generally have used desiccation times of between 10 and 15 min. 5. If the embryo has not been desiccated enough, or if too much DNA solution has been injected, cytoplasm may leak out of the egg, reducing its chances of sur- vival (see Fig. 2). We have found that increased survival rates can be achieved by removing the “bubbles” of cytoplasm. This is easily achieved by having a con- stant flow of DNA coming out of the needle, which is then brushed passed the line of embryos. References 1. Spradling, A. and Rubin, G. (1982) Transposition of cloned P elements into Droso- phila germ line chromosomes. Science 218, 341–347. 2. Rubin, G. and Spradling, A. (1982) Genetic transformation of Drosophila with transposable element vectors. Science 218, 348–353. 3. Robertson, H., Preston, C., Phillis, R., Johnson-Schlitz, D., Benz, W., and Engels, W. (1988) A stable genomic source of P element transposase in Drosophila melanogaster. Genetics 118, 461–470. 4. Spradling, A. (1986) P element mediated transformation, in Drosophila: A Prac- tical Approach (Roberts, D. B., ed.), IRL, Oxford, pp. 175–197. 5. Karess, R. and Rubin, G. (1984) Analysis of P transposable element functions in Drosophila. Cell 38, 135–146. Fig. 2. Microinjection of Drosophila embryos illustrating the region of the embryo targeted for injection. Note also the “bubble” of cytoplasmic material leaking from the embryo, which should be removed.
- 47. 36 O’Connor and Chia 6. Rio, D., Laski, F., and Rubin, G. (1986) Identification and immunochemical analy- sis of biologically active Drosophila P transposase. Cell 44, 21–32. 7. Laski, F., Rio, D., and Rubin, G. (1986) Tissue-specificity of P element transposi- tion is regulated at the level of mRNA splicing. Cell 44, 7–19. 8. Mismer, D. and Rubin, G. (1987) Analysis of the promoter of the nina E opsin gene in Drosophila melanogaster. Genetics 116, 565–578. 9. Klemenz, R., Weber, U., and Gehring, W. (1987) The white gene as a marker in a new P element vector for gene transfer in Drosophila. Nucleic Acids Res. 15, 3947–3959. 10. Pirrotta, V. (1988) Vectors for P element mediated transformation in Drosophila, in Vectors: A Survey of Molecular Cloning Vectors and Their Uses (Rodrigues, R. L. and Denhardt, D. T., eds.), Butterworths, Boston, MA, pp. 436–457. 11. Steller, H. and Pirrotta, V. (1985) A selectable P vector that confers selectable G418 resistance to Drosophila larvae. EMBO J. 4, 167–171. 12. Moses, K., Ellis, M., and Rubin, G. (1989) The Glass gene encodes a zinc finger protein required by Drosophila photoreceptor cells. Nature 340, 531–536. 13. Thummel, C., Boulet, A., and Lipshitz, A. (1988) Vectors for Drosophila P ele- ment-mediated transformation and tissue culture transfection. Gene 74, 445–456.
