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John M. Walker, SERIES EDITOR 518. Microinjection: Methods and Applications, edited by David J. Carroll, 2009
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Wnt Signaling, Volume 1: Pathway Methods and Mammalian Models, edited by Elizabeth Vincan, 2008
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Angiogenesis Protocols: Second Edition, edited by Stewart Martin and Cliff Murray, 2008
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489. Dynamic Brain Imaging: Methods and Protocols, edited by Fahmeed Hyder, 2009
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485. HIV Protocols: Methods and Protocols, edited by Vinayaka R. Prasad and Ganjam V. Kalpana, 2009
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480. Macromolecular Drug Delivery: Methods and Protocols, edited by Mattias Belting, 2008
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479. Plant Signal Transduction: Methods and Protocols, edited by Thomas Pfannschmidt, 2008
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478. Transgenic Wheat, Barley and Oats: Production and Characterization Protocols, edited by Huw D. Jones and Peter R. Shewry, 2008
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477. Advanced Protocols in Oxidative Stress I, edited by Donald Armstrong, 2008
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476. Redox-Mediated Signal Transduction: Methods and Protocols, edited by John T. Hancock, 2008
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475. Cell Fusion: Overviews and Methods, edited by Elizabeth H. Chen, 2008
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METHODS
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M O L E C U L A R B I O L O G Y TM
Microinjection Methods and Applications Edited by
David J. Carroll Florida Institute of Technology, Melbourne, FL, USA
Editor David J. Carroll Florida Institute of Technology Melbourne, FL USA
[email protected] Series Editor John M. Walker University of Hertfordshire Hatfield, Hert. UK
ISSN: 1064-3745 ISBN: 978-1-58829-884-3 DOI 10.1007/978-1-59745-202-1
e-ISSN: 1940-6029 e-ISBN: 978-1-59745-202-1
Library of Congress Control Number: 2008938642 # Humana Press, a part of Springer ScienceþBusiness Media, LLC 2009 All rights reserved. This work may not be translated or copied in whole or in part without the written permission of the publisher (Humana Press, c/o Springer ScienceþBusiness Media, LLC, 233 Spring Street, New York, NY 10013, USA), except for brief excerpts in connection with reviews or scholarly analysis. Use in connection with any form of information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed is forbidden. The use in this publication of trade names, trademarks, service marks, and similar terms, even if they are not identified as such, is not to be taken as an expression of opinion as to whether or not they are subject to proprietary rights. Cover illustration: The image shows how a holding pipet is used to immobilize a zebrafish oocyte for microinjection. A typical holding pipet produced by flame polishing of a glass capillary is used to pick up and immobilize an unfertilized oocyte. Magnification is indicated by the bar which represents 100mm. Photo credit: William H. Kinsey. Printed on acid-free paper springer.com
Preface David J. Carroll Abstract Cellular microinjection techniques have developed over the last century along with the evolution of other biological fields. For example, developmental biologists have used a variety of microinjection techniques to transfer cytoplasm between cells, inject antibodies and peptides, and express foreign genes in specific tissues to advance their understanding of cell specification and determination. Molecular biologists finely craft glass microneedles and tools for the manipulation of cells in order to study gene expression and communication between cells. As these techniques have matured, microinjection has become accessible and, thus, exploited by a larger segment of the scientific and medical community. As more and more information is gathered from the various genome projects, demand grows for methods to validate these new data. Microinjection can help address this need. In particular, microinjection has proven valuable for the confirmation and extension of in vitro results in an in vivo setting – the living cell. The technique has also found a home in the clinical setting, most obviously within the community of fertility specialists for in vitro fertilization methods and for those excited about the possibility of therapeutic cloning. This book explores the use of microinjection for a wide range of scientific uses. There are special considerations for each application of microinjection – whether it is the injection of antibodies, fusion proteins, DNA, and in vitro-synthesized RNA or the production of transgenic animals. It is hoped that these methods will be of interest for all biologists for use in the research laboratory, as well as for clinicians interested in applying this powerful method for treatment in the clinic.
1. A Brief History of Microinjection The technique of microinjection was born of necessity and owes its history to a combination of fields. Credit for the initial description of a coherent microinjection technique could be given to Marshall Barber, who developed methods for producing fine glass capillary pipettes for isolating and manipulating single bacterial cells (1,2), or to the embryologist Laurent Chabry, credited with developing the glass microcapillary and micromanipulator as tools for his studies on teratology in ascidian blastomeres (3,4). Barber incorporated techniques into his injection method that are still used today, including the first use of mercury for controlling the movement of small volumes of fluid (see below) and the use of a second pipette to hold the cells as the injection is completed (5,6). The technique of microinjection moved from being a useful method practiced by a few resourceful scientists to an exciting mainstream application when Gurdon and colleagues demonstrated that
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purified mRNA from one cell could be injected into the cytoplasm of another cell and actually get translated into protein (7). This was remarkable for several reasons: (1) the mRNA was stable, at least stable enough that it was capable of directing the synthesis of detectable levels of protein; (2) no special factors, other than the exogenous mRNA itself, were required for this to work; and (3) success was independent of the cell type of the donor RNA. In fact, species differences do not appear to matter, as they were able to obtain the synthesis of rabbit hemoglobin in Xenopus laevis oocytes. Since then, the method of expressing protein from microinjected mRNA has been used to great advantage in studies for developmental biology, neurobiology, cell biology, and signal transduction, just to name a few. An excellent overview of this method is given by Douglas Melton (8), in which he explains why the oocyte (and the Xenopus oocyte, in particular) provides the perfect system for this to work, along with describing details of the current methods at that time. One great advantage of this system, compared to translation in vitro or by induced expression in bacteria, is that the mRNA is placed in a living eukaryotic cell so the protein is processed properly and subject to post-translational modification. Of course, the technique is not only useful for microinjecting RNA into Xenopus oocytes. It has been utilized for transferring cytoplasm from one cell to another, such as was done in the experiments that led to the discovery that maturation promoting factor (MPF) was neither species nor cell-type specific (9–11). This fundamental work provided the groundwork for the eventual discovery of cyclins and cyclin-dependent kinases (12,13). Proteins have been injected into cells for the study of cell structure and function. One of the first examples of this utility was the microinjection of fluorescently labeled -actinin into living fibroblasts, allowing the visualization of this molecule integrating dynamically into the cytoskeleton (14). Of course, the concept of following labeled proteins in a living cell has since exploded with the availability of green fluorescent protein (GFP) vectors and protocols (15,16). One tremendous advantage of microinjection, when compared to other methods of introducing material into cells, such as electroporation or chemical membrane permeabilization, is the ability to be quantitative. Methods for quantitative microinjection have developed over the years, but began in earnest with a study by Hiramoto on the process of fertilization (17). In that study, live spermatozoa were microinjected into sea urchin eggs to develop an assay for the identification of a ‘‘substance or substances which trigger the train of fertilization reactions in the egg’’. No such substance was identified in that study (see Chapter 2), but methods to precisely control and quantify the volume of the injection using a mercury-filled needle connected to a screw-controlled
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syringe were described. Hiramoto’s basic method has been further refined (18–20), but is still being used today. This book provides methods of microinjection coupled with modern molecular techniques, such as RNAi, morpholino antisense oligonucleotides, GFP expression, or the production of transgenic cells or animals, for example. However, the book also revisits classic uses of microinjection, such as mRNA expression or nuclear transfer, with modern twists.
2. Book Content The classic technique of microinjecting Xenopus zygotes with mRNA prepared in vitro is revisited in Chapter 1 with a focus on studying proteins involved in the cell cycle. A discussion of the use of microinjection compared to other methods of manipulating proteins in a living cell is given. The chapter also provides consideration of choosing the appropriate plasmid vector with an eye toward downstream analysis of the effectiveness of the protein expression. This discussion should be very useful for investigators considering the use of mRNA microinjection for the first time. In Chapter 2, a very clever method using a luciferase chimera to visualize the expression of a protein, PLC, that causes calcium release and egg activation during fertilization in mammals is described (21). Methods are given for producing the luciferaselabeled cRNA, along with techniques for assessing the expression of the fusion protein by microscopy, which allows simultaneous imaging of fluorescent indicators (e.g., a Ca2+ indicator), or by luminometer for quantification of luciferase expression. This method could easily be applied to other molecules in any cell type that will express exogenous RNA. In particular, it promises to be useful for combining detection of low levels of protein expression with quantification of those molecules from a reasonable number of cells. In Chapter 3, a very straightforward technique for using antisense morpholino oligos to specifically remove 14-3-3 proteins in Xenopus laevis is described. Morpholino oligos are nonionic DNA analogs that possess an altered phosphodiester backbone. This apparently makes them more resistant to nucleases and, because they are not charged, less likely to interact nonspecifically with proteins (22). This technique complements their earlier work in which the 14-3-3 proteins were studied using peptide inhibitors or dominant-interfering GST fusion proteins (23). Their method utilizes morpholinos that target the initiation codon and the 22 ribonucleotides immediately downstream,
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which may produce more effective inhibition. This chapter outlines methods for (1) procuring the Xenopus sperm and eggs, (2) setting up and using a pressure microinjection apparatus with the Xenopus one- or two-cell embryo, (3) for analyzing and quantifying protein levels by western blotting, and (4) for analyzing the phenotypic effects of the morpholino injections. An alternate strategy for morpholino design that targets the 50 untranslated region of the target gene is also discussed. Methods for microinjecting peptides and fusion proteins into Xenopus laevis oocytes are given in Chapter 4. Following a very clear description of ovary dissection and oocyte procurement by collagenase treatment, the procedure for performing rapid microinjections into the Xenopus oocyte is given. Accompanied by excellent photographs, this chapter explains the process in detail from start to finish. For example, in the text and in the Notes section, suggestions are given regarding the exact size for the tip of the microinjection needle and when the needle should be changed, the maximum volume and concentration of protein that should be injected into an individual oocyte, and how to deal with multiple injections into the same oocyte. This type of detail fills the chapter and will help other investigators maximize success when they adopt this method. A method to combine microinjection with western blot analysis is described in Chapter 5. In this procedure, single oocytes are microinjected with a pharmacological inhibitor of RAS and then assayed for the presence of phosphorylated mitogen-activated protein kinase (pMAPK), which indicates an active enzyme, by immunoblotting. Thus far, this has been applied only for the analysis of the pMAPK during oocyte maturation and fertilization in starfish oocytes. However, it should also be useful in other large cells (such as Xenopus oocytes) and, as detection methods improve, it could easily be adapted to the analysis of other proteins in other systems. Analysis of single cells should prove useful because it eliminates the variability inherent to the analysis of cell populations. This book considers the use of many different model systems, including the zebrafish Danio rerio. The zebrafish has proven extremely useful for experimental study because it is amenable to genetic studies, the embryo develops rapidly, and it is optically clear (you can see inside). The zebrafish zygote and early embryo have been utilized as a system for microinjection because of the large size of the eggs and the ease with which the adults are maintained. However, for a variety of reasons it would be advantageous to inject the mature egg prior to fertilization and this has proven difficult. In Chapter 6, a practical method for microinjection, and subsequent insemination, of the unfertilized zebrafish egg is reported. This will open up more opportunities for exploiting an already useful model system.
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In the first method (Chapter 7) to deal with a ‘non-gamete’ system, techniques for antibody microinjection and oligofectamine transfection of RNAi for analysis of protein function in living tissue culture cells are compared. The chapter is very detailed which should allow these methods to be adapted to many different situations. The chapter also features a very extensive and useful notes section, detailing specifics of each of the methods. Methods for developing recombinant cells lines by microinjection into the nucleus are presented in Chapter 8. This very interesting article demonstrates that introduction of plasmid DNA for the GFP, in several cell types, leads to stable transduction as assayed by flow cytometry of GFP fluorescence up to 1 month after microinjection! The chapter details methods that could be adapted to virtually any cell type and for any DNA. Conditions for optimizing the production of the transformed cells lines are tested in this chapter and detailed notes are given to help any investigator interested in attempting this method. Different experimental methods, including transposons, I-SceI meganuclease, and direct injection of linearized DNA, have been used to produce transgenic Xenopus for the study of a variety of problems (24–26). However, these methods rely upon random insertion into the Xenopus genome or produce multiple copies. In Chapter 9, a technique is described for the targeted insertion of a single copy of the gene of interest into Xenopus laevis using phiC31 integrase. By incorporating insulator sequences into the plasmid design, they improve expression from the reporter gene making this method extremely useful for anyone wishing to express a transgene at approximately endogenous levels. Transgenics are also the topic of Chapter 10. A method for producing transgenic Caenorhabditis elegans by microinjecting DNA directly into the hermaphrodite gonad is provided. For those not familiar with C. elegans, this chapter provides an excellent introduction to this powerful system. It recounts the basic reproductive biology of the worm and includes a discussion of applications for microinjection. Techniques for cultivating the worms and special hints for maximizing success of microinjection are explained in detail. One of the great advantages of using microinjection to introduce molecules into living cells is the ability to perform quantitative experiments. Chapter 11 gives detailed methods for the quantitative microinjection of picoliter quantities into mouse oocytes and eggs in dishes on an inverted microscope. The technique is described in wonderful detail and covers all aspects from making the glass bottom dishes that hold the oocytes to the injection process itself. There are several items of note in the chapter. I particularly enjoyed learning how to use an old record player to construct a beveler capable of producing a nice 1–2 mm tip. But then, what will I do with my old Pink Floyd records?
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For precision, the microinjection needles need to include a mechanism for controlling the rate of injection. Two methods are given in Chapter 11 to achieve this: (1) using mercury to backfill the micropipette and relying upon the mercury to transduce and control pressure and (2) constructing a microneedle with a constriction near the tip which reduces fluid movement and provides fine control of the injection. Both types of microinjection needles can be used in the same overall system. A similar microinjection system is described in Chapter 12, with modifications that allow for injection of the mouse oocyte within the complete follicle. Because the mouse oocyte exists within the follicle when in the ovary, it is intimately associated with the surrounding follicle cells until after ovulation. This relatively new method allows for the microinjection to occur while the oocyte is cultured within the intact follicle under more physiologically relevant conditions. By microinjecting the follicle-enclosed oocytes, this group has been able to discover the mechanism(s) that maintain the immature oocyte arrested in meiosis and also to begin exploring the signaling mechanisms that are responsible for the reinitiation of meiosis (27–29). The technique is quite revolutionary and the concept of maintaining the proper physiological environment, as much as possible, is one that is applicable to all situations. In this chapter, you will also learn what type of music is most enjoyed by mouse oocytes. In some cases, microinjection would be very useful but not considered because of the perceived difficulty of injecting sufficient numbers of cells for further analysis. Chapter 13 describes a pressure-based method of injecting zygotes of the sea urchin Paracentrotus lividus. Using this method, up to several hundred embryos can be injected in a single session. In addition to the microinjection method, this chapter also describes experimental methods for perturbing gene function by either (1) producing and microinjecting synthetic mRNA produced in vitro, or (2) preparing linear DNA amplified by PCR for direct microinjection into the embryo. The advantages and disadvantages of these two different methods are discussed. The final two chapters describe practical uses of the technique of microinjection. Methods for imaging human gametes and zygotes after intracytoplasmic sperm injection (ICSI) are presented in Chapter 14. The techniques focus on the identification of cytoskeletal elements and the role these components play during fertilization. Procedures are given in great detail for the removal of follicle cells and the zona pellucida, and fixation procedures optimized for fluorescence immunocytochemistry and for examination by conventional electron microscopy and ultrastructural immunolocalization. Chapter 15 presents a method for somatic cell nuclear transfer (SCNT) in the mouse. This article directly addresses the fact
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that cloning by SCNT has always been difficult and inefficient (30, 31). In the protocol described here, donor nuclei from cumulus cells are injected directly into mouse oocyte. While this has been accomplished before, the Kishigami and Wakayama protocol results in a twofold to fivefold improvement in embryo developmental rates by the inclusion of trichostatin A, a histone deacetylase inhibitor. The TSA may ‘reprogram’ the somatic cell nuclei, making them more amenable to the early developmental program. Also significant in Chapter 15 is a description of the use of a piezo-actuated micromanipulator that allows the use of larger microinjection needle tips with less damage to the oocyte.
3. Conclusion The methods described in this book should allow any lab to incorporate the technique of microinjection into their experimental repertoire. Whether DNA, RNA, or protein is the molecule of interest, microinjection provides a mean of studying function within the context of the living cell. The technology is remarkably accessible and relatively inexpensive, while the possibilities are virtually endless.
Acknowledgments I thank Dr. Laurinda Jaffe of the University of Connecticut Health Center for introducing me to microinjection and other fun things; and to Dr. John Walker of the University of Hertfordshire for his patience and guidance during the development of this book.
References 1. Barber, M. (1904) A new method of isolating microorganisms. J. Kans. Med. Soc. 4, 489–494. 2. Barber, M.A. (1911) A technique for the inoculation of bacteria and other substances into living cells. J. Infect. Dis. 8, 348–360. 3. Chabry, L. (1887) Contribution a l’embryologie normal et teratologique des Ascidiens simples. Jour. de l’Anat. et de Physiol. 25, 167.
4. Fischer, J-L. (1990) Experimental embryology in France. Int. J. Dev. Biol. 34, 11–23. 5. Barber, M.A. (1914) The pipette method in the isolation of single microorganisms and in the inoculation of substances into living cells. Philippine J. Sci. B. 9, 307–360. 6. Korzh, V. and Strhle, U. (2002) Marshall Barber and the century of microinjection: from cloning of bacteria to cloning of everything. Differentiation 70, 221–226.
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7. Lane, C.D., Marbaix, G., and Gurdon, J.B. (1971) Rabbit haemoglobin synthesis in frog cells: the translation of reticulocyte 9S RNA in frog oocytes. J. Mol. Biol. 61, 73–91. 8. Melton, D.A. (1987) Translation of messenger RNA in injected frog oocytes. Meth. Enzymol. 152, 288–296. 9. Masui, Y., and Markert, C.L. (1971) Cytoplasmic control of nuclear behavior during meiotic maturation of frog oocytes. J. Exp. Zool. 177, 129–145. 10. Reynhout J.K. and Smith L.D. (1974) Studies on the appearance and nature of a maturation-inducing factor in the cytoplasm of amphibian oocytes exposed to progesterone. Dev. Biol. 38, 394–400. 11. Gurdon, J.B. (1968) Changes in somatic cell nuclei inserted into growing and maturing amphibian oocytes. J. Embryol. Exp. Morphol. 20, 401–414. 12. Evans, T., Rosenthal, E.T., Youngblom, J., Distel, D., Hunt, T. (1983) Cyclin: a protein specified by maternal mRNA in sea urchin eggs that is destroyed at each cleavage division. Cell 33, 389–396. 13. Nurse, P., Thuriaux, P., and Nasmyth, K. (1976) Genetic control of the cell division cycle in the fission yeast Schizosaccharomyces pombe. Mol. Gen. Genet. 146, 167–178. 14. Feramisco, J.R. (1979) Microinjection of fluorescently labeled a-actinin into living fibroblasts. Proc. Natl. Acad. Sci. USA. 76, 3967–3971. 15. Prasher, D.C., Eckenrode, V.K., Ward, W.W., Prendergast, F.G., Cormier, M.J. (1992) Primary structure of the Aequorea victoria greenfluorescent protein. Gene. 111, 229–233. 16. Shimomura, O. (2005) The discovery of aequorin and green fluorescent protein. J. Microsc. 217, 1–15. 17. Hiramoto, Y. (1962) Microinjection of the live spermatozoa into sea urchin eggs. Exp. Cell Res. 27, 416–426. 18. Kiehart, D.P. (1982) Microinjection of echinoderm eggs: apparatus and procedures. Methods Cell Biol. 25, 13–31. 19. Kishimoto, T. (1986) Microinjection and cytoplasmic transfer in starfish oocytes. Methods Cell Biol. 27, 379–394. 20. Jaffe, L.A., and Terasaki, M. (2004) Quantitative microinjection of oocytes, eggs, and embryos. Methods Cell Biol. 74, 219–242.
21. Saunders, C.M., Larman, M.G., Parrington, J., Cox, L.J., Royse, J., Blayney, L.M., Swann, K., and Lai F.A. (2002) PLC: a sperm-specific trigger of Ca2+ oscillations in eggs and embryo development. Development 129, 3533–3544. 22. Corey, D.R., and Adams, J.M. (2001) Morpholino antisense oligonucleotides: tools for investigating vertebrate development. Genome Biol. 2(5), reviews1015.1–1015.3. 23. Wu, C. and Muslin, A.J. (2002) Role of 143-3 proteins in early Xenopus development. Mech. Dev. 119, 45–54. 24. Yergeau, D.A., and Mead, P.E. (2007) Manipulating the Xenopus genome with transposable elements. Genome Biol. 8 (Suppl 1), S11. 25. Pan, F.C., Chen, Y., Loeber, J., Henningfeld, K., Pieler, T. (2006) I-SceI meganuclease-mediated transgenesis in Xenopus. Dev Dyn. 235, 247–252. 26. Etkin, L., Pearman, B., Roberts, M., Bektesh, S.L. (1984) Replication, integration and expression of exogenous DNA injected into fertilized eggs of Xenopus laevis. Differentiation 26, 194–202. 27. Mehlmann, L.M., Jones, T.L., Jaffe, L.A. (2002) Meiotic arrest in the mouse follicle maintained by a Gs protein in the oocyte. Science 297, 1343–1345. 28. Mehlmann, L.M., Saeki, Y., Tanaka, S., Brennan, T.J., Evsikov, A.V., Pendola, F.L., Knowles, B.B., Eppig, J.J., Jaffe, L.A. (2004) The Gs-linked receptor GPR3 maintains meiotic arrest in mammalian oocytes. Science 306, 1947–1950. 29. Freudzon, L., Norris, R.P., Hand, A.R., Tanaka, S., Saeki, Y., Jones, T.L., Rasenick, M.M., Berlot, C.H., Mehlmann, L.M., Jaffe, L.A. (2005) Regulation of meiotic prophase arrest in mouse oocytes by GPR3, a constitutive activator of the Gs G protein. J. Cell Biol. 171, 255–265. 30. Tian, X.C., Kubota, C., Enright, B., Yang, X. (2003) Cloning animals by somatic cell nuclear transfer – biological factors. Reprod Biol Endocrinol. 1, 98. 31. Campbell, K.H., Fisher, P., Chen, W.C., Choi, I., Kelly, R.D., Lee, J.H., Xhu, J. (2007) Somatic cell nuclear transfer: Past, present and future perspectives. Theriogenology. 68 Suppl 1, S214–S231.