- 48. Oocyte Injection 37 III TRANSGENESIS IN THE MOUSE: OOCYTE INJECTION
- 49. Oocyte Injection 39 39 From: Methods in Molecular Biology, vol. 180: Transgenesis Techniques, 2nd ed.: Principles and Protocols Edited by: A. R. Clarke © Humana Press Inc., Totowa, NJ 3 Oocyte Injection in the Mouse Gary A. J. Brown and Timothy J. Corbin 1. Introduction 1.1. Mouse Production Colony To provide fertilized eggs for microinjection, a production colony needs to be established. This should be carefully planned in order to provide enough material for your requirements. There are several items for consideration in this regard, detailed in the following sections. 1.1.1. Mouse Strain Consideration of the chosen strain is important because of the differences in genetic makeup, parental suitability, fecundity, response to administered gonadotropins, or future experiments involving breeding against a specific genetic background. Popular strains for use in transgenic mouse production as embryo donors are C57/BL6 x SJL F1 animals, FVB/N (1), or C57BL6 x C3H F1s, whereas those used for embryo recipients include Swiss Webster or C57BL/6 x DBA/2 F1s. 1.1.2. Colony Size It is necessary to plan colony size according to expected frequency of injec- tions. A typical day of microinjection at most institutions involves the injection of upward of 200 embryos. To provide this number of fertile embryos, and to constrain operational expenses by limiting cage per diem costs and the number of animals to be used, the technique of superovulation (2) is frequently employed. Typically between 7 and 10 animals are superovulated to yield suf- ficient embryos for one day’s microinjection. These animals should be used such that the administered gonadotropins initiate the animals’ first estrous cycle (3.5–4 wk of age). Additionally, the same number of male stud animals to mate
- 50. 40 Brown and Corbin to the superovulated females will be required. Male stud animals should be used no more than twice weekly and tracked as to their ability to plug females. For the embryo recipient mice, a colony large enough to consistently pro- duce sufficient animals in estrus for each day’s injection effort is required. Planning involves knowing the length of estrous cycle for the strain used for embryo recipients. To vaginally plug these animals, a stud colony of vasecto- mized males is required. These should be monitored for their ability to plug estrous females. It is recommended that a strain with a different coat color be used for embryo recipient animals and vasectomized studs than used for the embryo donors for microinjection, to allow easy detection of any vasectomized male that may have re-ligated a vas deferens and regained fertility. 1.1.3. Efficiency of Superovulation The ability of the intended embryo donor strain to be superovulated should be considered, because some strains do not react well to gonadotropin treat- ment. Hybrid strains tend to respond well and to produce substantially higher egg yields than by natural matings, although some inbred strains also have a consistently good performance. 1.1.4. Parental Suitability Some strains exhibit poor parenting and cannibalize their pups with a high frequency, and these should be avoided, if possible. Often outbred animals such as ICR or Swiss Webster are suitable for use as embryo recipients, although any strain with a high mean litter size and size (for best postoperative recovery) will be effective. Outbred strains are usually less expensive and bet- ter suited for this element of transgenic production than inbred or hybrid strains. 1.1.5. Pseudopregnancy It is necessary to generate animals that will have a receptive environment to implanted embryos. This is carried out by inducing animals to exhibit pseudo- pregnancy through sham fertilization by vasectomized males. The intended recipient females ideally should be between 8 and 16 wk old when sham fertil- ized and a strain with proven parental abilities chosen for this purpose. 1.2. Collection of Fertilized Eggs The fertilized embryos used for microinjection should be approx 0.5 d postcoitum (p.c.), typically obtained from female mice that have been mated to stud male mice in the afternoon of the previous day. The goal is to time the mating of the mice such that the sperm from the stud males has enough time to complete fertilization, and that the pronuclei from both gametes will be visible for several hours after embryo isolation has occurred. This will provide a time