Contents Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . v Contributors. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . xv
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Expression of Exogenous mRNA in Xenopus laevis Embryos for the Study of Cell Cycle Regulation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1 Jill C. Sible and Brian N. Wroble Use of Luciferase Chimaera to Monitor PLC Expression in Mouse Eggs . . . . . . . . . 17 Karl Swann, Karen Campbell, Yuansong Yu, Christopher Saunders and F. Anthony Lai Analysis of 14-3-3 Family Member Function in Xenopus Embryos by Microinjection of Antisense Morpholino Oligos . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 31 Jeffrey M. C. Lau and Anthony J. Muslin A Microinjectable Biological System, the Xenopus Oocyte, as an Approach to Understanding Signal Transduction Protein Function . . . . . . . . . . . . . . . . . . . . . . . . 43 Katia Cailliau and Edith Browaeys-Poly Combining Microinjection and Immunoblotting to Analyze MAP Kinase Phosphorylation in Single Starfish Oocytes and Eggs . . . . . . . . . . . . . . . . . . . . . . . . . 57 David J. Carroll and Wei Hua Analysis of Signaling Pathways in Zebrafish Development by Microinjection . . . . . . . 67 William H. Kinsey Protein Inhibition by Microinjection and RNA-Mediated Interference in Tissue Culture Cells: Complementary Approaches to Study Protein Function . . . . . . . . . . . . 77 Jane R. Stout, Rania S. Rizk, and Claire E. Walczak DNA Delivery by Microinjection for the Generation of Recombinant Mammalian Cell Lines . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 99 Sebastien Chenuet, Madiha Derouazi, David Hacker and Florian Wurm Bacteriophage fC31 Integrase Mediated Transgenesis in Xenopus laevis for Protein Expression at Endogenous Levels . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 113 Bryan G. Allen and Daniel L. Weeks Germline Transformation of Caenorhabditis elegans by Injection . . . . . . . . . . . . . . . 123 Pavan Kadandale, Indrani Chatterjee and Andrew Singson Quantitative Microinjection of Mouse Oocytes and Eggs . . . . . . . . . . . . . . . . . . . . . 135 Douglas Kline Microinjection of Follicle-Enclosed Mouse Oocytes . . . . . . . . . . . . . . . . . . . . . . . . . 157 Laurinda A. Jaffe, Rachael P. Norris, Marina Freudzon, William J. Ratzan, and Lisa M. Mehlmann Functional Studies of Regulatory Genes in the Sea Urchin Embryo . . . . . . . . . . . . . 175 Vincenzo Cavalieri, Maria Di Bernardo, and Giovanni Spinelli
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Exploring the Cytoskeleton During Intracytoplasmic Sperm Injection in Humans . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 189 Vanesa Y. Rawe and He´ctor Chemes Somatic Cell Nuclear Transfer in the Mouse . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 207 Satoshi Kishigami and Teruhiko Wakayama
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Index. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 219
Contributors BRYAN G. ALLEN Department of Biochemistry, University of Iowa, Iowa City, IA, USA EDITH BROWAEYS-POLY Universite´ des Sciences et Technologies de Lille, Laboratoire de Re´gulation des Signaux de Division, Villeneuve d’Ascq Cedex, France KATIA CAILLIAU Universite´ des Sciences et Technologies de Lille, Laboratoire de Re´gulation des Signaux de Division, Villeneuve d’Ascq Cedex, France KAREN CAMPBELL Department of Obstetrics and Gynaecology, School of Medicine, Cardiff University, Cardiff, UK DAVID J. CARROLL Department of Biological Sciences, Florida Institute of Technology, Melbourne, FL, USA VINCENZO CAVALIERI Dipartimento di Biologia Cellulare e dello Sviluppo ‘‘A. Monroy’’, Universita` di Palermo, Palermo, Italy INDRANI CHATTERJEE Waksman Institute, Rutgers University, Piscataway, NJ, USA HE´CTOR CHEMES CEDIE, Laboratorio de Fisiologı´ y Patologı´a Testicular, Hospital de Nin ˜ os ‘Ricardo Gutie´rrez’, Buenos Aires, Argentina SEBASTIEN CHENUET E´cole Polytechnique Fe´derale de Lausanne, EPFL-SV-IBI-LBTC, Lausanne, Switzerland MADIHA DEROUAZI E´cole Polytechnique Fe´derale de Lausanne, EPFL-SV-IBI-LBTC, Lausanne, Switzerland MARIA DI BERNARDO Istituto di Biomedicina e Immunologia Molecolare ‘‘A. Monroy’’, Consiglio Nazionale delle Ricerche, Palermo, Italy MARINA FREUDZON Department of Cell Biology, University of Connecticut Health Center, Farmington, CT, USA DAVID HACKER E´cole Polytechnique Fe´derale de Lausanne, EPFL-SV-IBI-LBTC, Lausanne, Switzerland WEI HUA College of Aqua-life Science and Technology, Shanghai Fisheries University, Shanghai, China LAURINDA A. JAFFE Department of Cell Biology, University of Connecticut Health Center, Farmington, CT, USA PAVAN KADANDALE Waksman Institute, Rutgers University, Piscataway, NJ, USA WILLIAM H. KINSEY Department of Anatomy and Cell Biology, University of Kansas Medical Center, Kansas City, KS, USA SATOSHI KISHIGAMI RIKEN, Center for Developmental Biology, Kobe, Japan DOUGLAS KLINE Department of Biological Sciences, Kent State University, Kent, OH, USA F. ANTHONY LAI Cell Signaling Laboratory, Wales Heart Research Institute, School of Medicine, Cardiff University, Cardiff, UK JEFFREY M.C. LAU Center for Cardiovascular Research, Department of Medicine, Department of Cell Biology & Physiology, Washington University School of Medicine, St. Louis, MO, USA
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LISA M. MEHLMANN Department of Cell Biology, University of Connecticut Health Center, Farmington, CT, USA ANTHONY J. MUSLIN Center for Cardiovascular Research, Department of Medicine, Department of Cell Biology & Physiology, Washington University School of Medicine, St. Louis, MO, USA RACHAEL P. NORRIS Department of Cell Biology, University of Connecticut Health Center, Farmington, CT, USA WILLIAM J. RATZAN Department of Cell Biology, University of Connecticut Health Center, Farmington, CT, USA VANESA Y. RAWE Centro de Estudios en Ginecologı´a y Reproducci´on (CEGyR), Buenos Aires, Argentina RANIA S. RIZK Department of Biology, Indiana University, Bloomington, IN, USA CHRISTOPHER SAUNDERS Cell Signaling Laboratory, Wales Heart Research Institute, School of Medicine, Cardiff University, Cardiff, UK JILL C. SIBLE Department of Biological Sciences, Virginia Polytechnic Institute and State University, Blacksburg, VA, USA ANDREW SINGSON Waksman Institute, Rutgers University, Piscataway, NJ, USA GIOVANNI SPINELLI Dipartimento di Biologia Cellulare e dello Sviluppo ‘‘A. Monroy’’, Universita` di Palermo, Palermo, Italy JANE R. STOUT Department of Biochemistry and Molecular Biology, Indiana University Medical Sciences, Bloomington, IN, USA KARL SWANN Department of Obstetrics and Gynaecology, School of Medicine, Cardiff University, Cardiff, UK TERUHIKO WAKAYAMA RIKEN, Center for Developmental Biology, Kobe, Japan CLAIRE E. WALCZAK Department of Biochemistry and Molecular Biology, Indiana University Medical Sciences, Bloomington, IN, USA DANIEL L. WEEKS Department of Biochemistry, University of Iowa, Iowa City, IA, USA BRIAN N. WROBLE Department of Biological Sciences, Virginia Polytechnic Institute and State University, Blacksburg, VA, USA FLORIAN WURM E´cole Polytechnique Fe´derale de Lausanne, EPFL-SV-IBI-LBTC, Lausanne, Switzerland YUANSONG YU Department of Obstetrics and Gynaecology, School of Medicine, Cardiff University, Cardiff, UK
Chapter 1 Expression of Exogenous mRNA in Xenopus laevis Embryos for the Study of Cell Cycle Regulation Jill C. Sible and Brian N. Wroble Abstract The microinjection of mRNA that is transcribed and capped in vitro into fertilized eggs and embryos of Xenopus laevis provides a powerful means for discovering the function of proteins during early development. Proteins may be overexpressed for a gain-of-function effect or exogenous protein function may be compromised by the microinjection of mRNA encoding ‘‘dominant-negative’’ proteins. This methodology is particularly suited for the investigation of the regulation of the cell cycle, checkpoints, and apoptosis in early development. Key words: Microinjection, mRNA, cell cycle, Xenopus laevis, early development, apoptosis, checkpoints, embryos, in vitro transcription.
1. Introduction 1.1. Applications
The microinjection of mRNA transcribed in vitro into oocytes, fertilized eggs, and embryonic cells from Xenopus laevis has become a classic methodology in developmental biology that takes advantage of the highly efficient translational capability of these cells. RNAs encoding membrane channels and transporters (other species) can be expressed in oocytes and eggs, providing a large surface area for patch clamping and other electrophysiological studies (1–3). Microinjection of mRNAs encoding X. laevis proteins enables one to determine the effect of ectopic expression of endogenous genes on early development. mRNAs of genes that have been mutated to encode ‘‘dominant-negative’’ proteins can be microinjected into embryos to assess the developmental
David J. Carroll (ed.), Microinjection: Methods and Applications, Vol. 518 Ó 2009 Humana Press, a part of Springer ScienceþBusiness Media, LLC DOI 10.1007/978-1-59745-202-1_1
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consequences of interfering with the function of a protein or pathway (4,5). Microinjection of mRNAs into X. laevis embryos provides a particularly powerful method for the investigation of cell cycle regulation. The first twelve cell cycles are driven exclusively by the translation of maternal cyclin mRNAs (6), and thus are amenable to manipulation by exogenous mRNA. After the twelfth cell cycle, the midblastula transition (MBT) begins, and cell cycle becomes lengthened and asynchronous (7). The embryo also becomes transcriptionally active at the MBT (8). Many maternal mRNAs that regulate the cell cycle are degraded and replaced by zygotic isoforms (9–11). Exogenous mRNAs encoding cell cycle-related proteins often alter the timing of cleavage cycles, the onset of the MBT, or both. The MBT also delineates a remodeling of the cell cycle with respect to cell cycle checkpoints. Prior to the MBT, embryos do not arrest cleavage divisions in response to damaged or unreplicated DNA, but rather undergo a maternally regulated program of apoptosis during early gastrulation (12–14). After the MBT, cell cycle checkpoints become operational, and damaged or unreplicated DNA triggers cell cycle arrest rather than apoptosis. Thus, the early embryo of X. laevis provides a dynamic system of cell cycle remodeling (15) that can be readily manipulated by microinjection of mRNA transcribed in vitro. 1.2. Relationship to Other Methods to Manipulate Gene Expression in X. laevis
In recent years, the suite of tools to manipulate gene expression in X. laevis and other externally developing organisms has been enhanced with new technologies. Notably, the development of transgenesis methodologies for X. laevis (16) and X. tropicalis (17) enables researchers to introduce genes under the control of specific promoters stably into the genome. Antisense morpholinos can be microinjected into the embryo to block translation of specific mRNAs through the late embryonic stages (18). Nonetheless, the more classic methodology of expressing genes by the microinjection of mRNAs transcribed in vitro maintains its value in the study of early embryogenesis. Because X. laevis embryos do not transcribe their genome until the MBT (cell cycle 12, approximately 6 h post-fertilization), transgenes will not be expressed before the MBT. Therefore, early developmental events including cell cycle remodeling and activation of the maternal program of apoptosis will not be affected by transgenesis, but they can be altered by microinjection of mRNAs, which are efficiently translated in the early embryo. Antisense morpholinos will block translation of maternal mRNAs but will not affect maternally supplied proteins, many of which regulate the cell cycle and apoptosis. In addition to being particularly suited for manipulation of gene expression during the earliest stages of development, microinjection of mRNAs is a relatively simple and inexpensive technique.
Expression of Exogenous mRNA in Xenopus
1.3. Summary
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Investigation of the regulation of cell cycle remodeling and other early developmental events can be facilitated by the microinjection of mRNA into early X. laevis embryos. Although the methodologies are straightforward, care should be taken to design plasmid constructions that encode protein tags and a poly-A tail, to generate high-quality RNA in an RNase-free environment, and to control for nonspecific effects of exogenous mRNA. By pairing this methodology with biochemical and morphological assays for cell cycle progression and apoptosis, we continue to build a systems-level view of the signaling networks that control early developmental events.
2. Materials 2.1. Template
1. pSP64poly(A) plasmid (Promega Corp., Madison, WI, USA) (see Note 1). 2. Restriction enzymes such as EcoRI and PvuII (see Note 2). 3. RNase-free phenol/chloroform (1:1). 4. 3-M sodium acetate. 5. 70% and 100% ethanol. 6. Tris-EDTA (TE) buffer: 10-mM Tris, 1-mM EDTA, pH 8.0.
2.2. RNA Preparation
1. mMessage mMachineTM (Ambion/Applied Biosystems, Inc., Austin, TX, USA). 2. Isopropanol. 3. Phenol: chloroform (1:1)
2.3. Fertilizing Eggs for Microinjection
1. Pregnant mare serum gonadotropin (PMSG; Calbiochem, catalog #367222). 2. Human chorionic gonadotropin (HCG; Sigma, catalog #C1063). 3. 0.1X MMR (0.5-mM HEPES, pH 7.8, 10-mM NaCl, 0.2-mM KCl, 0.1-mM MgSO4, 0.2-mM CaCl2, 0.01-mM EDTA).
2.4. Injecting the mRNA
1. Microcapillary tubes for making the microinjection needles. 2. Micropipet puller such as a Narishige PC-10 (Narishige Scientific Instrument Lab, Inc., Tokyo, Japan). 3. Sexually mature female adult Xenopus laevis. Commercial suppliers include (1) Xenopus I, Inc., Dexter, MI, USA; or (2) Xenopus Express Inc., Brooksville, FL, USA.
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2.5. Monitoring Expression of the mRNA
1. Mouse M2 Anti-FLAG monoclonal antibody (Sigma-Aldrich, St. Louis, MO, USA).
3. Methods 3.1. Preparing the Template
3.1.1. Constructing/ Selecting the Plasmid Vector Encoding the Experimental and Control mRNAs
Several considerations in the design and construction of the plasmid template should be made to optimize and verify the translatability of the mRNA to be injected. 1. The vector should have a bacteriophage T7, T3, or SP6 promoter upstream of a multicloning site (MCS). The pSP64poly(A) plasmid (Promega, catalog #P1241) is recommended because it encodes a poly-A tail downstream of the MCS followed by a unique restriction site (See Note 1). 2. The cDNA to be inserted into the MCS can be generated by excision from an existing plasmid followed by ligation into the in vitro transcription vector if compatible restriction sites exist. Alternatively, the desired portion of the cDNA (with or without UTRs) can be amplified by PCR. An advantage of PCR is that restriction sites and short epitope tags can be incorporated directly into the primers. 3. As an example of this strategy, the primer sequences and PCR conditions for generating a FLAG-tagged Chk2 cDNA to be cloned into pSP64polyA are shown in Fig. 1.1. A two-stage PCR was used because the 50 ends of each primer containing FLAG sequence and restriction sites will not anneal to the original cDNA template, necessitating a lower annealing temperature. After several rounds of amplification, enough product incorporating these primers is available as template and the annealing temperature can be raised to allow for a more efficient amplification. 4. The PCR product can then be digested with the appropriate restriction enzymes and cloned into digested pSP64polyA vector. Once clones with inserts have been selected, amplified, and sequenced, large-scale preparations of the plasmid should be made.
3.1.2. Restriction Digestion of the Template
1. Linearize 30-mg template plasmid by restriction digestion with an appropriate enzyme. 2. The restriction enzyme should not generate 30 overhangs. Try to choose a site that generates 50 overhangs (best choice) or blunt ends (OK). 3. A typical digestion reaction would include: a. 30-mg plasmid template b. 10-ml 10X buffer for restriction enzyme
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1 cycle @ 94°C for 2 min 5 cycles @ 94°C for 30 sec 52°C for 45 sec 68°C for 6 min 1 cycle @ 68°C for 20 min 20 cycles @ 94°C for 30 sec 62°C for 45 sec 68°C for 6 min 1 cycle @ 68°C for 20 min
Fig 1.1. PCR strategy for cloning X. laevis Chk2 cDNA into pSP64polyA. (A) Forward and reverse primers. On forward primer: 1, extra bases to facilitate restriction digestion of PCR product; 2, PstI site; 3, start codon; 4, Flag tag; 5, Chk2-specific sequence beginning after the start codon. On reverse primer: 6, extra bases to facilitate restriction digestion of PCR product, BamHI site; 8, reverse complement of stop codon; 9, reverse complement of the end of the Chk2 ORF. (B) Two-stage PCR program for amplifying Chk2 from cDNA clone using above primers.
c. 2-ml restriction enzyme d. H2O to 100-ml total volume 4. Incubate at 37°C for 5 h to overnight. 5. Digestion should be to completion because undigested plasmid can serve as template for transcription of concatamers of the entire plasmid. 6. To test the reaction for completion of digestion, remove 1 ml of the reaction and resolve by agarose gel electrophoresis alongside the same concentration of uncut plasmid. Linearized plasmid should resolve as a single discrete band that typically migrates slower than uncut plasmid. 7. After digestion, heat the reaction mixture at 65°C for 15 min to inactivate it. Extract with an equal volume of phenol/ chloroform (RNase-free). 8. From this point on, use only RNase-free reagents and plasticware and practice good RNA technique. 9. Precipitate the DNA with 1/10 volume 3-M sodium acetate and 2 volumes ethanol (EtOH) at –20°C overnight or –80°C for at least 15 min. 10. Centrifuge the DNA at 16,000 g for 15–30 min at 4°C, remove the 100% EtOH, wash the pellet with 70% EtOH (for RNA), centrifuge, remove the EtOH and air-dry the pellet.
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11. Resuspend in 30-mL TE (for RNA). 12. Take 1 ml and determine the concentration by spectrophotometry. 13. Dilute the DNA to 1 mg/ml. Store unused portion at 4°C or –20°C. This is your template DNA for in vitro transcription. 3.2. In Vitro Transcription of the mRNA
The recommended reagents for the in vitro transcription are supplied by Ambion (www.ambion.com) in the mMessage mMachineTM kit. Following the manufacturer’s instructions will generate sufficient amounts of mRNA suitable for microinjection. The protocol described below has been modified slightly to increase yields. 1. Set up the following reaction using your template DNA and reagents from the mMessage mMachine kit: a. 4-ml RNase-free water (there is some in the kit) b. 2-ml 10X transcription buffer (be sure this is completely thawed and there is not a precipitate in the tube) c. 10-ml 2X ribonucleotide mix d. 2-ml template DNA (0.5 mg/ml) e. 2-ml SP6, T3, or T7 polymerase (be sure you are using the right one) 2. Incubate at 37°C for 1 h. 3. Add 1 ml DNase I, mix well and incubate at 37°C for 15 min. 4. Add 115 ml RNase-free water and 15-ml NH4OAc (in kit). 5. Extract with 150-ml phenol/chloroform. 6. Add 150-ml isopropanol and precipitate at –20°C for a few hours to overnight. 7. Centrifuge at 16,000 g for 20 min at 4°C, wash with 70% EtOH (RNase-free) as described above, dry the pellet briefly and resuspend in 20 ml RNase-free water or TE. 8. Take1mlanddeterminetheconcentrationbyspectrophotometry. 9. Take 1 ml and resolve by denaturing gel electrophoresis to confirm that the RNA is intact and of the appropriate size. An example is shown in Fig. 1.2. 10. Store the remaining RNA at –80°C in several aliquots (see Note 3).
3.3. Fertilizing Eggs for Microinjection
1. To induce egg-laying, inject female X. laevis subcutaneously into the dorsal lymph sac with 75 IU PMSG (Calbiochem, catalog #367222) 3–5 days before eggs are desired. 2. Approximately 12–16 h before eggs are needed, inject the frogs with 550 IU HCG (Sigma, catalog #C1063). 3. The next day, collect freshly laid eggs directly into a petri dish containing 0.1X MMR (0.5-mM HEPES, pH 7.8,
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2.3 kb ~1.6 kb
1.3 kb MW
Chk2 mRNA
Fig. 1.2. Ethidium-bromide-stained product of in vitro transcription reaction. RNA was resolved on 1% MOPS agarose gel and visualized under UV light. MW ¼ molecular weight marker.
10-mM NaCl, 0.2-mM KCl, 0.1-mM MgSO4, 0.2-mM CaCl2, 0.01-mM EDTA). 4. Allow fertilization to proceed undisturbed for 10 min, then dejelly eggs in freshly prepared solution of 2% cysteine in 0.1X MMR (see Note 4). 5. Dejellying will take approximately 5–7 min and is complete when eggs pack tightly and geometrically together. As soon as eggs are dejellied, wash eggs 4–6 times in 0.1X MMR. 6. Examine eggs under the microscope. In fertilized eggs, the animal pole will be slightly contracted (occupying less than a full hemisphere of the egg) and will be oriented upward (Fig 1.3A). Fertilized eggs are typically firm when prodded with forceps, like a full balloon. Unfertilized eggs are soft, randomly oriented, and will often stick to the surface of the petri dish (Fig. 1.3B). Sometimes fertilized eggs will remain soft. These are viable but much harder to microinject (Fig. 1.3H). 7. Fertilized eggs should be left undisturbed for 30 min after fertilization before they are microinjected. This allows time for cortical rotation, which establishes the dorsal–ventral axis. 3.4. Injecting the mRNA
3.4.1. Preparing Needles for Microinjection
1. To inactivate contaminating RNases, the microcapillary tubes used to make the microinjection needles should be baked at 80°C for 2 h before they are pulled into needles. 2. The needles can then be pulled using a standard micropipet puller (e.g., Narishige PC-10). 3. Pulled needles should then be attached to the microinjector/ micromanipulator and calibrated using RNase-free water or TE. The tip of the needle typically needs to be broken with a fine, sharp pair of forceps (Fig. 1.3C). 4. The needle should be calibrated to inject the desired amount of RNA (50 ng) in the desired volume (50 nl).
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Fig. 1.3. Stereoscopic images of eggs and the microinjection procedure. (A) Fertilized eggs from X. laevis. Note the contraction of the pigments in the animal hemisphere. (B) A clutch with unfertilized eggs. Animal hemisphere is not contracted and egg remains soft. (C) The tip of a microinjection needle being broken with forceps. (D) The diameter of drops of the injection solution onto parafilm is used to estimate the volume. Note: micrometer is not visible because it is located in the eyepiece of the microscope. (E) Injecting a one-celled embryo. (F) Injecting a two-celled embryo. (G) Injecting a four-celled embryo. (H) Injecting a soft embryo. Embryo is fertilized but soft and therefore challenging to inject without breaking the needle or pushing the needle in too far. Scale bars ¼ 1 mm.
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An easy way to calibrate is to expel a drop of the injection solution onto a piece of parafilm and measure the radius (r) of the drop with a micrometer (Fig. 1.3D). The volume (V) of the drop can then be determined (V ¼ 4/3pr3). 5. The injection volume should then be adjusted (by manipulating injection pressure or time with an automated microinjector or volume directly with a manual microinjector) to achieve the desired volume (see Note 5). 3.4.2. Performing Injections
1. For injection at the one-cell stage (Fig. 1.3E), embryos can be injected from 30 min post-fertilization (pf) until signs that the first cleavage is beginning, usually 90 min pf, but sometimes as early as 60 min pf in warmer ambient temperatures. During this 30–60 min window, an experienced investigator can inject several hundred embryos with 2–5 different mRNAs. 2. Embryos should be injected near the interface of the animal and vegetal hemispheres with care to avoid the animal half where the nucleus resides. Most investigators find it easiest to position the needle so that it will inject in the proper place then bring each embryo to and from the needle with a pair of blunt forceps. 3. The embryos should be pushed onto the needle until it just breaches the membrane. For firm eggs, this will feel like inserting a needle into a full balloon (Figs. 1.3E–G). For very soft embryos, injections will be much more difficult, like piercing a very soft, understuffed pillow (Fig. 1.3H). 4. Depending on the experimental design, the investigator may wait to microinject mRNA at the two-cell stage (Fig. 1.3F). Injection of one of the two blastomeres leaves the remaining uninjected blastomere as an internal negative control. Some investigators inject both blastomeres at the two-cell stage because they have observed better viability than injections at the one-cell stage. 5. Likewise, one can inject a single blastomere at the 4- (Fig. 1.3G), 8-, 16-cell stage and so on. In these experiments, the investigator is typically targeting a particular blastomere of a specific cell lineage (19). These cell cycles last approximately 30 min each, and thus, the window of time for microinjection is shorter.
3.5. Monitoring Expression of the mRNA
1. Expression of the microinjected mRNA should be verified by western blotting of embryo extracts. 2. If the mRNA encodes an epitope tag, such as the FLAG tag described in Section 3.1.1, then blotting for that epitope can be performed with a commercially available antibody (Fig. 1.4A). 3. For the FLAG tag, the mouse M2 monoclonal anti-FLAG antibody works well although there is recognition of a
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nonspecific band at approximately 37 kDa. Blotting for an epitope tag allows the same antibody to be used to verify translation of all mRNAs including the control mRNA. 4. If the mRNA does not encode a tagged protein, then an antibody specific for the encoded protein may be used instead. In this case, endogenous protein will also be detected, and therefore, comparison of relative amounts of endogenous to exogenous protein can be made by comparing western blotting analysis of experimental embryos to uninjected or controlinjected embryos. This information is valuable for estimating the amount of dominant-negative exogenous protein relative to the endogenous, wild-type protein. 3.6. Assaying the Effects of the mRNA on Embryonic Development
3.6.1. Monitoring Embryonic Morphology
1. When using microinjection of mRNA to investigate cell cycle regulation and apoptosis in early embryos, phenotypic changes may manifest early in development, and thus, embryos should be monitored closely for the first 12 h of life (see Note 6). 2. Expression of some mRNAs will induce a modest delay in cleavage time, a delay that may not be appreciable for several cell cycles, but can be discriminated based on number and size of cells at the midblastula stage. 3. Examples of mRNAs that modestly delay cleavage include those encoding: 34-Xic1, an inhibitor of cyclin E-Cdk2 kinase (20); low doses of the cell cycle checkpoint kinase Chk2 (Fig. 1.4B) (5); and the Cdk inhibitory kinase Wee2 (11). 4. Likewise, mRNAs encoding cell cycle activators may accelerate cleavage cycles. Examples include cyclin B mRNA (21) and the Cdk-activating phosphatase Cdc25A (22).
3.6.2. Assays for Cell Cycle Progression
In addition to monitoring cleavage cycles by gross morphology, assays for biochemical and nuclear changes in the cell cycle are summarized here with references to articles providing examples and protocols. 1. Cell cycles can be monitored biochemically by western blotting for the mitotic cyclins or enzymatic assays for Cdk activity (20,21). For these experiments, approximately five embryos per time-point should be collected and snap-frozen. Time-points should be collected every 5–10 min in order to detect oscillations. Only well-synchronized clutches of embryos should be used in order to detect clear mitotic peaks. 2. To monitor cell cycle progression via rounds of DNA replication, incorporation of 3H thymidine into DNA can be followed (13).
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Fig 1.4. Assessment of the effect of microinjection. (A) Western blot for the FLAG epitope on the Chk2 protein. Each lane contains the equivalent of one embryo although extracts from five embryos were pooled for each sample. Note that embryos with low levels of FLAG-tagged protein did not show a cell cycle delay phenotype. (B) Cell cycle delay in embryos injected with Chk2 mRNA versus luciferase (mRNA). Delay is apparent because the cells from embryos injected with Chk2 are larger and fewer. (C) Embryo undergoing apoptosis. Arrows show the area where cells have detached and are filling the space delineated by the vitelline membrane. Scale bars ¼ 1 mm.
These assays provide a different perspective of the cell cycle since cleavages will occur even in the absence of DNA replication in pre-MBT embryos. 3.6.3. Assays for the Midblastula Transition
Expression of mRNAs that alter the rate of cleavage cycles will also affect the timing of most events of the MBT; however, some events, such as the degradation of maternal cyclin E, do not depend on reaching a critical nucleo-cytoplasmic ratio (20). Therefore, assays that monitor several hallmarks of the MBT
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should be employed to best understand the effect of the exogenous mRNA on early development. 1. The onset of zygotic transcription is a classic definition of the MBT and occurs at a critical nucleo-cytoplasmic ratio. Transcription can be monitored by northern blotting for expression of an early developmental gene such as GS17 (13). 2. Alternatively, a more global picture of transcription can be obtained by loading embryos with 3H uridine and then following incorporation of 3H into RNA (13). 3. At the MBT, a host of changes to cell cycle proteins occurs, as cell cycles lengthen and come under control of the zygotic genome. Cell cycle changes that can be assayed by western blot analysis include degradation of maternal cyclin E (20), Cdc25A (22), and Wee1 (23) as well as increased tyrosine phosphorylation of Cdk1 and Cdk2 (5,22). 3.6.4. Assays for Apoptosis
Many exogenous mRNAs trigger a maternally regulated program of apoptosis in early X. laevis embryos. Induction of apoptosis can be readily distinguished from a nonspecific toxic effect of an mRNA. Apoptosis will not be initiated until the early gastrula stage and is characterized by striking morphological events (Fig. 1.4C). 1. In apoptotic embryos, cell lose their attachments from one another and dissociated cells come ‘‘bursting’’ from the blastocoel, eventually filling up the cavity enclosed by the vitelline membrane. A clutch of embryos will undergo apoptosis with good synchrony, with the entire clutch showing an apoptotic phenotype within an hour. 2. Although this apoptotic morphology is distinct to the trained eye, apoptosis should be verified by one or more specific assays. The condensation of chromatin during apoptosis can be identified by electron microscopy or fluorescence microscopy of sectioned embryos stained with a dye that binds DNA (13). However, not every nucleus in the embryo may appear apoptotic and some agents that induce apoptosis, such as those that block DNA replication, do not result in typical apoptotic bodies (4). 3. The detection of DNA ladders by agarose gel electrophoresis is another common assay for apoptosis. These ladders can be detected in DNA isolated from apoptotic X. laevis embryos (12), but the resolution is often poor, obscured by abundant RNA, despite extensive treatment with RNases. 4. Assays that have proven more reliable in detecting apoptosis in X. laevis embryos are the whole-mount TUNEL assay and poly-(ADP ribose) polymerase (PARP) cleavage assay. The TUNEL assay is based on a modification of the whole-mount in situ hybridization protocol for X. laevis embryos (13,14).
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Embryos are fixed, and incubated with terminal deoxytransferase (TdT) and digoxigenin-labeled dUTP (digdUTP). TdT catalyzes the addition of dig-dUTP to free 30 OH ends of DNA An alkaline phosphatase conjugated anti-dig antibody and chromagens are used to detect the labeled nuclei. Because the chromagenic precipitate is a dark purple color that can be obscured by pigment, embryos generated from albino females are typically used. 5. In the PARP assay, embryos are collected when there is morphologic indication of apoptosis. Embryos are lysed and lysates are incubated with recombinant PARP which is then analyzed by western blotting (4). Detection of an 85-kDa cleavage fragment of PARP indicates activation of the apoptotic effector enzyme caspase 3. Because recombinant human PARP is effectively cleaved by apoptotic extracts from X. laevis embryos, all necessary reagents are commercially available.
4. Notes 1. The poly-A tail is thought to improve translatability and/or stability of the mRNA. However translatable mRNAs can be synthesized in vitro using templates that do not encode a polyA tail. Inclusion of 50 and 30 untranslated sequences from the gene may also improve efficiency of translation of the mRNA, but generally is not necessary. 2. For pSP64poly A, EcoRI is the preferred restriction endonuclease because it cuts just after the poly-A tail. If your cDNA contains an EcoRI restriction site, then PvuII is a good alternative. It cuts 182 bp downstream of the poly-A tail. 3. To improve yields and efficiency of recovery even further, set up duplicate or triplicate reactions, pool prior to phenol/chloroform extraction, and complete the procedure, adjusting volumes accordingly. In practice, yields are greater than 2–3 times that of a single reaction, probably due to more efficient recovery in the extraction and precipitation steps. 4. Prolonged exposure to cysteine will damage the eggs. 5. Some investigators find it useful to calibrate several needles ahead of time, but the calibration should be rechecked just
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prior to injecting the embryos. Experienced investigators typically calibrate needles ‘‘on-the-fly’’ as they are needed. 6. Physical penetration of an egg and expression of exogenous RNA could introduce artifacts that affect development. For example, cleavage cycles may lengthen if exogenous mRNA competes with cyclin mRNA for access to the translational machinery. Therefore, control embryos should be microinjected with mRNA encoding an inert or irrelevant protein. A FLAG-tagged luciferase cDNA cloned into pSP64poly A is one useful vector for making control mRNA (24). Alternatively, a cDNA clone encoding the protein of interest could be mutagenized to be catalytically inactive or otherwise inert.
References 1. Kinoshita-Kawada M., Oberdick J., and Xi Zhu M. (2004) A Purkinje cell specific GoLoco domain protein, L7/Pcp-2, modulates receptor-mediated inhibition of Cav2.1 Ca2+ channels in a dose-dependent manner. Brain Res. 132, 73–86. 2. Brandt S. and Fisahn J. (1998) Identification of a K+ channel from potato leaves by functional expression in Xenopus oocytes. Plant & Cell Physiol. 39, 600–6. 3. Morales M.M., Carroll T.P., Morita T., Schwiebert, E.M., Devuyst O, Wilson P.D., Lopes A G., Stanton B.A., Dietz H.C., Cutting G.R., and Guggino W.B. (1996) Both the wild type and a functional isoform of CFTR are expressed in kidney. Am. J. Physiol. 270, F1038–48. 4. Carter A., and Sible J. (2003) Loss of XChk1 function leads to apoptosis after the midblastula transition in Xenopus laevis embryos. Mech. Devel. 120, 315–23. 5. Wroble B., Sible J. (2005) Chk2/Cds1 protein kinase blocks apoptosis during early development of Xenopus laevis. Dev. Dyn. 233, 1359–65. 6. Murray A.W., and Kirschner M.W. (1989) Cyclin synthesis drives the early embryonic cell cycle. Nature 339, 275–80. 7. Newport J. and Kirschner M. (1982) A major developmental transition in early Xenopus embryos: I. Characterization and timing of cellular changes at the midblastula stage. Cell 30, 675–86. 8. Newport J. and Kirschner M. (1982) A major developmental transition in early Xenopus
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embryos: II. Control of the onset of transcription. Cell 30, 687–96. Howe J.A., Howell M., Hunt T., Newport J.W. (1995) Identification of a developmental timer regulating the stability of embryonic cyclin A and a new somatic A-type cyclin at gastrulation. Genes Dev. 9, 1164–76. Howe J.A., and Newport J.W. (1996) A developmental timer regulates degradation of cyclin E1 at the midblastula transition during Xenopus embryogenesis. Proc. Natl. Acad. Sci. USA 93, 2060–4. Leise W.F., III and Mueller P.R. (2002) Multiple Cdk1 inhibitory kinases regulate the cell cycle during development. Dev. Biol. 249, 156–73. Anderson J.A., Lewellyn A.L., and Maller J.L. (1997) Ionizing radiation induces apoptosis and elevates cyclin A1-Cdk2 activity prior to but not after the midblastula transition in Xenopus. Mol. Biol. Cell 8, 1195–206. Sible J.C., Anderson J.A., Lewellyn A.L., and Maller J.L. (1997) Zygotic transcription is required to block a maternal program of apoptosis in Xenopus embryos. Dev. Biol. 189, 335–46. Hensey C., and Gautier J. (1997) A developmental timer that regulates apoptosis at the onset of gastrulation. Mech. Devel. 69, 183–95. Frederick D.L., Andrews M.T. (1994) Cell cycle remodeling requires cell–cell interactions in developing Xenopus embryos. J. Exp. Zool. 270, 410–6. Kroll K.L., Amaya E. (1996) Transgenic Xenopus embryos from sperm nuclear transplantations
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reveal FGF signaling requirements during gastrulation. Development 122, 3173–83. Offield M.F., Hirsch N., and Grainger R.M. (2000) The development of Xenopus tropicalis transgenic lines and their use in studying lens developmental timing in living embryos. Development 127, 1789–97. Heasman J. (2002) Morpholino oligos: making sense of antisense? Dev. Biol. 243, 209–14. Moody S.A. (2000) Cell lineage analysis in Xenopus embryos. Methods Mol. Biol. 135, 331–47. Hartley R.S., Sible J.C., Lewellyn A.L., and Maller J.L. (1997) A role for cyclin E/Cdk2 in the timing of the midblastula transition in Xenopus embryos. Dev. Biol. 188, 312–21.
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21. Hartley R.S., Rempel R.E., and Maller J.L. (1996) In vivo regulation of the early embryonic cell cycles in Xenopus. Dev. Biol. 173, 408–19. 22. Kim S., Li C., and Maller J. (1999) A maternal form of the phosphatase Cdc25A regulates early embryonic cell cycles in Xenopus laevis. Dev. Biol. 212, 381–91. 23. Murakami M.S., and Woude G.F.V. (1998) Analysis of the early embryonic cell cycles of Xenopus; regulation of cell cycle length by Xe-wee1 and Mos. Development 125, 237–48. 24. Kappas N.C., Savage P., Chen K.C., Walls A.T., Sible J.C. (2000) Dissection of the XChk1 signaling pathway in Xenopus laevis embryos. Mol. Biol. Cell 11(9), 3101–8.
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Chapter 2 Use of Luciferase Chimaera to Monitor PLCz Expression in Mouse Eggs Karl Swann, Karen Campbell, Yuansong Yu, Christopher Saunders and F. Anthony Lai Abstract The microinjection of cRNA encoding phospholipase C (PLC zeta) causes Ca2+ oscillations and the activation of development in mouse eggs. The PLC protein that is expressed in eggs after injection of cRNA is effective in causing Ca2+ oscillations at very low concentrations. In order to measure the amount and timecourse of protein expression we have tagged PLC with firefly luciferase. The expression of the luciferase protein tag in eggs is then measured by incubation in luciferin combined with luminescence imaging, or by the lysis of eggs in the presence of Mg-ATP and luciferin in a luminometer. The use of luciferase to monitor protein expression after injection of cRNA is a sensitive and effective method that efficiently allows for sets of eggs to be used for PLC quantitation, Ca2+ imaging, and studies of embryo development. Key words: Luminescence, luciferase, phospholipase, egg.
1. Introduction Mammalian eggs are large cells (100 mm in diameter) and readily amenable to microinjection. We have used pressure-based microinjection as a means of introducing molecules into mouse eggs for many years. Our particular interest over the last few years has been focused on the role of a sperm-specific phospholipase C (PLC zeta) in causing the Ca2+ changes that lead to egg activation in mammals. This protein can trigger repetitive Ca2+ oscillations that are very similar to those seen at fertilization in mouse, pig, and human eggs (1). We have proposed that PLC is the ‘‘sperm factor’’ that is delivered by the sperm into the egg following David J. Carroll (ed.), Microinjection: Methods and Applications, Vol. 518 Ó 2009 Humana Press, a part of Springer ScienceþBusiness Media, LLC DOI 10.1007/978-1-59745-202-1_2
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gamete fusion (2). It has been shown that recombinant PLC can cause Ca2+ release when it is injected into mouse eggs (3). We, and others, have also carried out biochemical studies of recombinant PLC (3,4). However, one of the major problems of working with recombinant PLC protein is that its Ca2+ oscillation-inducing activity is very labile, consistent with our observation that the PIP2 hydrolysis enzymatic activity is difficult to maintain. Consequently, we have used cRNA injection as a general means of introducing PLC into mouse eggs. The injection of cRNA for PLC also has the advantage that no protein contaminants are introduced into cells, in contrast to native protein isolation procedures. One of the disadvantages of injecting cRNA PLC is that the level of expressed protein cannot be readily determined in living cells. An effective way to measure how much protein is being expressed is to inject cRNA for PLC that has been tagged with firefly luciferase (4). Using luciferase luminescence to measure protein expression is a highly sensitive technique. Luminescence has an advantage over the use of fluorescence-based methods in that it does not suffer from interference from auto-fluorescence which is quite considerable in mammalian eggs (5). The issue of sensitivity is particularly important since mouse PLC is active in mouse eggs at concentrations of 1–10 nM (3), and this very low level of protein is on the limit of detection for the most sensitive fluorescent proteins used in eggs (6). Furthermore, since human and monkey PLC appear to be more potent in causing Ca2+ oscillations in eggs than the mouse PLC (7), the physiological levels of PLC expression in many species may be undetectable using fluorescently tagged PLC. The chief disadvantage in using luciferase luminescence is that the localization of the expressed protein is poor compared to fluorescence probes, where high-resolution confocal imaging can be used. However, with photon imaging cameras it is certainly possible to identify which individual eggs, or cells, are expressing luciferase and it is possible to quantitatively estimate how much luciferase-tagged protein is being expressed. In this chapter, we describe how we study the effects of PLC, and other PLC isoforms, or various mutant versions of PLCs, by injection of cRNA encoding luciferase-tagged versions of the proteins. In many experiments, we inject the cRNA and then measure Ca2+ signals from eggs for several hours after injection, before calibrating luciferase expression at a set time-point. In other cases, we monitor Ca2+ oscillations and luciferase expression from eggs and then place them in culture for further studies on their development. The methods we describe for PLC can be readily applied to other proteins and, for example, we have also injected and studied cyclin B levels in mouse eggs using a luciferase-tagged cRNA (unpublished data).
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2. Materials 2.1. DNA and RNA Preparation
1. To produce sufficient quantities of DNA plasmid, we use Qiagen’s Plasmid Maxi Kit. Restriction enzymes are routinely purchased from New England Biolabs. To produce polyadenylated cRNA, mMessage mMachine T7/T3/SP6 kits and Poly (A) Tailing Kits are used, along with SUPERase-In RNAse Inhibitor (Ambion). Rabbit Reticulocyte Lysate (Promega) is used to assess RNA quality. 2. RNAse-free solutions and plasticware are prepared using treatment with diethyl pyrocarbonate (DEPC, Sigma). 3. Media for mouse eggs consists of M2 and acidified Tyrode’s solution (Sigma). We also use KSOM and a Hepes version of KSOM (HKSOM) which is made from stock using embryotested chemicals (Sigma) and clinical-grade water. The constituents of KSOM and HKSOM are given in reference (8) and (9). Hyaluronidase M2 and acid Tyrodes solution are stored in aliquots at 20°C. The M2 or HKSOM media are stored at 4°C and used for 2–3 weeks. For all imaging studies, firefly luciferin is added to HKSOM media. The luciferin (L6882 from Sigma) is made up at 100 mM in distilled water and stored in the 20°C freezer for 1 month. It is diluted into HKSOM shortly before use to give a final concentration of 100 mM (see Note 1). 4. The injection buffer consists of KCl/Hepes (120-mM KCl, 20-mM HEPES, pH 7.2). The buffer is made up in plastic vessels and then mixed with 1% Chelex 100 beads (Sigma) for 1 h (to remove divalent cations) before being filter-sterilized. For experiments where intracellular Ca2+ is to be measured Oregon Green BAPTA dextran (OGBD) (Molecular Probes, www.probes.com) is added to the injection buffer. Aliquots of injection buffers are stored in the 20°C freezer.