- 51. Oocyte Injection 41 window in which the embryos may be successfully injected with prepared DNA, before the pronuclei fuse and the membranes are no longer visible. To minimize the number of animals to be used for embryo donors, and to boost the efficiency of embryos recovered per donor, the technique of superovula- tion (7) is frequently employed. 1.2.1. Superovulation Superovulation is achieved by the injection of gonadotropins to stimulate and increase natural ovulation. This is most commonly done by administering pregnant mare’s serum gonadotropin (PMSG) to mimic the endogenous effects of follicle-stimulating hormone, followed by human chorionic gonadotropin (hCG) to mimic the effect of luteinizing hormone (LH). The importance of this hormone treatment is twofold: to increase the number of ovulations for each female and to control the timing of ovulation independent from the natural estrous cycle. It has been documented, however, that the administration of hor- mones to elicit superovulation does increase the rate at which there are chro- mosomal errors in the embryos obtained through this process (8). PMSG is most commonly supplied as a lyophilized powder and, for best results, should be stored at –80°C until ready for use. PMSG will remain stable as a powder but is very unstable once reconstituted and should only be resus- pended immediately before administration. For convenience, aliquots of PMSG can be stored at –20°C, but this can greatly decrease its efficacy over time. Most commonly, PMSG is administered intraperitoneally at a dose of 5 IU (international units)/mouse. For administration, PMSG is resuspended in ster- ile water or 0.9% NaCl. The second gonadotropin administered is hCG. It is given to induce the rup- ture of the mature follicle and is typically administered 42–48 h following administration of PMSG. hCG is also typically supplied as a lyophilized pow- der and is far more stable than PMSG when reconstituted. For administration, hCG is resuspended at 500 IU/mL in sterile water or 0.9% NaCl and is divided into 100-μL aliquots. These can be stored for approx 1 mo at –20°C. For administration of hCG, these aliquots may be diluted in 1 mL of sterile water or 0.9% NaCl for a final concentration of 50 IU, and then 0.1 mL is injected into each animal at a dose of 5 IU/mouse. 1.2.2. Mating Mice The mice to be used for the generation of fertilized embryos for microinjec- tion and those for use as pseudopregnant embryo recipients should be mated on the day before microinjection is to occur. This will give rise to fertilized embryos that are 0.5 dp.c., and recipients that are timed appropriately to receive those that have been successfully injected.
- 52. 42 Brown and Corbin The mice should be mated in the late afternoon following the administration of hCG, to ensure that the majority of the mice mate during the dark period of the room’s light cycle. Mice that mate earlier may yield a much higher number of fertilized embryos that have already fused their pronuclei and lost any vis- ible pronuclear membrane, or have advanced past the first cellular division to the two-cell stage of development. Neither of these embryos can be success- fully microinjected, because the placement of the injection needle to deliver the DNA construct cannot be determined. Any mice to be used as pseudopreg- nant embryo recipients for these injected eggs should be mated in synchrony with these mice also. The stud males’ ability to plug female mice should be carefully monitored and tracked to ensure maximum mating efficiency and a high yield of fertilized eggs harvested per superovulated female. 1.2.3. Light Cycle Several factors control the reproductive performance of superovulated females and stud males. Breeding conditions such as light cycle and timing must be carefully controlled and regulated. The time of administration of PMSG and hCG and the light–dark cycle of the animal facility is critical to the synchronous development of the eggs and the number harvested from the superovulated female. If female mice are ordered from an outside supplier, they should be allowed approx 1 wk to adjust to the animal room’s light–dark cycle before administering superovulatory hormones. This will also synchro- nize the endogenous release of LH stimulated by the PMSG injection. Control- ling the release of endogenous LH is important because the hCG must be administered prior to the natural LH surge in order to precisely control ovula- tion. The endogenous LH release is controlled by the light–dark cycle and gen- erally occurs approx 18 h following the second dark period after administration of PMSG. A typical injection and mating schedule is as follows: For a 12-h light–dark cycle set at 6 AM lights on and 6 PM lights off, PMSG is given at approx 12:00 noon. hCG should then be given 46–48 h later at approx 11:00 AM to 12:00 noon. This will allow several hours before endogenous release of LH. Following the administration of hCG, mice should be monogamously mated one female to one stud male. The female is then carefully checked the following morning for the presence of a copulatory plug. 1.3. Microinjection of DNA It is critically important that the DNA to be used for microinjection is as clean as and of the highest quality possible. Vector sequences can significantly alter the expression of transgenes (mechanism unclear) and must be separated