2.2. Mouse Eggs
1. We regularly use the MF1 strain of mice, but have obtained similar results with other strains of mice such as CD-1, or with F1 hybrid cross strains. The hormones were purchased from Dunlops Veterinary Supplies (www.dunlops.com). The superovulation of mice and collection of eggs is described in laboratory manuals (10). 2. Eggs were manipulated in M2 media (Sigma) using fine-bore glass pipettes that were pulled in a flame to a diameter of approximately 80–100 mm.
2.3. Microinjection
1. For microinjection, borosilicate glass capillaries (GC150F, Harvard Apparatus Ltd., 1.5-mm outer diameter and 0.86mm inner diameter) with an internal filament were pulled on a
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vertical pipette puller (Model P-30; Sutter Instruments). The pipettes used for injection should be checked for appropriate tip size. This can be done by finding the minimal pressure required to blow bubbles in ethanol (11). Injection needles are backfilled with sterile microloader pipette tips (eppendorf). 2. Injection needles are clamped in a holder with a silver wire and side port (World Precision Instruments Inc, www.wpi-europe.com). The holder is plugged into a preamplifier that is electrically connected to an intracellular amplifier (e.g., Electro 705 or Cyto 721; WPI). The preamplifier is held in the micromanipulator. 3. Pressure is applied to the back of the needle by pulses from a pressure injection system (Picopump, WPI) connected to the side port of the needle holder with stiff silicone tubing. 4. Mouse eggs are injected while being held by a ‘‘holding’’ pipette (Hunter Scientific) using suction via a syringe system (Narashige) containing embryo-tested mineral oil (Sigma). 5. The preamplifier and needle holder, and the ‘holding pipette’, are mounted on hydraulic manipulators (Narashige) that are fixed to the inverted microscope (TE2000, Nikon UK Ltd). 2.4. Imaging and Quantifying Luciferase
1. For imaging, the eggs are maintained in drops of media in a heated chamber (Intracel Ltd.) on the stage of an inverted fluorescence microscope (either a Nikon TE2000, or Zeiss Axiovert S100). Each microscope has the facility to direct 100% of the light from the eggs to the camera via either a side port or the base port. 2. The light collected from injected eggs is imaged using photoncounting imaging cameras. The cameras we currently use are cooled intensified CCD cameras (Photek Ltd.; www.photek.com). Photek’s software is used for data collection and analysis (see Note 1). 3. The imaging systems are contained in a lightproof dark box. In one case, we have the facility to direct bright field or fluorescent illumination to the eggs via fiber optical cables. The gating of these light sources is controlled via Photek’s software that controls electro-mechanical shutters (Uniblitz; www.uniblitz.com) that are integrated into the light box. In another case, the microscope light sources are inside the box and simply switched off during luminescence imaging (see Note 2). 4. After imaging, the eggs can be lysed in order to quantify luciferase expression. This is done in a lysis buffer using a custom-made luminometer (see reference (12)). This essentially consists of a dark-proof tube holder that is adjacent to a cooled photomultiplier tube (S20 type tube, with a
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cooled housing and amplifiers from Electron Tubes Ltd.; www.electrontubes.com). The light output is measured using photon-counting discriminators and amplifiers with software supplied by Electron Tubes Ltd. Commercially available tube luminometers would also be suitable. 5. The lysis buffer consists of phosphate-buffered saline with 1mM MgATP and 100-mM luciferin. Eggs are lysed with Triton X-100 and the amount of luciferase calibrated with recombinant luciferase protein. All these reagents are purchased from Sigma.
3. Methods 3.1. Synthesis of cRNA
3.1.1. Preparation of DNA
1. RNAse-free solutions and plasticware are prepared by incubation in a solution of 0.1% DEPC overnight in a fume hood. Following this incubation, residual DEPC is removed by autoclaving. We routinely generate microgram quantities of the required DNA plasmid using Qiagen’s Plasmid Maxi Kit, following the manufacturer’s instructions (a standard molecular biology/microbiology textbook (13) provides further information on Escherichia coli strains, transformation, and handling). Depending on the plasmid copy number, 500 ml of DNA with a concentration of 0.5–2 mg/ml can be harvested from 250 ml of culture. 2. Due to the high processivity of RNA polymerases, it is necessary to linearize the circular plasmid DNA to prevent the production of extremely long RNA molecules. This can be achieved by digestion with a suitable restriction enzyme, obviously avoiding those which cut between the promoter and gene of interest. A 100-ml restriction digest containing 10 mg of DNA and 20 U of enzyme is incubated at a suitable temperature overnight. Complete linearization can be confirmed by running 2 ml on an agarose gel if necessary. 3. The linearized DNA is cleaned up by phenol/chloroform extraction. Essentially, an equal volume of TRIS-buffered phenol:choloroform:isoamylalcohol (25:24:1 v/v/v) is added and mixed by vigorous inversion for 30–60 s. The phases are separated by microcentrifugation at 14000 g for 3 min and the top, aqueous phase is transferred to a DEPC-treated microfuge tube using a DEPC-treated tip, taking care not to transfer any of the proteinaceous, white interphase. This extraction is repeated twice more.
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4. The DNA is precipitated by addition of 80 ml of isopropanol and 18 ml of 3-M sodium acetate, pH 5.2 and, following an incubation at 80°C for 1 h, is pelleted by microcentrifugation at 14000 g for 20 min at 4°C. The supernatant is removed and the pellet washed with 80% ethanol. Following a further 20min spin, the pellet is left to air-dry for 10–15 min.
3.1.2. Translation and Polyadenylation of RNA
1. The DNA pellet is resuspended in 6 ml of nuclease-free water containing 20 U of SUPERase-In RNAse Inhibitor. This is then transferred to a fresh microcentrifuge tube, and a transcription reaction assembled at room temperature by adding 10 ml of ice-cold 2xNTP/CAP mix, 2 ml of room-temperature 10X reaction buffer, along with 2 ml of enzyme. 2. The reaction is then incubated at 37°C for 2 h. Addition of a poly-A tail, to enhance RNA longevity, is achieved by addition of 36-ml nuclease-free water, 20-ml E-PAP buffer, 10-ml 25mM MgCl2, 10-ml 10-mM ATP, and 4-ml E-PAP to the 20 ml transcription reaction. The reaction is again incubated at 37°C for a further 2 h. 3. The polyadenylated RNA is precipitated by addition of 150-ml lithium chloride precipitation solution, and incubated at 80°C for >1 h. The RNA is pelleted by 4°C microcentrifugation at 14000 g for 30 min. The supernatant is discarded, the pellet washed with 80% ethanol, and further centrifuged for 15 min. Following air-drying, the RNA is resuspended by addition of 9 ml of nuclease-free water and 1 ml (20 U) of SUPERase-In RNAse Inhibitor.
3.1.3. Quantification and Dilution of RNA
1. The RNA is quantified by measuring its absorbance at 260 nm in a 500-ml quartz cuvette. We routinely use 1 ml of sample, giving a dilution factor of 1 in 500. The amount of RNA is then calculated using the standard equation: RNA conc ðmg=mlÞ ¼ 40 A260 500 2. The RNA can stored at 80°C in 1-ml aliquots. We commonly store RNA as either undiluted stock aliquots, or working aliquots diluted to 2, 0.2, or 0.02 mg/ml. When diluting, we commonly add 20U SUPERase-In to the RNA prior to aliquotting. 3. Just before injection, the RNA is mixed with other components, such as OGBD, and then kept on ice for the period over which injections are carried out (<1 h). If there is any remaining solution in the aliquots after injection, it is discarded and a fresh aliquot is used for each injection session.
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3.1.4. Assessment of RNA Quality
Initially, it may be necessary to check batches of RNA for signs of RNAse contamination, which leads to degradation of the RNA. Commonly this can be achieved by running an aliquot of RNA on a denaturing agarose gel, checking for the presence of a single species of defined size without a smear of lower-molecular-weight fragments, indicative of RNA degradation. However, due to the heterogeneity in size of the poly-A tail, a single species is rarely seen, leading to some degree of uncertainty about the quality of the RNA. Instead, we check that the RNA can be translated into a protein of the predicted molecular weight. This is achieved using 1–2 mg of RNA in a Rabbit Reticulocyte Lysate reaction (Promega). We label the protein with [35S]-Pro-Mix, and, following separation on SDS-PAGE, use autoradiography to determine its molecular weight. Alternatively, we have also satisfactorily used ‘‘cold’’ methionine in the reaction, and then used antibodies to detect the protein of interest on a western blot following the SDS-PAGE.
3.2. Microinjection of Eggs
1. Zona intact mouse eggs are placed in a shallow drop (1 ml) of media covered in oil, in the lid of a petri dish. The dish is placed on the microscope stage without heating. A silver wire is placed in the injection drop and this wire is held in place via a small manipulator. This wire is connected to a longer standard copper wire to the chassis ground of the electrical amplifier. This wire allows for an electrical circuit to form between the ground and the tip of the pipette once it is place in the media (a circuit is indicated by the amplifier). If this does not occur, then the pipette should be replaced because the tip is probably blocked. 2. The RNA solution to be injected should be spun (13,000 rpm in a benchtop microcentrifuge) for several minutes before injection. For RNA injection, we generally use tips of 0.75–0.9 mm in diameter. The injection pipettes are backfilled with <1 ml of injection solution containing the RNA. 3. This pipette is then held in the specialized holder (containing a silver wire) that is then fitted onto the preamplifier that is itself clamped into the micromanipulator. 4. For experiments where fluorescence is also measured, to look at changes in Ca2+ dynamics within the egg, the luciferasetagged cRNA is mixed with an equal volume of 1-mM OGBD prepared in KCl Hepes pH 7.2. Even if a Ca2+ dye is not to be used, it is useful to mix the RNA solution 1:1 with KCL Hepes buffer so as to have some salts present in the injection buffer. 5. For injection, each egg is held by suction with the holding pipette and then the tip of the injection pipette is manipulated so that it will touch the plasma membrane. The injection pipette is then pushed into egg in a way that deforms the zona pellucida. At some point, the zona will jump back into
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shape, which is a sign that the zona pellucida is penetrated. At this point, the negative capacitance is applied to the amplifier that is connected to the back of the specialized pipette holder. This causes the pipette tip to enter the cytoplasm. The operator should then make sure that the tip of the injection pipette is in focus and a pressure pulse is applied from the picopump to push a bolus of solution into the egg. 6. The pressure pulses we use are typically 100 ms1 s long, at a pressure of 20 psi. The volume of solution injected is estimated by the diameter of cytoplasmic displacement caused by the injection and should correspond to 3–5% of the egg volume. In practice, the first egg can be used for a test with the pulse duration and pressure being adjusted to suit the amount of solution injected. Once familiar with this system, it is possible to inject >30 eggs in 20 min. However, tips often become blocked during injection and need to be replaced. 1. After injection, the eggs are placed in drops of media for imaging. In most experiments, the eggs are left zona intact and placed in a small (50 ml) drop of HKSOM media, which is under mineral oil in a heated (37°C) chamber with a glass coverslip that sits on the inverted microscope. The HKSOM media contains BSA (4 mg/ml) and 100-mM luciferin (see Note 3). Figure 2.1 shows the luminescence from a single mouse egg injected with cRNA for luciferase alone. The timecourse of luciferase expression lasts at least 10 h with a peak at around 4–5 h post-injection. 2. For some experiments where we want to add extra compounds, or sperm, during the course of imaging, the eggs have to be stuck down. We do this by briefly treating the eggs with acid Tyrode’s solution to remove the zona pellucidas and then immediately placing the eggs in 1-ml drop of the HKSOM in a chamber that has a polylysine-coated (1 mg/ml) coverslip.
Photons per minute
3.3. Imaging of Luciferase Luminescence
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0
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Fig. 2.1. Luminescence (in photon counts per minute) from a mouse egg injected with luciferase cRNA. The egg was injected with 10 pl of 2 mg/ml of cRNA and imaged in media in the presence of 100-mM luciferin 10–20 min after injection.
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3. To image intracellular Ca2+ in eggs (injected with OGBD), we monitor fluorescence for the period during which Ca2+ signals occur (5 h). At the end of this period, the luminescence is then measured on the same set of eggs (see Note 4).
a)
Fluorescence (a.u.)
4. Fluorescence or luminescence is imaged in the eggs using either 20x 0.65 NA or 10x 0.5 NA Fluor objectives. The light (100%) is directed via either a sideport or baseport to the ICCD camera. We use the same ICCD camera to monitor both fluorescence and luminescence. The main difference is that during fluorescence measurements, the eggs are exposed to excitation light. This means that a standard fluorescence filter block is in place to enable epifluorescence illumination. For OGBD we use a FITC block or else a modified block with a 500-nm longpass filter. Figure 2.2a shows the relative changes in fluorescence from eggs injected with OGBD and PLC-luc cRNA. The spike-like increases indicate intracellular Ca2+ oscillations, as described previously.
b)
Luminescence (photons per minute)
30 min 250
egg 2 egg 1
0 5 Hours
c) 1 2
Fig. 2.2. Mouse eggs injected with PLC-luc cRNA. Eggs were injected with 10 pl of 0.2 mg/ml cRNA. In (a) the fluorescence (in arbitrary units, a.u.) of Oregon Green BAPTA dextran is shown. The oscillations in fluorescence indicate intracellular Ca2+ oscillations are occurring. In (b) the luminescence from two other eggs injected with PLC-luc cRNA and incubated in 100-mM luciferin is shown. The luminescence is recorded continuously with this experiment. In (c) an image of the group of eggs injected with PLC-luc cRNA is shown for different time periods. The arrows point to eggs 1 and 2 that are shown in (b).
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5. The excitation light source is from a halogen lamp (see Note 5). Fluorescence is monitored in injected eggs for 4 h or 5 h, and then by measuring the OGBD fluorescence with low-level excitation light, the luminescence is measured from the same set of eggs by recording the light from eggs with the excitation light turned off. During the fluorescence recording, the luminescence signal may increase slightly and so the recorded fluorescence signal actually contains a small component of luminescence. However, the fluorescence signals are typically more than 100 times greater than the luminescence signals, so can be ignored in practice. During fluorescence recording, the camera’s sensitivity can be reduced to 10%. 6. At the end of the fluorescence measurements, the same set of eggs are then monitored for luminescence by integrating light emission (in the absence of fluorescence excitation) for 20 min using the same ICCD camera. The cameras we use, typically have a very low background count such that the background noise from an area the size of one egg is about 1 photon per minute. The luminescence signal starts to increase above background within 10 min of the start of recording (Fig. 2.2). The signal then continues to increase for several hours and does not start to decrease until about 8–10 h. The level of signal depends upon the amount of cRNA injected and the particular construct used (see Note 6). Figure 2B and C shows the images and luminescence integrated from different time periods, as well as the timecourse of expression from two of the eggs in the group that illustrate the range of variation in luminescence. 7. If the experiment only requires a measure of the relative amounts of lufciferase expression, then zona intact eggs can be removed from the imaging drop and placed in KSOM media in drops under oil in a 37°C 5% CO2 incubator. If an absolute calibration of expression is required then the eggs are lysed in a luminometer. 3.4. Quantifying Luciferase Expression in a Luminometer
1. Imaging the luciferase luminescence from eggs can ensure that all eggs counted in an experimental group express the luciferase-tagged PLC. To measure the amount of luciferase protein expressed, groups of eggs are collected from the imaging drop and then lysed in a buffer in a luminometer (see Note 7). For each experiment, groups of eggs, verified as being luminous, are collected and placed in a test tube containing phosphatebuffered saline with 1-mM Mg2+ATP and 100-mM luciferin. 2. The eggs are then lysed with 0.5% Triton X-100 and the steadystate light compared to that emitted from serial dilutions of recombinant firefly luciferase (Sigma) in the same buffer. The amount of luciferase activity measured for each group of eggs is then divided by the number of luminous eggs to obtain the
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mean value for protein expression of each type of PLC-luciferase. We have found that injection of mouse PLC-luciferase into mouse eggs can lead to Ca2+ oscillations and the expression of 0.1–0.2 pg of protein in a 4-h period following injection (4).
4. Notes 1. We have previously used an imaging photon detector (IPD) system which was set up by, and used software from, Sciencewares (www.sciencewares.com). The IPD camera (Photek Ltd.) uses a different principal for light collection from the ICCD cameras. We have not noticed any significant difference in sensitivity of these two types of photon-counting detectors. 2. It is essential that some form of dark box is constructed around the microscope. The light level in a typical darkened room, where standard fluorescence microscopy is carried out, is usually much too high and causes considerable interference when imaging luminescence. Any light sources within the darkbox should be removed or covered with black tape. If the microscope used is motorized in any way, it will probably be necessary to switch it off completely during luminescence imaging since internal LEDs will cause an elevated background light. 3. We use 100 mM for mouse eggs, but a range of luciferin concentrations (1 mM to 1 mM) are cited for use in luciferaseimaging of cells in general. The higher concentrations are not always the most effective, because the luciferin luciferase reaction shows ‘‘flash kinetics’’ which means that the reaction rate can decrease with time due to inhibition from the reaction product, oxyluciferin (12). The best concentration to use can depend upon a range of factors that are specific to a cell type, and it is best to test different concentrations. The luciferase also depends upon ATP and can be used to monitor ATP concentrations in eggs (5, 14). For monitoring the timecourse of expression this is not a major issue because the changes in luminescence caused by ATP increases in activating eggs accounts for a 10–20% change in the total light output, and this is hardly detectable in studies using effective amounts of PLC-luc, where the luminescence signals are about 1–10 photons per second for each egg. In order to effectively measure the ATP change at fertilization, we have to inject high concentrations of recombinant luciferase protein, which results in luminescence values of 100–1000 photons per second per egg (14). 4. We use OGBD to measure Ca2+ because the imaging systems we use can only monitor fluorescence at a single wavelength.
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Consequently, a dye such as OGBD is used, since it undergoes an increase in fluorescence intensity with an increase in Ca2+, and being dextran-linked it does not undergo compartmentalization which can be a problem in mouse eggs (15). Other dyes, such as fura 2, which permits ratio excitation, could be used in conjunction with luciferase monitoring. However, it is worth noting that luciferin is fluorescent when excited with UV light, so if fura 2 is to be used, the luciferin should not be added to the media until the fluorescence imaging is finished. 5. A halogen light source with a stabilized power supply is used because this can easily be reduced to the minimum level required. The excitation light used with photon-counting cameras is generally much lower than with standard cooled CCD cameras and so the use of Xenon lamp (for example) with multiple neutral density filters creates unnecessary light and heat. 6. It is important to optimize the length of the spacer residues present between the protein of interest and luciferase, as we have found that this can alter the expression level of the protein. This may be related to the potential for changes in the protein secondary structure, consequent to tagging with luciferase, and will vary with each protein. For PLCs, we find that a spacer of four residues works well. 7. The relative expression of a PLC can be easily assessed and this can be used for studies on egg activation, or for studies on the effects of PLC on later embryo development. However, it is not simple to calibrate the absolute amount of protein expressed in single eggs using the luminescence from living eggs on the microscope. This is partly because the light emitted from firefly luciferase depends upon ATP and pH. Consequently, the precise level of free ATP and pH in an egg would have to be known to calibrate the absolute amount of luciferase.
Acknowledgments Our work is supported by the BBSRC, Wellcome Trust, and Cardiff Partnership Fund.
References 1. Swann, K., Saunders, C.M., Rogers, N. and Lai, F.A. (2006) PLC (zeta): A sperm protein that triggers Ca2+ oscillations and egg activation in mammals. Sem. Cell & Dev. Biol. 17, 264–73.
2. Saunders, C.M., Larman, M.G., Parrington, J., Cox, L.J., Royse, J., Blayney, L.M., Swann, K. and Lai F.A. (2002) PLC: a sperm-specific trigger of Ca2+ oscillations in eggs and embryo development. Development 129, 3533–44.
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3. Kouchi, Z., Fukami, K., Shikano, T., Oda, S., Nakamura, Y., Takenawa, T and Miyazaki, S. (2004) Recombinant phospholipase Czeta has high Ca2þ sensitivity and induces Ca2þ oscillations in mouse eggs. J. Biol. Chem. 279, 10408–12. 4. Nomikos, M., Blayney, L.M., Larman, M.G., Campbell, K., Rossbach, A., Saunders, C.M., Swann, K. and Lai, F.A. (2005) Role of phospholipase C-(zeta) domains in Ca2+-dependent phosphatidylinositol 4,5-bisphosphate hydrolysis and cytoplasmic Ca2+ oscillations. J. Biol. Chem. 280, 31011–18. 5. Dumollard, R., Marangos, P., Fitzharris, G., Swann, K., Duchen, M. and Carroll, J. (2004) Sperm-triggered [Ca2+] oscillations and Ca2+ homeostasis in the mouse egg have an absolute requirement for mitochondrial ATP production. Development 131, 3057–67. 6. Yoda, A, Oda, S., Shikano, T., Kouchi, Z., Awaji, T., Shirakawa, H., Kinoshita, K and Miyazaki, S. (2004) Ca2+ oscillation-inducing phospholipase C zeta expressed in mouse eggs is accumulated in the pronucleus during egg activation. Devel. Biol. 268, 245–57 7. Cox, L.J., Larman, M.G., Saunders, C.M., Hashimoto, K., Swann, K. and Lai, F.A. (2002) Sperm phospholipase C from humans and cynomolgus monkeys triggers Ca2+ oscillations, activation and development of mouse oocytes. Reproduction 124, 611–23.
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8. Summers, M.C., Bhatnagar. P.R., Lawitts. J.A., Biggers. J.D. (1995) Fertilization in vitro of mouse ova from inbred and outbred strains: complete preimplantation embryo development in glucose-supplemented KSOM. Biol. Reprod. 53, 431–7. 9. Summers, M.C., McGinnis, L.K., Lawitts, J.A., Raffin, M., Biggers, J.D. (2000) IVF of mouse ova in a simplex optimized medium supplemented with amino acids. Hum. Reprod. 15, 1791–801. 10. Hogan, B., Costantini, F. and Lacy, E. (1986). Manipulating the Mouse Embryo: A Laboratory Manual. Cold Spring Harbour Laboratory, Cold Spring Harbor, NY. 11. Schnorf, M., Potrykus, I. and Neuhaus, G. (1994) Microinjection technique: routine system for characterization of microcapillaries by bubble pressure measurement. Exp. Cell. Res. 210, 260–7. 12. Campbell, A.K. (1988) ‘Chemiluminescence’ Ellis Horwood Series on Biomedicine. 13. Sambrook, J. and Russell, D.W. (2001) Molecular Cloning: A Laboratory Manual. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY. 14. Campbell, K. and Swann, K. (2006) Ca2+ oscillations stimulate an ATP increase during fertilization. Devel. Biol. 298, 225–53. 15. Carroll, J., Swann, K., Whittingham, D. and Whitaker, M.J. (1994) Spatiotemporal dynamics of intracellular [Ca2+]i oscillations during growth and meiotic maturation of mouse oocytes. Development 120, 3507–17.
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Chapter 3 Analysis of 14-3-3 Family Member Function in Xenopus Embryos by Microinjection of Antisense Morpholino Oligos Jeffrey M. C. Lau and Anthony J. Muslin Abstract The 14-3-3 intracellular phosphoserine/threonine-binding proteins are adapter molecules that regulate signal transduction, cell cycle, nutrient sensing, apoptotic, and cytoskeletal pathways. There are seven 143-3 family members, encoded by separate genes, in vertebrate organisms. To evaluate the role of individual 14-3-3 proteins in vertebrate embryonic development, we utilized an antisense morpholino oligo microinjection technique in Xenopus laevis embryos. By use of this method, we showed that embryos lacking specific 14-3-3 proteins displayed unique phenotypic abnormalities. Specifically, embryos lacking 14-3-3 t exhibited gastrulation and axial patterning defects, but embryos lacking 14-3-3 g exhibited eye defects without other abnormalities, and embryos lacking 14-3-3 appeared completely normal. These and other results demonstrate the power and specificity of the morpholino antisense oligo microinjection technique. Key words: Morpholino, Xenopus, microinjection, embryogenesis, 14-3-3.
1. Introduction 14-3-3 proteins are intracellular dimeric phosphoserine-binding molecules that regulate important aspects of cell physiology (1–3). 14-3-3 family members participate in signal transduction, cell cycle, apoptotic, metabolic, and cytoskeletal pathways. In vertebrate organisms, there are seven 14-3-3 family members encoded by separate genes, named 14-3-3 , g, ", , , t, and (2). When tested in vitro, all seven family members bind with similar affinity to phosphoserine-containing peptides containing the RSxSxP motif, where x is any amino acid (4,5). Despite their similar binding properties in vitro, each family member has a unique expression pattern in embryonic development, and there David J. Carroll (ed.), Microinjection: Methods and Applications, Vol. 518 Ó 2009 Humana Press, a part of Springer ScienceþBusiness Media, LLC DOI 10.1007/978-1-59745-202-1_3
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may be a secondary contact point that differentiates the binding properties of individual 14-3-3 proteins in vivo (6,7). In previous work, we investigated the role of 14-3-3 proteins in Xenopus laevis development by use of a peptide inhibitor of all 14-3-3 proteins, the R18 peptide (8). We injected RNA encoding the R18 peptide linked to glutathione S-transferase (GST). Microinjection of GST-R18 into two-cell embryos resulted in major phenotypic abnormalities, including reduced mesoderm induction with abnormal gastrulation. These and other experiments showed that global 14-3-3 inhibition blocked FGFmediated mesodermal differentiation and patterning. To test the role of individual 14-3-3 proteins in early Xenopus development, we used a morpholino antisense oligo microinjection technique (6). In this method, morpholinos that target individual Xenopus 14-3-3 family members were injected into two-cell-stage embryos. The ability of the morpholinos to specifically knockdown target protein levels was documented by analysis of embryonic protein lysates by Western blotting with family member-specific anti-14-3-3 antibodies. Phenotypic abnormalities in injected embryos were evaluated by visual inspection and by analysis of gene expression by whole-mount in situ hybridization. These experiments demonstrated that embryos that were lacking 14-3-3 t, and to a lesser degree 14-3-3 ", exhibited gastrulation and axial patterning defects, but embryos lacking 14-3-3 exhibited eye defects without other abnormalities. Embryos lacking 14-3-3 appeared completely normal. Therefore, individual 14-3-3 proteins have distinct roles in the regulation of Xenopus development.
2. Materials 2.1. Sexually Mature Xenopus laevis
1. African clawed frogs, Xenopus laevis, are obtained from Xenopus One (Dexter, MI, http://www.xenopusone.com), Xenopus Express (Brooksville, FL, http://www.xenopus.com), or Nasco (Modesto, CA, http://www.enasco.com). 2. Wild-type sexually mature males (7.5–9 cm) and females (9–14 cm) are used for in vitro fertilization experiments. For whole-mount in situ hybridization and immunostaining experiments, sexually mature albino males and females are used to obtain non-pigmented embryos.
2.2. Morpholino Antisense Oligos
1. Morpholinos were purchased from GeneTools, LLC (Philomath, OR). Morpholinos are designed to be antisense to the initiation AUG and subsequent 22 ribonucleotide residues in the mRNAs encoding specific Xenopus proteins (see Note 1).
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2. Sequence of the specific morpholinos are the following: 14-3-3 morpholino, 5-TCTGTACCAGTTCACTCTTGTCCAT-30 ; 14-3-3 morpholino, 50 -GCAACTGCTGCTCCCGATCAGCCAT-30 ; 14-3-3 " morpholino, 50 -ACACTAAATCCTCT CGCTCTTCCAT-30 , and 50 -TACACTAAATCCTCTCGCTC TTCCA-30 ; 14-3-3 g morpholino, 50 -GCACCAGCTGCTCG CGGTCCACCAT-30 ; 14-3-3 t morpholino, 50 -TCTGGATTT GTGCGGTCCTGTCCAT-30 , and 50 -GTCTGGATTTGTGC GGTCCTGTCCA-30 ; and 14-3-3 morpholino, 50 - TCTGGA CCAGTTCATTTTTATCCAT-30 . The control morpholino is a scrambled version of the experimental morpholino commercially available from GeneTools, LLC (Philomath, OR). 3. Morpholinos are re-constituted in RNAase-free distilled water to a final concentration of 1 mmol/ml and stored in aliquots at 80°C. 2.3. Family-MemberSpecific Anti-14-3-3 Antibodies and Western Blotting
1. 14-3-3 family member-specific antibodies (Santa Cruz Biotechnology, Inc., Santa Cruz, CA) are used to detect specific knockdown of 14-3-3 family member protein levels on immunoblots. Total ERK protein level is measured with anti-p44/p42 antibody (#9102, Cell Signaling, Danvers, MA) as loading control. 2. Primary antibodies: 14-3-3 (A-15, sc#17288), " (T-16, sc#1020), g (C-16, sc#731), (K-12, sc#17286), t (C17, sc#732), (C-16, sc#1019) primary antibody stock solutions are stored at 4°C. Working solutions are prepared by making a 1:500 dilution of the stock antibody in 1X TBS/T containing 5% bovine serum albumin (#A9647, Sigma) and 0.05% sodium azide. Working solutions are stored at 4°C and can be re-used multiple times. 3. Primary antibody: p44/p42 total ERK antibody (#9102, Cell Signaling, Danvers, MA) is stored at 20°C. Working solution is prepared by making a 1:1000 dilution of the stock antibody in 1X TBS/T containing 5% bovine serum albumin (#A9647, Sigma) and 0.05% sodium azide (Sigma). The working solution is stored at 4°C and can be re-used multiple times. 4. 10x TBS/T: 100-mM Tris-HCl pH 8.0, 1.5-M NaCl, 0.5% Tween 20. Dilute to 1X concentration with distilled water to make 1X working solution. Store both 10X and 1X solutions at room temperature. 5. Secondary antibody solution: Horseradish peroxidase (HRP)-conjugated secondary antibody derived from the correct species (Cell Signaling, Danvers, MA) is diluted 1:5000 in 10 ml of 1X TBS/T containing 5% fat-free dry milk powder.
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6. ECL Western blotting detection reagents (Amersham Biosciences, England) and ISO-MAX autoradiography/X-ray film (#GX-330810, Scimart, St. Louis, MO). 7. NP40 lysis buffer: 20-mM Tris HCl pH 7.5, 137-mM NaCl, 50-mM NaF, 0.5% NP40, 2-mM EDTA, 0.017 mg/ml aprotinin, 1-mM phenylmethylsulfonyl fluoride (PMSF), 1-mM Na3VO4. Store at 4°C. Aprotinin (Sigma), PMSF, and Na3VO4 are added to the lysis buffer from 100x stock immediately before use. 8. PMSF 100x stock solution (100 mM): Dissolve 174.2 mg PMSF in 10-ml ethanol. Aliquot and store at 20°C. Add 1:100 to NP40 lysis buffer. 9. Na3VO4 100x stock solution (100 mM): Dissolves 184-mg Na3VO4 in 10-ml water, adjust to pH 10.0 with NaOH and HCl, boil until it turns colorless, and re-adjust pH to 10.0. Aliquot and store at 20°C. Add 1:100 to NP40 lysis buffer. 10. 2x SDS loading buffer: 950 ml Laemmli sample buffer (#1610737, BioRad, Hercules, CA) is combined with 50 ml betamercaptoethanol (#M3148, Sigma) to make 2x SDS loading buffer. Store at room temperature. 11. Western stripping buffer: We use either 0.2x NaOH solution or Restore Western blot stripping buffer (Pierce Biotechnology, Inc., Rockford, IL) and both are effective at completely removing signal from western membranes. 2.4. Removal of Testes
1. 20x Modified Barth’s Solution (MBS) Solution: 1.76-M NaCl, 20-mM KCl, 48-mM NaHCO3, 16.4-mM MgSO4, 200-mM Hepes, 6.6-mM Ca(NO3)2 4H2O, 8.2-mM CaCl2 6H2O, pH 7.4. Store the 20X stock solution at 4°C. Sterilize by filtration. Dilute in distilled water to make 1X or 0.1X working solutions. Working solutions are stored at room temperature. 2. Testes solution: 1X MBS containing 20% fetal calf serum (FCS, Sigma) and 1x penicillin/streptomycin. Sterilize by filtration. Store at 4°C. 3. Tricane methanesulfonate (MS-222, Sigma). 4. Penicillin/Streptomycin 100X stock solution (#P0781, Sigma).
2.5. Microinjector Apparatus
1. Microinjector (PLI-100, Harvard Medical Systems Corp, Greenvale, NY). 2. Joystick micromanipulator (MN-151, Narishige, Japan). 3. Micropipette puller (P30, Sutter Instrument Co., Greenvale, NY). 4. Filamented borosilicate glass capillaries (BF100-50-10, Sutter Instrument Co., Greenvale, NY).
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5. Compact stereomicroscope (SMZ-2B, Nikon) with fiber optic cold light source (KL1500, Schott, Elmsford, NY). 2.6. In Vitro Fertilization of Embryos
1. Human chorionic gonadotropin (CG-10, Sigma) is dissolved in 10-ml sterile water to yield a final concentration of 1000 IU/ml and stored at 4°C. 2. Embryo medium: 0.2X MBS solution, 1X penicillin/streptomycin. Sterilize by filtration. Store at 4°C and warm up to room temperature before use. 3. Cysteine solution: Dissolve 2-g/L-cysteine (#168149, Sigma) in 100-ml distilled water, adjust to pH 7.6–7.8 with NaOH. Prepare immediately before use and keep at room temperature.
3. Methods 3.1. Removal of Xenopus Testes
1. Anesthetize a sexually mature male frog by placing it in a container with 0.3% tricane methanesulfonate (MS-222, Sigma) for >20 min. Euthanize by cervical translocation with a wire cutter. 2. Make a lower abdominal sagittal incision and remove the testes. The testes are cream-colored oval structures that are attached to the anterior, ventral surface of the kidneys. 3. Rinse the testes in 1X MBS and transfer to a 50-ml conical tube containing 10-ml testis solution. Store at 4°C. These testes can stay fresh up to 2 weeks for use in in vitro fertilization experiments.
3.2. Preparation of In Vitro Embryos
1. The evening prior to spawning, inject 800 IU of human chorionic gonadotropin (CG-10, Sigma) into the dorsal lymph sac of a sexually mature female frog. While eggs from one frog may be enough for an entire day’s experiments (up to 8000 eggs laid in a day), it may be best to spawn multiple frogs (we spawn three at a time) to ensure that at least one frog lays eggs of sufficient quality for in vitro fertilization experiments. 2. Female frogs are maintained between 14°C and 19°C overnight. It takes longer to start spawning if a frog is kept at a colder temperature. In general, it takes approximately 10–12 h to start spawning at 19°C, 11–13 h at 16°C, and >15 h at 14°C. 3. Once a frog starts spawning, collect the eggs into a 100-mm petri dish containing 1x MBS. 4. Cut off about one-quarter of a testis with a razor blade and homogenize it in 1-ml 1X MBS in a 1.5-ml eppendorf tube.
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The 1X MBS is removed from the petri dish containing the eggs, and approximately 250 ml of the sperm solution is applied to the eggs. Mix well and keep at room temperature for 4 min to allow fertilization to occur. 5. Fill the petri dish with 0.1X MBS. 40 min later, pour off the 0.1X MBS and replace with 2% cysteine solution. Place the petri dish on a rotating platform with slow agitation for 2–4 min until the jelly coat outside the embryos is dissolved. 6. Pour off the cysteine solution and wash five times with 0.1X MBS. After the last wash, replace the 0.1X MBS solution with embryo medium. Keep the embryos in a 19°C incubator before microinjection. 3.3. Setting Up the Microinjector Apparatus
1. These instructions assume the use of a Harvard Medical System #PLI-100 microinjector system. They are easily adaptable to other constant-pressure microinjectors that use compressed nitrogen gas to propel ejection from the micropipette. 2. Assemble the microinjection apparatus, which includes the following parts: a microinjector (PLI-100, Harvard Medical Systems corp, Greenvale, NY); a joystick micromanipulator (MN-151, Narishige, Japan) for mounting the micropipette; a compressed nitrogen gas tank to supply air pressure in the microinjector; two foot paddles to control aspiration and ejection from the micropipette. Adjust the setting on the PLI-100 to the following: Pclear ¼ 580 kPa, Pbalance ¼ 4 kPa, injection time ¼ 500 ms. Do not change the settings once the micropipette is calibrated. 3. Prepare the microinjection needles (micropipettes) used for penetrating embryos and delivering antisense morpholinos in the following way: filamented borosilicate glass capillaries (BF100-50-10, Sutter Instrument Co., Greenvale, NY) can be vertically pulled with a Model P-30 vertical micropipette puller (Sutter Instrument Co., Greenvale, NY) to generate micropipettes for microinjection experiment. Adjust the settings on the P-30 micropipette puller to Heat 999, Pull force 999. Cut micropipettes with a razor blade 1 mm from the tip to make an opening with a diameter approximately 0.3 mm. 4. Place a compact stereomicroscope (SMZ-2B, Nikon) immediately next to the microinjector to monitor microinjection under magnification. Use a fiber optic cold light source (#KL1500, Schott, Elmsford, NY) to provide ample lighting without increasing the temperature on the microinjection platform. 5. Prepare the microinjection platform in the following way: cover a microscope glass slide with a layer of parafilm and place it face up at the base of the microscope. Embryos will be microinjected on the parafilm surface of the slide.
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6. Calibrate micropipettes in the following way: transfer 1 ml of RNAase-free distilled water onto the parafilm surface. Aspirate as much water as possible into the micropipette, being careful to avoid aspirating air into the micropipette. Eject the water from the micropipette by repeatedly pressing the ejection foot paddle. Count the total number of times the paddle was pressed. Ejection volume can be calculated by the following equation: Ejection volumeðnlÞ ¼ 1000 nl=Number of ejections To change the ejection volume, adjust ‘‘injection time’’ on the microinjector proportional to the desired change of the ejection volume. Ejection volume is linearly proportional to injection time within the range of 100–900 ms. 7. Load the micropipette with morpholino solution: transfer 1 ml of morpholino solution onto the parafilm surface. Aspirate as much morpholino solution as possible into the barrel of the micropipette while avoiding air. Transfer 100 ml of RNAasefree distilled water onto the parafilm surface, and dip the micropipette in water to prevent the tip from drying. The microinjector apparatus is now ready for microinjection experiments. 3.4. Microinjection of Morpholinos into Embryos
At room temperature, it takes approximately 90 min for fertilized eggs to enter cleavage stage, 120 min to enter two-cell stage, and 150 min to enter four-cell stage (9). Depending on the particular experiment, select the appropriate embryonic stage of embryos for morpholino injection. For global protein knockdown, morpholinos can be injected bilaterally into cleavage stage or two-cell-stage embryos. To knockdown protein expression on only one side of the embryos, morpholinos can be injected into only one of the two blastomeres in two-cell embryos, or two same-side blastomeres in four-cell embryos (see Note 2). For Western blots, use 1–4 embryo-equivalents of protein lysates per lane on SDS-polyacrylamide gels. It is best to combine protein extracts from >10 embryos. More embryos are often needed if they are used for scoring gross phenotypes (n>100), or for in situ hybridization experiments (n>20). The following protocol for morpholino microinjection is adapted from those published by Moon and Christian (10): 1. Immediately before microinjection, transfer embryos from embryo medium to injection solution (see Note 3). 2. With a transfer pipette (#13-711-5A, Fisher), transfer 10–20 embryos onto the parafilmed glass slide. All embryos should be submerged in the injection solution.
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3. Gently insert the micropipette tip approximately 0.5-mm deep inside the embryos. Inject the desired amount of morpholinos by pressing the ejection paddle. In one swift motion withdraw the micropipette from the embryo and proceed to the next embryo. 4. With the non-operating hand, use a fine forceps or a p200 pipette tip to nudge the embryos towards the micropipette for microinjection. 5. After microinjection, keep the injected embryos in injection solution for 1 h to allow the embryos to heal. After 1 h, transfer the embryos to embryo medium (see Note 4).
3.5. Evaluation of Morpholino Effect on Protein Levels by Immunoblotting
1. Maintain morpholino-injected embryos in embryo medium at 19°C for at least 12 h and up to 96 h before harvesting the proteins. 2. Transfer approximately 10 embryos into a 1.5-ml eppendorf tube. Aspirate as much solution as possible so that only the embryos remain in the tubes. Quick freeze with liquid nitrogen for >2 min. The embryos can be stored in 80°C freezer for an extended period of time. 3. If the injected morpholino target mRNAs that encode nonmembrane-associated proteins, lyse the embryos or oocytes in a low-stringency NP40 lysis buffer. Use 10–20 ml lysis buffer per embryo. If the morpholino target membrane-bound proteins, a lysis buffer that contains a stronger detergent is preferred. 4. Homogenize embryos manually with a dounce homogenizer (20 strokes), followed by repeated pipetting (20 times) with a p200 Pipetman. 5. Centrifuge the embryo homogenate at 13,000 rpm for 10 min at 4°C. Remove the supernatant into a new 1.5-ml eppendorf tube. This embryonic protein lysate may be stored at 80°C for up to 6 months. 6. Quantify and normalize the amount of protein in each sample with BioRad protein assay reagent (#500-0006, Biorad). Mix 2 ml of protein assay reagent with 8 ml of distilled water to create the 1X working solution. Mix 5 ml of Xenopus embryonic protein lysate with 400 ml of the working solution, and read light absorbance at 595-nm wavelength with a standard spectrophotometer. 7. Prepare, load, and run SDS-PAGE gels with standard molecular techniques. Proteins with size around 30 kDa (the size of 14-3-3 proteins) are best separated with a 12% gel.
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8. Transfer the SDS-polyacrylamide gel onto nitrocellulose membrane with standard molecular techniques. Block with 1X TBS/T containing 5% milk for 1 h at room temperature. 9. Incubate with 10-ml 14-3-3 family-member-specific primary antibody solution in a small sandwich box with agitation overnight at 4°C. After overnight incubation, wash blots with 1X TBS/T three times at room temperature, 15 min each time. 10. Incubate blots with HRP-conjugated secondary antibody for 45 min at room temperature. After secondary antibody incubation, wash with 1X TBS/T three times for 15 min each time. 11. Visualize 14-3-3 bands with an ECL Western Blotting Detection Reagent (Amersham Biosciences, Buckinghamshire, England) in an X-ray film cassette. 12. Once a satisfactory exposure on the X-ray film is obtained, the immunoblot membrane is stripped and reprobed with an antibody that recognizes total ERK proteins. This provides a loading control that confirms equal loading. Add 20-ml stripping buffer (0.2-N NaOH, or ‘‘Restore’’ stripping buffer, Pierce Biotechnology, Inc., Rockford, IL) to the blot for 20 min with constant agitation. Once the blot is stripped, extensively wash with 1X TBS/T 15 min three times, and then incubate overnight with primary antibody that recognizes total ERK proteins, followed by washes, secondary antibody, and ECL detection as above. 13. Bands on scanned immunoblots are quantified by use of the computer program Image J (NIH, Bethesda, MD). The target 14-3-3 band densities are normalized to the amount of protein loaded in each lane (i.e., total ERK).