- 53. Oocyte Injection 43 from the insert by restriction digest. DNA that contains particulate matter will clog the injection needle and slow the operator and if fine enough will pass into the cell with possibly deleterious effects. Additionally, purification reagents that contaminate the sample to be microinjected are frequently toxic to the cell, and nuclear condensation or lysis will occur. Several protocols exist to purify injection-quality DNA, in print and also on the Internet. One such protocol is detailed in Subheading 2.3. (9). 1.4. Oviduct Transfer Following microinjection of foreign DNA, the manipulated embryos must be transferred to pseudopregnant recipient females. Embryos from the one-cell through to the morula stage (0.5–2.5 d p.c.) are transferred into the ampulla by oviduct transfer. Generally, microinjected eggs are transferred at the one- to two-cell stage. One-cell embryo transfer is best performed after allowing injected embryos a period in which to recover (1–3 h) in culture media, such as M16 (Sigma, St. Louis, MO) Z (see Note 5). This allows better evaluation of the cells’ survival and easier identification of viable cells for transfer. Eggs also can be incubated in culture overnight and allowed to develop to the two- cell stage. This gives an even better indication of cell viability because only healthy, undamaged cells will divide to the two-cell stage. However, it is best to minimize the time in culture, so the increased confidence in transferring embryos at the two-cell stage must be balanced against the increased in vitro exposure. The number of eggs transferred to a recipient female also depends on whether the transfer is performed at the one- or two-cell stage. In general, approx 20–30 one-cell eggs can be delivered into each recipient mouse. This number can be reduced to approx 15–20 for transfer of two-cell embryos. These numbers generally will produce litters of approx 5–10 pups, considering that 50–75% of transferred embryos will develop to term for one- and two-cell embryos, respectively. 1.5. Alternate Technologies and Strategies In recent years, new methods of transgenic production have been developed. These alternate strategies may allow some benefits over the established “stan- dard” technique of transgenic production outlined in previous chapters. Although none of the techniques described yield higher efficiencies (indeed they often have lower yields per treated embryos), these do afford other tan- gible benefits that may prove valuable. Injection into fertilized embryos normally requires placement of the DNA insert into the pronuclear envelope, a skill that may take many months to acquire and gain proficiency. A technique has been described in which the DNA for injection is complexed with a polylysine mixture (13), enabling the
- 54. 44 Brown and Corbin generation of transgenic mice by injection into the cytoplasm. Clearly, such a technique has value in the training of a new microinjection operator, whereby even unsuccessful injections where the pronucleus is not injected within the membrane but beside it may yield viable founder animals. In such a manner, training periods would still be able to contribute meaningfully to the injection projects at hand. A methodology has been described whereby previously vitrified embryos can be injected (14) into the pronucleus by standard means and give rise to transgenic founders at a similar rate to nonvitrified embryos. This has the advantage that numbers of embryos can be obtained either over time or when it is convenient to do so and be maintained in a cryopreserved state. When micro- injection needs to be performed, sufficient embryos for a day’s injection are thawed and injected as normal. This has the advantage of limiting the costs of maintaining a large production colony of mice, although it lends itself more to facilities where microinjection is a relatively infrequent activity. Additionally, the use of adenoviral vector delivery has been used to gener- ate transgenic mice (15). These viruses are replication defective and are used to infect one-cell fertile embryos that have had the zona pellucida removed. This strategy is particularly relevant to many researchers because it eliminates the requirement for a considerable component of the standard equipment used in transgenic animal production, thus representing a considerable saving in startup costs. An additional benefit is that this system delivers a single copy of the gene of interest rather than the multiple copies that the “standard” tech- nique often imparts by concatomer formation (16). This allows the evaluation of the insert without the complication of gene dosage. 2. Materials 2.1. Mouse Production Colony 2.1.1. Vasectomy 1. One pair of curved serrated forceps. 2. One pair of straight serrated forceps. 3. Two pairs of watchmaker’s forceps. 4. One pair of 4-in. sharp/sharp dissecting scissors. 5. One pair of Autoclip applicators. 6. Autoclips. 7. 4/0 Silk suture, with curved needle swagged on. 8. One-half of a plastic 10-cm Petri dish half full of 70% EtOH (ethanol). 2.1.2. Anesthetic for Surgery 2,2,2 Tribromoethanol (avertin) is found to be quite effective. For the method of preparation, see ref. 3. This agent should be stored wrapped in tin
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