3.6. Phenotype Analysis of Embryonic Development
The developmental processes in morpholino-injected embryos can be observed with a dissecting microscope. 14-3-3 "- and tmorpholino-injected embryos develop prominent gastrulation defects and fail to undergo mesoderm specification. These phenotypes are assessed by visual inspection of gross embryos and in situ hybridization with gene-specific RNA probes. 14-3-3 "- or t-morpholino-injected embryos also undergo extensive apoptosis at the start of gastrulation. These are assessed by an ELISA apoptosis assay (Roche Applied Science, Indianapolis, IN). For gross morphologic evaluations, embryos are observed under light microscope every 2–3 h after microinjection. Exogastrulation in 14-3-3 t or "-morpholino-injected embryos becomes
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apparent at stage 10.5 (approximately 11 h post-fertilization at room temperature). A loss-of-eye phenotype in 14-3-3 g-morpholino-injected embryos can be assessed in tadpole-stage embryos (2 days post-fertilization). 3.7. Analysis of Marker Gene Expression in Microinjected Embryos by In Situ Hybridization
1. Preparation and storage of embryos prior to in situ hybridization: after morpholino injection, allow albino embryos to develop to the desired stage(s). Fix in 4% paraformaldehyde and keep in 100% ethanol for up to 6 months. 2. Preparation of templates and probes: The DNA template is linearized with a restriction enzyme, extracted with phenol/ chloroform, and precipitated prior to use. Digoxigenin-labeled probes are made with mMessage (Roche Applied Sciences, Indianapolis, IN) by the method of Sive (11). 3. Incubation of embryos with in situ hybridization probes: Embryos are rehydrated, permeabilized with proteinase K, and hybridized overnight with RNA probe as described (11). 4. Visualization of gene expression pattern: Embryos are washed, incubated with anti-digoxigenin antibody and subjected to chromogenic reaction by the method of Sive (11).
4. Notes 1. Alternative design of morpholino antisense oligos: Some other investigators have used an alternative approach to design antisense morpholinos in Xenopus (12). Morpholinos are made antisense to the 50 UTR region of targeted genes. Since Xenopus laevis is a pseudo-tetraploid organism, there may be two slightly different 50 UTR sequences for the same gene. The morpholino is made antisense to an identical region in both 50 UTR sequences. If such a region does not exist, two separate morpholinos, each targeting one of the 50 UTRs, are combined and injected together to knockdown the expression of a particular gene. 2. Injection of embryos before they enter the cleavage stage is not recommended because this frequently results in embryonic death. Furthermore, injection of morpholinos at high doses may result in the development of non-specific abnormalities such as cell cycle arrest or exogastrulation. Therefore, it is recommended that the injection dose be kept under 20 pmol/embryo and the injection volume be kept under 20 nl/embryo. 3. The Ficoll and high salt content prevent leakage of cytosolic contents during and after microinjection.
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4. Do not keep embryos in injection solution past stage 8 (approx. 5 h post-fertilization). Embryos left in high-salt media during gastrula stages develop exogastrulation defects and subsequently die.
References 1. Muslin, A. J., Xing, H. (2000) 14-3-3 proteins: regulation of subcellular localization by molecular interference. Cell Signal. 12, 703–9. 2. Fu, H., Subramanian, R. R., Masters, S. C. (2000) 14-3-3 proteins: structure, function, and regulation. Annu. Rev. Pharmacol. Toxicol. 40, 617–47. 3. Aitken, A. (2006). 14-3-3 proteins: a historic overview. Semin. Cancer Biol. 3, 162–72. 4. Muslin, A. J., Tanner, J. W., Allen, P. M., Shaw, A. S. (1996) Interaction of 14-3-3 with signaling proteins is mediated by the recognition of phosphoserine. Cell 84, 889–97. 5. Yaffe, M. B., Rittinger, K., Volinia, S., Caron, P. R., Aitken, A., Leffers, H., Gamblin, S. J., Smerdon, S. J., Cantley, L. C. (1997) The structural basis for 14-3-3: phosphopeptide binding specificity. Cell 91, 961–71. 6. Lau, J. M., Wu, C., Muslin, A. J. (2006) Differential role of 14-3-3 family members
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in Xenopus development. Dev. Dyn., 235, 1761–76. Obsil, T., Ghirlando, R., Klein, D. C., Ganguly, S., Dyda, F. (2001) Crystal structure of the 14-3-3 zeta:serotonin N-acetyltransferase complex. A role for scaffolding in enzyme regulation. Cell. 105, 257–67. Wu, C. , Muslin, A. J. (2002) Role of 14-3-3 proteins in early Xenopus development. Mech. Dev. 119, 45–54. Nieuwkoop, P. D., Faber, J. (1967) Normal Table of Xenopus laevis, North-Holland Publishing Company, Amsterdam. Moon, R. T., Christian, J. L. (1989) Microinjection and expression of synthetic mRNAs in Xenopus embryos. Technique – J. Methods Cell. Mol. Biol.. 2, 76–89. Sive, H. L., Grainger, R. M., Harland, R. M. (2000) Early Development of Xenopus laevis: A Laboratory Manual, Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY. Heasman, J. (2002) Morpholino oligos: making sense of antisense? Dev. Biol. 243, 209–14.
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Chapter 4 A Microinjectable Biological System, the Xenopus Oocyte, as an Approach to Understanding Signal Transduction Protein Function Katia Cailliau and Edith Browaeys-Poly Abstract To study protein function in cellular signaling, manual microinjection is a direct technique, but limited by the small size of many cells. The giant vertebrate cell, the Xenopus laevis oocyte, is a perfect model system to perform these studies. Oocytes are numerous and synchronous cells, arrested in the G2 phase of the cell cycle and easily amenable to biochemical, electrophysiological, and cytological studies. We describe how to microinject proteins or peptides in this model and we study, as an example, the Grb2 transduction cascade. Key words: Microinjection, peptides, proteins, Xenopus laevis oocytes, cellular signaling.
1. Introduction Signal transduction pathway proteins are involved in pleiotropic cellular functions and in several pathological processes (1–4). These proteins, particularly receptors, belong to large families and display numerous signaling cascades that increase the difficulty of understanding their role. Several technological approaches are possible to dissect these cascades and are mainly based on the loss and rescue function of the targeted protein. Some somatic cell systems are approached by introduction of foreign DNA or RNA molecules (5,6) or by permeable recombinant proteins (7,8). However these techniques remain long and fastidious to perform. A more rapid approach would be to directly microinject mimetic peptides, dominant-negative proteins, pharmacological tools, or antibodies to affect the protein function and David J. Carroll (ed.), Microinjection: Methods and Applications, Vol. 518 Ó 2009 Humana Press, a part of Springer ScienceþBusiness Media, LLC DOI 10.1007/978-1-59745-202-1_4
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to record the induced effects. For most somatic cells, their small size is a constraint for these studies and renders microinjection difficult. However, giant cells that are easily microinjectable exist: the germinal biological system, the Xenopus oocyte. These vertebrate cells have a large size (diameter > 1.2 mm, for fully grown oocytes), are numerous, synchronous, have the capacity to survive in vitro for a long time, and display a high translational capacity (9), properties that permit many types of study. In addition, they are amenable to cytological, electrophysical, and biochemical manipulations and have been extensively used to study the nongenomic signaling cascade of the cell cycle, particularly the G2/M transition. Xenopus oocytes are physiologically arrested at the G2 stage of the first meiotic prophase. Progesterone, the natural inducer, reinitiates their entry into the M phase and leads to a germinal vesicle break-down (GVBD) used as an indicator of the maturation process (the transition G2/M). Insulin or growth factors can also induce GVBD when binding to the appropriate receptor. Insulin and IGF1 receptors are naturally present in oocytes, and exogenous growth factor receptors can be expressed by injection of their respective mRNA (10–13). Here, we describe how to manipulate and microinject Xenopus oocytes with various peptide or protein samples to analyze their positions and roles in cascade transduction pathways. Using inhibitory peptides and dominant-negative Grb2 proteins, we showed that microinjected Grb2 protein initiates a Ras-dependent cascade, requiring SH2 and SH3 domains, without external stimulation (14).
2. Materials 2.1. Frogs
Mature Xenopus laevis females (preferably around 2 years) were ordered from the ‘‘Centre de ressources biologiques Xe´nope’’ (Universite´ de Rennes, France). Frogs were acclimatized in our laboratory at least 3 weeks before oocyte handling.
2.2. Media and Solutions
1. ND96 freshly prepared each week is kept at 14°C and in the dark: 96-mM NaCl, 2-mM KCl, 1-mM MgCl2, 1.8-mM CaCl2,, 5-mM HEPES adjusted to pH 7.4 with NaOH, supplemented with 50 mg/ml streptomycin/penicillin, 225 mg/ ml sodium pyruvate, 30 mg/ml soybean trypsin inhibitor, 1 ml/ ml tetracycline. Solutions used in our experiments do not usually require sterilization, but great care should be taken to use clean water and reagents. Tetracycline allows an optimal conservation and a good recovery of oocytes after the various microinjection treatments (15).
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2. High-salt stock solution: 2.6-M NaCl, 0.5-M KCl, 0.2-M MgCl2, 0.2-M CaCl2. 3. Tetracycline stock solution: 50 mg/ml in ethanol 50% stored at –20°C. 4. Anesthetic solution: MS-222 also called tricaine methane sulphonate (Sandoz). 5. Collagenase solution: 1 mg/ml collagenase A from Clostridium histolyticum (Roche Diagnostics), dissolved in ND96 without tetracycline. 2.3. Reagents/Special Items
1. Oocyte lysis: Eppendorf pipetman, micropipettes (1 ml or 200 ml). 2. Microinjection material: hard glass capillary, mineral oil (Sigma), insulin syringe with a millipore filter 0.45 mm adapted on the syringe. 3. Surgery tools: forceps type Dumont MC 40 (Moria) and scissors cleaned before use with ethanol 70%, and sewing thread. 4. Petri dishes 50 mm. 5. Culture plates, 24 wells.
2.4. Equipment
1. A horizontal micropipette puller (World Precision Instruments) (Fig. 4.1A). 2. A micromanipulator modified to accept the microinjection pipette. At the base of the micromanipulator, we fixed a plastic ring, composed of two parts that are screwed together, that received the microinjection pipette (Fig. 4.1B). 3. A microinjection pipette: capillary micropipette (Nichiryo model 800) with glass capillary 1–5 ml (Nichiryo cap-5) (Fig. 4.1B). 4. A stereomicroscope with a 10-fold magnification permits an adapted magnification for fine interventions.
Fig. 4.1. Equipment for microinjection. (A) Micropipette puller (World Precision instruments), with the glass capillary used for microinjection; (B) Microinjection station equipped with the microinjector, the stereomicroscope, and the optic fiber.
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Fig. 4.1. (continued)
5. A fiber optic light source that provides sufficient illumination and avoids oocyte desiccation during the procedure. 6. Two incubators to keep injected oocytes at 19°C and to maintain ovaries at 14°C. 7. Plastic transfer pipettes, single use, 3 ml (Dutscher, France).
3. Methods 3.1. Frog Maintenance
Before keeping and utilizing Xenopus laevis in the laboratory, authorizations should be obtained from the concerned authorities. Adults are maintained at room temperature on a 12 h light/ dark cycle in large plastic tanks. A continuously filtered tap water circuit furnishes the tanks and avoids daily cleaning. The cleaning is usually performed once a week after the animals are nourished with granules commonly used for trout.
3.2. Ovary Removal and Oocyte In Vitro Handling
1. Female Xenopus laevis are anesthetized by placing them in a 1-liter beaker of MS-222 (1 g/l) for about 45 min. The beaker is covered to prevent escape.
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Fig. 4.2. Xenopus laevis should be incised as indicated to remove one ovary.
2. Once completely sedated, the animal is washed with soap, rinsed with tap water, and placed on its back on clean aluminum foil. 3. The skin of the lateral part of the abdomen is kept tight with a forceps, in order to perform a small incision (about 1 cm) with scissors (Fig. 4.2). Depending on the amount of oocytes needed, only one ovary can be removed. In this case the second can be removed 3–4 weeks later. 4. The underlying abdominal wall is also cut to allow the ovary to be excised. The ovary appears as bags containing various stages of oocyte development, wrapped in connective tissue with blood vessels. 5. The ovary lobes are carefully dissected away and washed 4 times to remove the remaining blood in a ND96 petri dish and stored covered at 14°C. 6. From this point onwards, it is vital to keep the ovaries submerged in ND96 complemented with antibiotics all times, including during the following transfers. This permits them to be kept for at least a week. In our experience, Xenopus laevis only produce low-quality oocytes during the summer. 7. The abdominal wall and the skin are separately sutured using sewing thread with 3–4 stitches first on the muscles and then on the skin. 8. The frogs are each placed in a beaker without water and their skin moistened with tap water until they recommence walking. The beaker is again covered to prevent escape.
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9. The tank is filled with tap water where the animal remains for 1 week before going to a collective tank (about 10 animals). 3.3. Oocyte Isolation
1. Groups of 5–10 oocytes are separated from small parts of the ovary lobes using two forceps under the stereomicroscope and immersed in ND96. A close-up view of an opened bag shows oocytes at different stages (16). 2. Oocytes from the final stage, stage VI, displaying the largest diameter (>1.2 mm), are used in our experiments. These oocytes are pigmented brown in one hemisphere (animal hemisphere). The other hemisphere (vegetal hemisphere) shows the yellow color of the egg yolk. The equatorial region sometimes presents a ring. To avoid confusion with other stages, stage V oocytes appear smaller, darker in their animal hemisphere and never show an equatorial ring. 3. Oocytes incubated in a collagenase solution are gently shaken for 45 min on an agitator to easily remove the surrounding follicular cells, and the connective tissue containing blood vessels. Oocytes are washed 3–4 times with ND96 before being kept at 14°C in fresh ND96 medium. We do not use a stronger or a longer collagenase treatment due to the risk of destroying oocyte surface proteins and causing poor viability. Collagenase treatments can induce spontaneous GVBD. 4. At this step oocytes are stored some hours before defolliculation is manually achieved by dissection. Waiting 3–4 h renders the process easier. A small group of oocytes is selected, held with one forceps and the stage VI oocyte is carefully dislodged from the surrounding tissue with a second forceps. 5. The oocytes are sorted and the healthiest are kept at 19°C in the final ND96 medium. 6. Successful microinjection depends upon selecting the correct defolliculated oocytes. Stage VI oocytes are the largest ones in the sample and show good contrast between the brown animal hemisphere and the creamy-colored vegetal hemisphere. Oocytes which are ‘‘jelly-like’’ or which have indistinct poles should be discarded. Healthy oocytes should be smooth, quite firm, and should not burst too easily when gently manipulated.
3.4. Preparation of Microinjection Pipettes
1. Good micropipettes are essential to the success of microinjection. The capillary micropipette must be large enough to deliver the desired volume, but not too fine as it will often get clogged. We use a micropipette horizontal puller to taper small glass capillary tubes. Different pulling conditions have to be tried to find the best one. We always ensure obtaining a large micropipette of about 5–6 cm for use and a smaller micropipette that will be discarded (Fig. 4.1A). The capillary tip
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diameter should be about 20 mm, or the filling will not succeed. Short capillaries will not fit on the microinjection pipette piston. 2. Under a stereomicroscope, the extreme tip of the capillary is broken up using forceps, to create a blunt end. 3. Backfill capillary with mineral oil using an insulin syringe equipped with a millipore 0.45-mm filter mounted between the syringe needle and core. This pipette is a nanoliter injector based on the displacement of a mineral oil/aqueous sample interface by a small metal piston. 4. The capillary micropipette is mounted onto the microinjection pipette. It is gently placed on the thin metal piston of the microinjection pipette and tightly screwed into place. Some mineral oil will come out of the capillary tip during the procedure and afterwards it will be necessary to remove about 60–80 nl more oil by turning the micropipette knurl (Fig. 4.1B). 5. The protein or peptide solution, with a volume as small as 0.5 ml, is dropped on Saran wrap or in a petri dish and immediately transferred into the capillary pipette. 6. Under the stereomicroscope, the capillary tip is pushed into the drop before filling by turning the microinjector button backwards. 7. A magnification of 10 will be sufficient for all the microinjection experiments. 8. A pulse of 20–40 nl from the sample is ejected from the capillary tip to verify that the system is ready for microinjection. 3.5. Oocyte Microinjection
1. Microinjections are performed in petri dishes with the bottom scraped in a pattern of squares (with forceps) to generate an adhesive surface for the oocytes and filled with ND96. Most of the time oocytes will stick to the bottom of the plastic dish (Fig. 4.3A). After defolliculation, a delay of 1–2 h at 14°C is necessary for oocyte recovery before microinjection. 2. Several oocytes can be lined up along a scraped lane and injected one after another with the capillary micropipette at an angle of 45° (Fig. 4.3B). 3. The oocytes are injected in the equatorial zone, just below the pigmented animal zone. This protocol allows the optimal diffusion of the sample in the oocyte cytoplasm and avoids the alteration of the nucleus in the animal hemisphere with the glass micropipette tip (Fig. 4.3B). 4. Only the thinnest part of the glass capillary tip is inserted into the oocyte, for about 150–200 mm inside, to avoid the formation of a large hole that could prevent oocyte recovery.
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Fig. 4.3. (A) Petri dish prepared for microinjection. (B) Diagram illustrating how to inject proteins into Xenopus laevis oocytes.
5. The optimal injected volume is 60 nl but volumes up to 100 nl or 120 nl can be injected very slowly when needed. For these larger volumes a control using distilled water instead of the sample is also recommended. Once the glass capillary tip is inserted into the oocyte it should not be withdrawn before 5–10 s to avoid sample escape. 6. After capillary tip removal oocytes will sometimes leak a small amount of vitellus but this will not affect their survival rate. 7. After 2–3 microinjections are performed, it will probably be necessary to bring the glass capillary out of the bath solution and generate a small pulse to ensure the tip is not clogged with vitellus platelets. If this is the case the capillary tip should again be broken with forceps or replaced by a new one when the tip exceeds 30 mm in diameter. 8. After the oocytes are injected, the petri dish is manually displaced and a new oocyte is injected with the same procedure; this avoids losing the micropipette set-up and also avoids the displacement of the injected oocyte. Moreover, in this setting the capillary pipette can be changed easily and will not need a new calibration. 9. Microinjections are performed in an organized manner: this will allow the investigator to keep track of injected oocytes
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with minimal effort and optimal rapidity. When a second injection has to be done the same day on the same oocyte, the initial hole should be re-used if found. The total injected volume should not exceed 120 nl. 10. The oocytes are gently transferred to petri dishes or culture plates (24 wells) filled with fresh ND with a 3-ml largediameter plastic transfer pipette. The medium of injected oocytes is replaced when necessary. 11. The quantity of injected protein or peptide will vary according to their toxicity. We commonly microinject about 100 ng and up to 400 ng of dominant-negative protein or mimetic peptide. The ratio of the number of molecules injected should be taken into account when competition experiments are realized between two samples. The final concentration of every sample can be determined assuming the total volume to be 1 ml for a stage VI oocyte. 12. We recommend using three batches of 15–20 oocytes from three different animals per experiment to avoid non-specific results.
4. Notes 1. After defolliculation or microinjection, selecting oocytes is an important step and has to be performed very carefully to obtain the best results. Under the stereomicroscope a fine selection of oocytes is performed, all oocytes that present defects are removed: those that do not present a good contrast between the brown and cream halves, and discolored or damaged oocytes. Damaged and lysed oocytes release cytoplasm into the medium and this affects the survival of other oocytes. 2. Frequently, recombinant proteins are produced from cDNA inserted into a pGEX vector flanked by a GST tag. Grb2, Grb2-GST, and mutated Grb2 were prepared as described in Ref. (17). Considering that the GST tag can produce nonspecific results, ideally it should be removed, using thrombin cleavage. The subsequent removal of thrombin from the protein is performed easily in small centricon microfuge tubes. Some recombinant proteins are available from manufacturers. Two disadvantages should be noted in the use of these proteins. The low final concentration of the sample renders the microinjection of high concentrations into oocytes impossible. The storing buffer can sometimes be toxic for oocytes and requires dialysis.
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Most of the time, samples are resuspended in deionized water and should not be resuspended in any buffer such as PBS or ND96 that are toxic for the oocytes. When ethanol or DMSO are present in the samples, they have to be diluted at least to a ratio of 1/1000 or they will spontaneously trigger GVBD. Synthetic peptides offer the advantage of incorporating posttranslational modifications such as phosphorylation, highly useful for the study of signal transduction cascades. Disadvantages include the cost, as it is sometimes necessary to synthesize several peptides encompassing an interacting or a functional domain to generate the most efficient blocking peptide. 3. When mimetic or inhibitory peptides or proteins are injected, a minimum 1 h incubation should be included before oocyte stimulation to allow for sample distribution in the vitellus-rich cytoplasm. 4. To study receptors, cRNAs are microinjected and expressed for 48 h according to the same microinjection method, and oocytes are kept in ND96 at 19°C before any peptides or proteins are further microinjected (10,11). Sometimes it is useful to co-inject two samples at the same time: in these cases, small drops of each sample can be mixed on a petri dish before they are loaded in the capillary micropipette. 5. GVBD determination. The oocyte nucleus is localized closer to the animal pole and called the germinal vesicle. When the maturation process begins, the germinal vesicle migrates close to the oocyte surface and temporarily displaces the pigment from the underneath surface generating a white spot at the animal pole, defined as the first phase of the GVBD process, after 6–15 h (Fig. 4.4). Oocytes exhibiting these white spots are counted and plotted on graph for statistical analysis (Fig. 4.5A). After the second phase of the process has occurred, the nuclear envelope breaks down, the pigment is replaced, and the white spots disappear. At this time the metaphase spindles forms. When the determination of the GVBD is
Fig. 4.4. External morphology of stage VI Xenopus laevis oocyte. (A) Immature oocyte of stage VI; (B) Stage VI oocyte that presents a white spot indicating GVBD.
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Fig. 4.5. Role played by the SH2 and SH3 domains of Grb2 in the Xenopus oocyte-induced G2/ M transition. Oocytes were injected with Grb2 (100 ng), mutated proteins Grb2-P49L (N-SH3) or Grb2-G203R (C-SH3) (100 ng), SH2-Grb2 domain (160 ng), or phosphorylated peptide (PP) with a high affinity for Grb2 (PSpYVNVPN, 10 mM). SH2-Grb2 domains and PP were injected at least 45 min before Grb2. (A) Percentage of GVBD observed after microinjection shows that the induction of meiosis requires the SH2 and SH3 domains of Grb2. (B) Western blot analysis of Erk2 phosphorylation state shown by the upper band. Fifteen hours after microinjection with Grb2 and mutated Grb2 proteins or competing SH2 domain, oocytes were homogenized and electrophoresed on a 15% modified polyacrylamide gel (30% acrylamide and 0.2% bisacrylamide). Proteins were transferred to a hybond ECL membrane (Amersham Life Sciences). Membranes were incubated 1 h with an anti-ERK2 antiserum (Santa Cruz Biotechnology 1/ 2500) in TBS (Tris-HCL 15 mM, NaCl 150 mM, Tween 20.1%, pH 8 containing 10% bovine serum albumin). Grb2 triggers the phosphorylation of ERK2, only when both SH2 and SH3 domains are functional. The use of mutated protein or a competing domain abolished this effect. (Reproduced from Ref. (14) with permission from Elsevier Science).
uncertain, oocytes are boiled for 5 min at 100°C in water and opened up at the animal pole with two forceps under a stereomicroscope to determine the presence (or the absence) of the germinal vesicle which appears as a hard white globe. 6. Effects of microinjections are easily studied in oocytes by separating cytoplasmic and membrane fractions at different
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time-points or after the GVBD has occurred, and analysis by immunoprecipitation or Western blotting (Fig. 4.5B). For these experiments, oocytes are homogenized in the ice cold buffer: 50-mM Hepes, pH 7.4, 500-mM NaCl, 0.05% SDS, 5mM MgCl2, 1 mg/ml bovine serum albumin, 10 mg/ml leupeptin, 10 mg/ml aprotinin, 10 mg/ml soybean trypsin inhibitor, 10 mg/ml benzamidine, 1-mM PMSF, 1-mM sodium vanadate. Back and forth movements generated by a micropipette tip (100, 200 ml) are used to break the oocytes. Centrifugation at 10,000 g for 15 min removes the lipids (upper phase) and the membrane fraction (bottom phase). For membrane protein enrichment, the bottom phase is resuspended in the buffer described before added with 1% Triton X-100 and again homogenized vigorously. A final centrifugation removes oocyte debris (at the bottom) and some remaining lipids (at the surface).
Acknowledgments We thank Dr. R. Pierce (Pasteur Institute, Lille) for reading the manuscript. This study was supported by two successive grants from the ‘‘Ligue Contre le Cancer, Comite´ de l’Aisne’’ and the ‘‘Ligue Contre le Cancer, Comite´ du Nord’’ to KC and EBP.
References 1. Lowenstein EJ, Daly RJ, Batzer AG, Li W, Margolis B, Lammers R, Ullrich A, Skolnik EY, Bar-Sagi D, Schlessinger J. (1992) The SH2 and SH3 domain-containing protein GRB2 links receptor tyrosine kinases to ras signaling. Cell 70, 431–442. 2. Downward J. (1994) The GRB2/Sem-5 adaptor protein. FEBS Lett. 338, 113–117. 3. Chardin P, Cussac D, Maignan S, Ducruix A. (1995) The Grb2 adaptor. FEBS Lett. 369, 47–51. 4. Maignan S, Guilloteau JP, Fromage N, Arnoux B, Becquart J, Ducruix A. (1995) Crystal structure of the mammalian Grb2 adaptor. Science 268, 291–293. 5. Heiser W.C. Ed. (2003) Nonviral Gene Transfer Techniques, Gene Delivery to Mammalian Cells in Methods in Molecular Biology, vol. 245. Humana Press.
6. Heiser W.C. Ed. (2003) Viral Gene Transfer Techniques, Gene Delivery to Mammalian Cells in Methods in Molecular Biology, Vol. 246. Humana Press. 7. Dunican DJ, Doherty P. (2001) Designing cell-permeant phosphopeptides to modulate intracellular signaling pathways. Biopolymers 60, 45–60. 8. Lindgren M, Hallbrink M, Prochiantz A, Langel U. (2000) Cell-penetrating peptides. Trends Pharmacol. Sci. 21, 99–103. 9. Gurdon JB, Lane CD, Woodland HR, Marbaix G. (1971) Use of frog eggs and oocytes for the study of messenger RNA and its translation in living cells. Nature 233, 177–182. 10. Browaeys-Poly E, Cailliau K, Vilain JP. (1998) Fibroblast and epidermal growth factor receptor expression in Xenopus oocytes displays distinct calcium oscillatory
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patterns. Biochim. Biophys. Acta. 1404, 484–489. 11. Browaeys-Poly E, Cailliau K, Vilain JP. (2000) Signal transduction pathways triggered by fibroblast growth factor receptor 1 expressed in Xenopus laevis oocytes after fibroblast growth factor 1 addition. Role of Grb2, phosphatidylinositol 3-kinase, Src tyrosine kinase, and phospholipase C gamma. Eur. J. Biochem. 267, 6256–6263. 12. Cariou B, Perdereau D, Cailliau K, Browaeys-Poly E, Bereziat V, Vasseur-Cognet M, Girard J, Burnol AF. (2002) The adapter protein ZIP binds Grb14 and regulates its inhibitory action on insulin signaling by recruiting protein kinase C zeta. Mol. Cell. Biol. 22, 6959–6970. 13. Liu XJ. Ed. (2006) Xenopus Protocols, Cell Biology and Signal Transduction, Methods in Molecular Biology, vol. 322. Humana Press.
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14. Cailliau K, Browaeys-Poly E, Broutin-L’Hermite I, Nioche P, Garbay C, Ducruix A, Vilain JP. (2001) Grb2 promotes reinitiation of meiosis in Xenopus oocytes. Cell. Signal. 13, 51–55. 15. Elsner HA, Honck HH, Willmann F, Kreienkamp HJ, Iglauer F. (2000) Poor quality of oocytes from Xenopus laevis used in laboratory experiments: prevention by use of antiseptic surgical technique and antibiotic supplementation. Comp. Med. 50, 206–211. 16. Dumont JN. (1972) Oogenesis in Xenopus laevis (Daudin). I. Stages of oocyte development in laboratory maintained animals. J. Morphol. 136, 153–179. 17. Guilloteau JP, Fromage N, Ries-Kautt M, Reboul S, Bocquet D, Dubois H, Faucher D, Colonna C, Ducruix A, Becquart J. (1996) Purification, stabilization, and crystallization of a modular protein: Grb2. Proteins 25,112–119.
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Chapter 5 Combining Microinjection and Immunoblotting to Analyze MAP Kinase Phosphorylation in Single Starfish Oocytes and Eggs David J. Carroll and Wei Hua Abstract The starfish oocyte has proven useful for studies involving microinjection because it is relatively large (190 mm) and optically clear. These oocytes are easily obtained from the ovary arrested at prophase of meiosis I, making them useful as a model system for the study of cell cycle-related events. In this chapter, a method for combining microinjection with immunoblotting of single cells is described. Individual starfish oocytes are injected, removed from the microinjection chamber, and analyzed by immunoblotting for the dual-phosphorylated form of mitogen-activated protein kinase (MAPK). This method will allow for experiments testing the regulation of MAPK in single cells and for the manipulation of these cells by a quantitative microinjection technique. Key words: Mitogen-activated protein kinase, MAPK, oocyte, starfish, echinoderm, meiosis, maturation, fertilization, quantitative microinjection.
1. Introduction In many animals, oocytes are arrested in prophase of meiosis I and incompetent to respond fully to sperm until after meiotic maturation. The immature oocytes are stimulated to re-enter the cell cycle by an external signal that is linked to internal signaling pathways that ultimately activate a maturation promoting factor (MPF), composed of a cyclin–cdc2 complex (1). The mechanisms controlling this meiotic maturation differ between animals, but in a number of cases protein synthesis and/or the activation of mitogen-activated protein kinase (MAPK) are required for MPF activation (1–3). MAPK is an enzyme that plays a pivotal role in mitogenic signaling pathways and in pathways involved in cell differentiation David J. Carroll (ed.), Microinjection: Methods and Applications, Vol. 518 Ó 2009 Humana Press, a part of Springer ScienceþBusiness Media, LLC DOI 10.1007/978-1-59745-202-1_5
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and development (3). MAPK is part of a highly conserved signaling cascade that includes the upstream MAPK kinase (or MEK) and MAPK kinase kinase (or Raf/Mos) (4,5). This pathway is activated by a large number of extracellular stimuli and it targets multiple downstream targets, notably transcription factors and other regulatory proteins (4). These three molecules (MAPK, MEK, and Raf) are Ser/Thr kinases that form the core group of a MAP kinase module found in almost all cell types and throughout the plant and animal kingdoms (5). MAP kinase becomes phosphorylated, and presumably activated, during maturation of oocytes from diverse animal species including mice, frogs, starfish, sea urchins, the nemertean worm Cerebratulus lacteus, Caenorhabditis elegans, and the parchment worm Chaetopterus (2, 6–13). However, the exact role played by MAPK, if any, during oocyte maturation is unclear and may, in fact, vary depending upon the animal or stimulus involved (2, 14–16). In the starfish, MAPK becomes phosphorylated following treatment of prophase I-stage immature oocytes with the hormone 1-methyladenine and then it becomes rapidly dephosphorylated after fertilization (13). There is evidence that MAPK plays a role in regulating transitions that occur during completion of meiosis in the starfish oocyte, and in the transition from meiosis to mitosis that occurs following fertilization (7,13). Additionally, MAPK is also involved in the decision to stimulate apoptosis in eggs that are not fertilized within a certain period of time (7). In this chapter, a method for combining the microinjection of starfish oocytes with an assay to study changes in the phosphorylation state of MAPK in single cells is described. The microinjection method has been described in great detail previously (17), but the method for recovering the injected cells and preparing them for further analysis is novel. The great advantage of this technique is the ease with which the injection volume can be quantified and the ability to perform concentration curves within a single chamber. This method also makes possible the ability to measure a biological phenomenon (such as calcium release or DNA synthesis) in a living cell and then later analyze that exact same cell using biochemical methods such as western blotting.
2. Materials 2.1. Supplies for Oocyte Dissection and Culture
1. Starfish (Asterina miniata) were obtained from either Marinus Scientific Inc. (Garden Grove, CA) or from Santa Barbara Marine Biologicals (Santa Barbara, CA) and maintained in re-
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circulating seawater tanks at 17°C. Each starfish was fed biweekly with approximately half piece of a bay scallop. 2. 1-Methyladenine (Acros Organics, Geel, Belgium). Prepare a 10-mM stock of 1-methyladenine in distilled water. Aliquot and store at 80° C for long-term storage or at room temperature for use within 6 months. 3. Dissection tools from Fine Science Tools, Inc. (Foster City, CA): 3-mm sample corer (Item #18035), Graefe forceps (Item #11051-10), Noyes spring scissors (Item #15012-12). 4. Miscellaneous items such as coverslips, beakers, Pasteur pipettes, etc., can be obtained from your normal supplier. We typically use Fisher Scientific Research (Pittsburgh, PA) or Midwest Scientific (St. Louis, MO). 5. Unless otherwise indicated, chemicals for reagents were purchased from Fisher Bioreagents (Fair Lawn, NJ) or SigmaAldrich (St. Louis, MO). 6. Calcium-free seawater (CFSW) and artificial seawater (ASW) prepared according to the Marine Biological Laboratory (Woods Hole, MA) recipes. CFSW: 436.71-mM NaCl, 9mM KCl, 22.94-mM MgCl2, 25.50-mM MgSO4, 2.14-mM NaHCO3, pH 8.2. ASW: 423-mM NaCl, 9-mM KCl, 9.27mM CaCl2, 22.94-mM MgCl2, 25.50-mM MgSO4, 2.14-mM NaHCO3, pH 8.2. 2.2. Microinjection
1. Standard light microscope with left-hand stage controls rather than the standard stage that has the x–y controls on the righthand side. 2. Narishige SM-20 micromanipulator is recommended (Narishige Scientific Instrument Company, Tokyo, Japan). 3. Horizontal micropipette puller is recommended using a wide filament (6 mm) to produce a long taper on the microneedle. We currently use the PUL-1 puller from World Precision Instruments (Sarasota, FL). 4. Glass capillaries for microneedles (Drummond Scientific, Broomhall, PA, #2-000-100) and for mouth pipettes (Drummond Scientific, #9-000-1061). 5. Scotch double-coated tape 667 (3 M products, St. Paul, MN); available at most office supply stores. 6. Cover Glass No.1, 22-mm square from Corning Labware & Equipment (Corning, NY). Clean and rinse thoroughly before use (17). 7. Dow Corning High Vacuum Grease (Dow Corning Corp., Midland, MI). 8. Microinjection chamber support slide: Machined from ¼’’ plexiglass to same size as a standard microscope slide to have
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a U-shaped cut-out in the middle to make a chamber that is 1/ 16’’ high. This can be ordered from the Instrument Development Lab of the Marine Biological Laboratory in Woods Hole, MA. See reference (17) for more details. 9. FPT Inhibitor III. Farnesyltransferase inhibitor dissolved in water (344152, EMD Chemicals, San Diego, CA). 2.3. Gel Electrophoresis and Western Blotting
1. SuperSignal West Femto Maximum Sensitivity substrate (Catalog number 34095; Pierce Biotechnology, Rockford, IL). 2. Tris-Saline-Tween-20 (TST) for western blotting: 2.42-g Tris and 8.0-g NaCl, 0.1% Tween-20 in deionized water to a total volume of 1 l. 3. Blotto: TST + 5% non-fat dry milk. 4. Primary antibody specific for the dual-phosphorylated form of MAPK. Both the mouse monoclonal anti-phosphoMAPK (#9106, Cell Signaling Technology, Danvers, MA) or the rabbit polyclonal anti-phosphoMAPK (#9101, Cell Signaling Technology) work equally well in the starfish oocyte and egg. 5. ImmunoPure Goat Anti-Rabbit IgG, Peroxidase-Conjugated (#31460, Pierce Biotechnology, Rockford, IL) or ImmunoPure Goat Anti-Mouse, Peroxidase-Conjugated (#31430, Pierce Biotechnology, Rockford, IL). 6. Pierce CL-XPosure film, product #34090 (Pierce Biotechnology, Rockford, IL). 7. Kodak GBX developer (P7042, Sigma-Aldrich, St. Louis, MO) and Kodak Rapid Fixer (P7167, Sigma-Aldrich).
3. Methods 3.1. Oocyte Dissection and Preparation
1. Each starfish has a pair of gonads at the base of each arm. To dissect the ovary or testis, make a small hole on one side of the base of an arm in the aboral side of the starfish using the 3-mm sample corer. In most cases, you will immediately see whether you have a male (white testis) or female (orange-yellow ovary). 2. Using the fine curved forceps, gently remove the testis to an eppendorf tube placed on ice or the ovary to a small (50 ml) beaker filled with CFSW. 3. Using the fine scissors, chop up the ovary in the CFSW. The oocytes will be released into the CFSW. Allow the ovary pieces to settle to the bottom of the beaker. Pour the oocytes into a clean 50-ml beaker to separate them from the ovary pieces.
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Take a small sample of the oocytes and check them under the light microscope. Healthy oocytes will be approximately 185–190 mm in diameter with a uniform cytoplasm. The large nucleus (germinal vesicle) will be clearly visible, as will the nucleolus. The oocyte will be surrounded by clear jelly and a layer of follicle cells. The purpose of the CFSW rinse is to remove the follicle cells which will interfere with microinjection and which will complicate the protein sample for later analysis. 4. After the oocytes settle by gravity to the bottom of the beaker, pour off the CFSW and add filtered natural seawater to rinse the oocytes (see Note 1). Rinse the oocytes several times in filtered natural seawater and keep them at 17–20°C. 3.2. Oocyte Microinjection
The method used for microinjection is described in great detail elsewhere (17). Here we describe the basic method and modifications that are useful to recover the single cells for further analysis. 1. The procedure and methods used for starfish oocyte microinjection are identical to those published by Laurinda Jaffe and Mark Terasaki (17). 2. Oocytes are held between two clean glass coverslips separated by Scotch double-coated tape (see Note 2). The thickness of this tape is approximately 100 mm. Therefore, two layers of tape are needed to hold the 190-mm starfish oocytes. 3. During the microinjections, maintain a cool temperature at the stage where the oocytes are being injected, or in your microscope and injection room. We accomplish this using a portable airconditioner that can be directed toward the microscope stage. 4. Following microinjection, the oocytes are removed from the injection chamber. Because of their small size, care must be taken to follow each individual oocyte. Using the injection needle, gently roll the oocyte off of the coverslip ledge. 5. Using the coarse focus of the microscope, follow the oocyte as it falls to the bottom coverslip of the chamber. At this magnification, you will be able to identify the injected oocyte by the presence of the oil droplet that was introduced into the cytosol during the microinjection process. 6. Recover the oocyte using the mouth pipette and transfer it to an eppendorf tube (Fig. 5.1; Note 3). 7. Confirm the presence of the oocyte in the eppendorf tube under the dissecting microscope (Fig. 5.1; Note 3). 8. Freeze the oocyte at the appropriate time-point by immersion of the eppendorf tube into liquid nitrogen. 9. Store frozen oocytes at 80°C until needed for gels and western blot (see below).
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Fig. 5.1. Assaying pMAPK in single starfish oocytes. (A) A single immature starfish oocyte (arrow) in a small volume of seawater within a capillary tube following recovery after microinjection. The menisci at the left and right of the oocyte show the boundary of the seawater. This oocyte will then be transferred to a 1.7-ml eppendorf tube for sample preparation. (B) Single starfish egg (arrow) in the bottom of a 1.7-ml eppendorf tube with the associated volume of seawater. This will be snap frozen by dropping the tube into liquid nitrogen and stored at –80°C until assayed by western blotting. (C) Western blot of phosphoMAPK in single starfish oocytes. Immature starfish oocytes were injected with either deionized water or with 25-mM FPT III. After 5 min, the oocytes were treated with 1-mM 1-MA and incubated within the microinjection chamber. At 30 min after 1-MA treatment, the oocytes were removed from the microinjection chambers and placed in 1.7-ml eppendorf tubes. At 40 min after 1-MA treatment, the samples were snap-frozen in liquid nitrogen. Control oocytes were monitored for GVBD by viewing under a dissecting microscope (see (B) above). The proteins from individual oocytes were separated by SDS-PAGE and the presence or absence of phosphorylated MAPK was determined by western blotting. Lanes 1–6 show oocytes injected with FPT III and lanes 7–12 show control oocytes. FPT III inhibits GVBD and phosphorylation of MAPK in response to 1-MA.
3.3. Sample Preparation
1. Prepare fresh working solution of 2X Laemmli sample buffer (2X LSB) by adding 100 ml of -ME to 900 ml of LSB stock (18). Mix by vortexing. If needed, prepare 1X LSB by diluting 2X LSB with equal volume of deionized water. 2. Estimate total volume of oocyte and seawater in each eppendorf tube and add an equivalent volume of 2X LSB. If necessary, bring total volume of each tube to 20 ml using 1X LSB. 3. Heat samples for 3 min at 95°C (see Note 4). 4. After heating, centrifuge sample at 10,000 rpm for 30 s to collect evaporated liquid back to the bottom of the tube (if not
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using mineral oil) (see Note 4). Keep sample at room temperature until loading onto the gel; however, the gel should be loaded as quickly as possible after heating. 3.4. Gel Electrophoresis and Western Blotting
1. Pour a 10% polyacrylamide:bis-acrylamide (37.5:1 ratio) resolving gel with a 5% stacking gel. Load the entire 20-ml sample into one lane of the gel. Run samples at 100–110 volts using Tris-Glycine-SDS buffer in upper and lower chambers. In a mini-gel format, the gel will run for approximately 1.5 h. 2. Transfer proteins from gel to nitrocellulose using western transfer buffer (see above). Arrange the transfer to accumulate 1000 mA-h (see Note 5). After the transfer, the nitrocellulose sheet will contain the egg proteins preferentially on the side that was facing the gel. Make a note of this by writing on the nitrocellulose using a number 2 pencil (see Note 6). 3. Block non-specific binding using the appropriate blocking buffer (see Note 7). For the anti-pMAPK antibodies, use 5% non-fat dry milk in TST, pH 7.6. Incubate in blocking buffer for 1 h at room temperature with gentle agitation. We utilized a rotary-type shaker set at low speed. 4. Discard blocking buffer and add primary antibody solution to the nitrocellulose. The primary antibody is diluted to between 0.1 and 1 mg/ml in blotto. Incubate with gentle agitation for 2 h at room temperature or overnight at 4°C. 5. At the end of the incubation period, discard primary antibody solution and add 50 ml of TST to the blot. Incubate for 10 min with gentle agitation at room temperature. Discard wash solution at each change. Repeat for three washes. 6. Dilute secondary antibody (GAM-HRP or GAR-HRP, as needed) to 0.05–0.5 mg/ml in blotto. Discard final TST wash and add antibody to blot in same container. Incubate blot with secondary antibody for 1–2 h at room temperature. 7. Discard secondary antibody solution. Wash blot four times with TST as indicated above (step 5). 8. Prepare ECL reagent according to manufacturer’s instructions. Briefly, mix equal volumes of luminol solution to hydrogen peroxide. For each blot, prepare at least 1.5 ml of total ECL solution. 9. Remove blot from final TST wash and remove excess fluid by touching a corner of the blot to a piece of kimwipe. Place each blot, protein-side up, onto a piece of plastic wrap. 10. Overlay the blot with the ECL reagent prepared in step 8 above. Gently lift the edges of the plastic wrap to ensure even coverage of the ECL reagent over the blots. Incubate at room temperature for 5 min.
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11. At the end of the 5-min incubation, gently remove the excess ECL reagent by touching a corner of the blot to a new kimwipe. Be careful not to remove too much – the surface of the blot should remain moist. 12. Lay the blot, this time protein-side down, onto a new piece of plastic wrap. Carefully wrap the blot with the plastic wrap so that there is only a single layer of plastic over the front (protein side) of the blot and the plastic is neatly folded over the back of the blot. 13. Tape the wrapped blot onto a support so that it does not move during exposure. 14. Beginning 10 min after initial incubation with the ECL reagent, expose film to blot for 10 min. Overlay blot with second piece of film while developing first film (step 15 below). 15. Develop and fix film: 2 min in Kodak GBX Developer; rinse 30 s in room-temperature water; 2 min in Kodak rapid fixer; rinse in room-temperature water and dry at room temperature. 16. Adjust the exposure time for film number 2 (step 14 above) in response to the signal obtained for film number 1.
4. Notes 1. The CFSW rinses are useful in removing the follicle cells around each oocyte; however, the oocytes cannot be left indefinitely in the CFSW or they will begin to die. As soon as you see the follicle cells begin to fall off of the oocytes, you should allow the oocytes to settle to the bottom of the beaker and rinse them into filtered natural seawater. Oocytes left too long in CFSW take on a characteristic appearance of a darkening and coarse cytoplasm. 2. For a thorough discussion of the use of double-coated tape, please see the article by Jaffe et al. (17). This double-coated tape can be purchased at any office supply store. 3. Be conscious of how much seawater is transferred with each oocyte into the eppendorf tube. With practice, you will be able to keep this to a minimum. 4. Heating at 95°C results in evaporation of a significant portion of the 20-ml volume within the eppendorf tube. We dealt with this by overlaying the sample with mineral oil in some experiments; in other experiments, we allowed the evaporation to
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occur and then recovered the liquid by brief centrifugation (as noted above). Inclusion of mineral oil successfully prevented evaporation in these samples. However, no difference in experimental western blotting results was observed whether or not mineral oil was used. 5. The term ‘‘mA-h’’ is used to standardize the current to which the gels are exposed during the western transfer. This allows you to adjust the length of your transfer and get similar results. For example, 1000 mA-h can be obtained by transferring for 10 h at 100 mA or for 20 h at 50 mA. In both situations, you would expect to get similar results from the transfer process. 6. You can write directly on the nitrocellulose blot immediately following the electrotransfer. Place the blot, face-up, onto a clean lab bench or any clean hard surface. Using a fairly blunt number 2 (medium-soft) graphite pencil, mark the sample, date, and your initials directly onto the nitrocellulose. It is not recommended to use ink because the ink may be lost if you re-probe the blot. 7. Several brands of non-fat milk have been used for these experiments and no difference in results has been observed. We are currently using Carnation non-fat instant milk and Publix Supermarket brand non-fat instant milk in our experiments. It is important to thoroughly dissolve the milk in the TST; therefore, after adding the powdered milk to the TST it should be stirred for at least 30 min at room temperature. The Blotto can be stored at 4°C and used for approximately 24 h.
References 1. Haccard, O., and Jessus, C. (2006) Oocyte maturation, Mos, and cyclins. Cell Cycle 5(11), 1152–1159. 2. Liang, C-G., Su, Y-Q., Fan, H-Y., Schatten, H., and Sun, Q-Y. (2007) Mechanisms regulating oocyte meiotic resumption: roles of mitogen-activated protein kinase. Mol. Endocrinology 21, 2037–2055. 3. Avruch, J. (2007) MAP kinase pathways: the first twenty years. Biochim. Biophys. Acta 1773, 1150–1160. 4. Shaul, Y.D., and Seger, R. (2007). The MEK/ERK cascade: from signaling specificity to diverse functions. Biochim. Biophys. Acta 1773, 1213–1226. 5. Caffrey, D.R., O’Neill, L.A.J., Shields, D.C. (1999) The evolution of the MAP kinase pathways: coduplication of interacting proteins
6.
7.
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leads to new signaling cascades. J. Mol. Evol. 49, 567–582. Posada, J., and Cooper, J.A. (1992) Requirements for phosphorylation of MAP kinase during meiosis in Xenopus oocytes. Science 255, 212–215. Kishimoto, T. (2004) More than G1 or G2 arrest: useful starfish oocyte system for investigating skillful MAP kinase. Biol. Cell 96, 241–244. Philipova, R., and Whitaker, M. (1998) MAP kinase activity increases during mitosis in early sea urchin embryos. J. Cell Sci. 111, 2497–2505. Kumano, M., Carroll, D.J., Denu, J.M., and Foltz, K.R. (2001) Calcium-mediated inactivation of the MAP kinase pathway in sea urchin eggs at fertilization. Dev. Biol. 236, 244–257.
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10. Smythe, T.L., Stricker, S.A. (2005) Germinal vesicle breakdown is not fully dependent on MAPK activation in maturing oocytes of marine nemertean worms. Mol. Reprod. Dev. 70, 91–102. 11. Miller, M.A., Nguyen, V.Q., Lee, M.H., Kosinski, M., Schedl, T., Caprioli, R.M., and Greenstein, D. (2001) A sperm cytoskeletal protein that signals oocyte meiotic maturation and ovulation. Science 291, 2144–2147. 12. Eckberg, W.R. (1997) MAP and cdc2 kinase activities at germinal vesicle breakdown in Chaetopterus. Dev. Biol. 191, 182–190. 13. Tachibana, K., Machida, T., Nomura, Y., and T. Kishimoto (1997) MAP kinase links the fertilization signal transduction pathway to the G1/S-phase transition in starfish eggs. EMBO J. 16, 4333–4339. 14. Posada, J., Yew, N., Ahn, N.G., Vande Woude, G.F., and Cooper, J.A. (1993) Mos stimulates MAP kinase in Xenopus oocytes
15.
16.
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and activates a MAP kinase kinase in vitro. Mol. Biol. Cell. 13, 2546–2553. Matten, W.T., Copeland, T.D., Ahn, N.G., and Vande Woulde, G.F. (1996) Positive feedback between MAP kinase and Mos during Xenopus oocyte maturation. Dev. Biol. 179, 485–492. Lutz, L.B., Cole, L.M., Gupta, M.K., Kwist, K.W., Auchus, R.J. (2001) Evidence that androgens are the primary steroids produced by Xenopus laevis ovaries and may signal through the classical androgen receptor to promote oocyte maturation. Proc. Natl. Acad. Sci. USA 98, 13728–13733. Jaffe, L.A. and Terasaki, M. (2004). Quantitative microinjection of oocytes, eggs, and embryos. Meth. Cell Biol. 74, 219–242. Laemmli, U.K. (1970). Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227, 680–685.
Chapter 6 Analysis of Signaling Pathways in Zebrafish Development by Microinjection William H. Kinsey Abstract The zebrafish oocyte differs substantially from the zygote and cleavage-stage embryo with regard to the ease with which it can be microinjected with proteins or reagents that modify subsequent development. The objective of this chapter is to describe methods developed in this and other laboratories for microinjection and calcium imaging in the unfertilized zebrafish egg. Methods of immobilizing the oocyte include a holding chamber and a holding pipette. The holding chamber allows imaging of three or four oocytes simultaneously, while the holding pipette facilitates imaging of localized regions in the oocyte. Injection of calcium green dextran via holding chambers allowed detection of global changes in Ca2+ release following fertilization and development through early blastula stages. Injection and imaging with the holding pipette method allowed discrimination of calcium changes in the egg cortex from that in the central regions of the cell. The results demonstrate the highly localized nature of calcium signaling in the zebrafish zygote and the implications of this signaling for embryonic development. Key words: Zebrafish, egg, oocyte, zygote, fertilization, Fyn, Yes, Src.
1. Introduction The zebrafish has many advantages for the study of fertilization and early embryonic development due largely to the effectiveness of genetic and mutagenesis approaches in this model system. The optical clarity of the zebrafish egg and early embryo facilitates observation of dynamic signaling events by fluorescence microscopy and a considerable knowledge base has been developed on calcium signaling events in this species. The resting free calcium level in the zebrafish oocyte has been estimated at 60 nM (1) and the dynamics of calcium signaling following fertilization in this species have been studied with fluorescent reporters such as David J. Carroll (ed.), Microinjection: Methods and Applications, Vol. 518 Ó 2009 Humana Press, a part of Springer ScienceþBusiness Media, LLC DOI 10.1007/978-1-59745-202-1_6
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aequorin (2), calcium green (3), as well as ratiometric dyes such as Fura-2 (4). The benefits and limitations of these different reagents have been reviewed recently (5). The calcium signaling events triggered by fertilization of the zebrafish oocyte include an initial activation wave that traverses the egg at a velocity of around 9 mm/s in the egg cortex and 69 mm/s in the central cytoplasm (1) followed by more localized events associated with the streaming of cytoplasm toward the animal pole during blastodisc formation. At the first cell division, repeated calcium transients are associated with the cleavage furrow and are involved in furrow deepening (6). The cytokinesisassociated transients are repeated with subsequent cell cycles and are easily observed since these cell cycles remain synchronized for the first several cycles (1). While measurement of the total calcium-induced fluorescence in the zygote or embryo provides a certain amount of information, we have now used more detailed morphological analysis to quantitate calcium signaling in different subcellular regions of the zygote. The results demonstrate how localized changes in calcium signaling characterize the egg activation process. These studies have required further refinement of microinjection methods to overcome the physical difficulties inherent in the unfertilized zebrafish egg.
2. Materials 2.1. Buffers
1. Hank’s BSA: 137-mM NaCl, 5.4-mM KCl, 0.25-mM Na2HPO4, 1.37-mM CaCl2, 1.0-mM MgSO4, 4.2-mM NaHCO3, pH 7.2, 5 mg/ml BSA. 2. Injection buffer: KCl, 0.15 M; NaCl, 3 mM; KH2PO4, 10 mM (pH 7.2); glutathione, 1 mM; sucrose 80 mM; and 10-kDa calcium green dextran (50 mM) (Molecular Probes, Eugene, OR). 3. Sperm extender buffer: 10-mM HEPES; 80-mM KCl, 45-mM NaCl, 45-mM NaOAc, 0.4-mM CaCl2, 0.2-mM MgCl, pH 7.2.
2.2. Microinjection Chambers
1. The holding insert based on that originally described (2) was used to immobilize unfertilized zebrafish oocytes for microinjection. 2. Cut insert (1.5–2.0 cm) from a sheet of opaque plastic approximately 0.75-mm thick. 3. Drill holes as a row along one edge of the plastic using a handheld jewelers drill and a pin series bit #65 from Huot Corp. St. Paul MN, USA obtained at a local specialty tool retailer.
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Fig. 6.1. Design of an oocyte holding chamber. The row of holes approximately 800 mm in diameter were drilled along one edge of the plastic and openings were cut with a scalpel to admit the injection pipette (A). The cover was cut from a plastic coverslip as were guides to hold the cover on the insert but still allow it to move (B). Panel (C) shows the cover slid over the oocyte chambers to prevent their escape upon addition of aquarium water to initiate fertilization.
4. Cut slots to open these holes to the edge of the plastic sheet using a scalpel. 5. Assemble a sliding cover from sections cut from a plastic coverslip (Fig. 6.1). 6. Trim surfaces free of plastic fragments and attach the insert to the bottom of a plastic culture dish using two-part epoxy glue. 2.3. Microinjection Pipettes
1. Pull injection pipettes using borosilicate glass capillaries 1 350 mm of thick wall construction (World Precision Instruments, Sarasota, FL, USA). 2. Pipettes are beveled with a rotating disc pipette beveler (World Precision Instruments, Sarasota, FL, USA) at an angle of 30° to produce a tip diameter of approximately 2.5–3.5 mm.
2.4. Holder Pipettes
1-mm o.d. capillaries were flame-polished to reduce the opening to approximately 25% of the egg diameter, then bend over a flame or microforge to allow the tip to lay parallel to the bottom of the dish used for microinjection. The pipette was then filled with mineral oil and Hank’s BSA was drawn into the tip before picking up an egg (Fig. 6.2).
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Fig. 6.2. Holding pipette used to immobilize a zebrafish oocyte. A typical holding pipette produced by flame-polishing of a glass capillary is used to pick up and immobilize an unfertilized oocyte. Magnification is indicated by the bar which represents 100 mm.
2.5. Microinjector
We have used both a Picospritzer II pressure injector (United Valve Corp. Fairfield, NJ, USA) and a CellTram Vario syringe injector (Eppendorf Corp. Hamburg, Germany).
2.6. Confocal Microscopy
The eggs were imaged by confocal fluorescence microscopy on an inverted Nikon TE2000U microscope using a 4x or 20x super fluor objective. Long-working distance objectives were needed to focus through the plastic dish as well as the 700-mm egg. Illumination was provided with a 488-nm Spectra Physics (Mountainview, CA) laser with pinhole settings set to obtain a theoretical 24-mm optical section through the equator of the embryo. Emitted fluorescence was recorded at 15 s intervals and separate images were collected using a transmission detector to obtain a ‘‘brightfield’’ image and a 515-nm/30-nm bandpass filter to obtain calcium green fluorescence.
3. Methods 3.1. Zebrafish Culture Conditions and Egg Collection
1. Wild-type zebrafish between 3 and 6 months of age were maintained at 28°C under wide-spectrum fluorescent lighting
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(Coralife Trichromatic, Energy Savers Unlimited, and Carson, CA) with a 13 h-on/11-h off light/dark cycle. 2. Diet consists of Tetramarine flake food (Tetra Sales, Blacksburg, VA) supplemented daily with freshly hatched, live brine shrimp (Aquatic Lifeline Inc., Salt Lake City, UT) (see Note 1). 3. For sperm and egg collection, fish were anesthetized with 0.02% tricane (Sigma-Aldrich, St. Louis, MO) and squeezed between gloved fingers to express eggs or sperm. 4. Sperm were suspended in sperm extension buffer (2), which maintains them in an immotile state, and stored on ice for up to 2 h. 5. Eggs were stored in Hank’s BSA at 25°C and used within 30 min of collection (see Note 2). 3.2. In Vitro Fertilization, Microinjection Chamber Method
1. The microinjection chamber holes were filled with Hank’s BSA and a droplet (200 ml) of Hank’s BSA was formed across the edge containing the pipette slots. 2. Freshly obtained unfertilized eggs were transferred to the microinjection chamber with a yellow pipette tip cut to accommodate the large-diameter egg. Every effort was made to position the micropyle at 90° to the pipette slot so that it would not be damaged by the injection pipette. 3. Then the clear camber cover was slid over the egg to prevent its escape (Fig. 6.1C). 4. Next the sperm suspension (2 ml of 2 mg/ml sperm protein containing approximately 1.23 106 sperm) was added to the 200-ml Hank’s BSA. 5. The microscope field was then positioned to include three eggs (4x objective) or a single egg (20x objective) and three or four images were recorded to obtain pre-fertilization calcium levels. 6. Fertilization was triggered by adding 2 ml of aquarium water to activate sperm motility and allow fertilization to proceed.
3.3. In Vitro Fertilization, Holder Pipette Method
1. Unfertilized eggs were placed in a droplet (200 ml) of Hank’s BSA in the center of a 35- or 50-mm culture dish in the view field of an inverted microscope. The holder pipette was then used to rotate one oocyte so that the micropyle could be identified. The oocyte was then held by applying suction to the holder pipette with a screw-driven syringe pump (Fig. 6.2). 2. Once immobilized, the oocyte was injected with calcium green dextran or other reagents and the injection pipette withdrawn.
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3. After a recovery period of approximately 5 min, sperm were added to the droplet as above and image acquisition was begun. 4. Fertilization was induced by adding aquarium water (3 ml) to the dish which reduced the ionic strength and allowed sperm to become motile (see Note 3). 3.4. Image Acquisition and Analysis
1. An example of calcium release during activation of the zebrafish egg in a holding chamber is shown in Fig. 6.3. Within 60 s of fertilization, calcium release is evident in the cortical cytoplasm, but little change has occurred in the central cytoplasm. 2. Calcium green fluorescence (see Note 4) intensity can be measured simultaneously in several zygotes (see Note 5). 3. Once data collections were complete, calcium green fluorescence of an optical section through the entire oocyte was quantitated by Metamorph software and is presented graphically in Fig. 6.4 (upper panel). These measurements made from confocal image sections revealed that fertilization is followed by an initial low-amplitude calcium transient beginning as early as 15 s after sperm addition (black arrows) followed by two high-amplitude transients, one reaching a maximum at 2–3 mpf (grey arrows) and the second reaching a maximum at 8 mpf. These three initial transients are followed by later calcium signaling events associated with blastodisc expansion and cytokinesis as reported elsewhere (6,7).
Fig. 6.3. Fertilization-induced changes in calcium green fluorescence. Zebrafish oocytes were injected with calcium green dextran and an inactive protein (glutathione-S-transferase) then imaged by confocal fluorescence microscopy before fertilization (left), or at 60 s post-insemination (right). The increase in calcium green is most apparent in the egg cortex at 60 s post-insemination. Magnification is indicated by the bar which represents 100 mm.
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Fig. 6.4. Quantitation of fertilization-induced changes in calcium green fluorescence. Oocytes were injected with calcium green dextran and GST, then images were recorded after a 10-min recovery period. Fertilization was initiated by addition of a mixture of sperm and water at time 0 and images were recorded every 15 s. Fluorescence was quantitated by pixel intensity quantitation using Metamorph 6.1 software. The top panel represents global fluorescence measured from a cross-section of the entire zygote. The middle panel represent fluorescence measured within a circular region in the center of the zygote (central cytoplasm) comprising approximately 25% of the total cross-sectional area. The bottom panel represents fluorescence measured within an arc traced over the cortex (cortical cytoplasm) as near as possible to the micropyle. This arc comprised approximately 20% of the perimeter of the zygote.
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4. To more accurately study the localized calcium changes in the cortex and central cytoplasm, these regions were quantitated separately by measuring pixel intensity in different regions of the image. The cortical region was drawn as an arc to include approximately 20% of the egg cortex centered over the initial site of calcium release. The central cytoplasm region was drawn as a circle encompassing approximately 25% of the cross-sectional area of the oocyte. Both regions had to be relocated during the time series as the zygote moved during chorion elevation, and during cortical contractions; however, this analysis allowed detection of small, localized changes that were poorly represented in measurements from the entire oocyte. 5. Quantitation of the cortical and central cytoplasmic fluorescence demonstrated the difference between calcium signaling in these two compartments of the egg. For example, the initial low-amplitude spike at 15–30 s post-insemination was detected only in the cortical region (Fig. 6.4, bottom panel, black arrow) which is reminiscent of the cortical ‘‘hot spots’’ observed in Xenopus oocyte (8,9). The first high-amplitude transient (grey arrow), which reached a maximum between 2 min and 3 min post-insemination, was also more prominent in the cortical region than in the central cytoplasm. Later cortical transients did not occur in a consistent pattern, but did indicate that the cortical cytoplasm remained highly active. In contrast, the central cytoplasm exhibited only moderate changes until 5–6 min post-insemination when the second high-amplitude transient began, primarily involving the central cytoplasm rather than the cortical region (Fig. 6.4, middle panel). 6. Analysis of more localized changes in calcium signaling required that the egg be totally immobilized in a holding pipette to prevent movements from taking a given region out of the focal plane. This method allows exact positioning of the oocyte so that localized features such as the micropyle can be observed. At fertilization in fish, sperm penetrate the chorion through an opening (the micropyle) so the investigator can visualize the actual point of sperm–egg contact. As seen in Fig. 6.5 the cortical region under the micropyle responds to fertilization with increased cytoplasmic calcium within 30 s of sperm activation (arrows). 7. In summary, the unfertilized zebrafish oocyte can be effectively microinjected if methods are employed to immobilize the egg and allow the injection pipette to penetrate the very tough chorion. In addition, the method chosen to immobilize the egg can allow precise orientation so that specific subcellular regions can be observed. These techniques have shown the extent to which calcium signaling is compartmentalized in the zygote.
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Fig. 6.5. Calcium green fluorescence in the region of the micropyle. An unfertilized oocyte was picked up with a holding pipette and rotated until the micropyle was visible. The egg was then injected with calcium green dextran and fertilized as above. The images shown here were obtained at 50 s post-insemination and demonstrate the localized calcium green fluorescence in the region of the micropyle (arrows). Magnification is indicated by the bar which represents 100 mm.
4. Notes 1. Fish Maintenance: Fish are fed approximately 2 g of flake food plus brine shrimp hatched from 0.5 g of dried eggs each day. This high level of feeding results in improved egg production but also causes rapid acidification of the aquarium system. It is important to adjust the pH daily with Na2CO3 and wash the filter media each week. 2. Handling gametes: Unfertilized eggs are susceptible to aging and to rough handling and can undergo parthenogenic activation if they are exposed to too much agitation. Aging causes the chorion to harden and become even more resistant to injection pipettes. Sperm can be activated by exposure to aquarium water, so we rinse the anesthetized male fish with a few drops of sperm extender buffer before collecting sperm. 3. Full activation of sperm requires that the Hank’s BSA be diluted at least 10–15-fold by aquarium water. Therefore, a 200-ml droplet of Hank’s BSA would be diluted with 3 ml of aquarium water to initiate fertilization. 4. The calcium reporter calcium green dextran reports only the relative calcium changes and does not allow calculation of actual free calcium levels. It does have the major advantage that it can be excited with a 488-nm laser which causes much less damage to the cell than a shorter-wavelength ultraviolet source such as used for the fura series of ratiometric dyes. 5. The spacing of the holes in the egg holding chamber can be spaced so that three or four oocytes can be imaged simultaneously with a 4x objective.
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Acknowledgments Supported by NICHD-HD14846.
References 1. Creton, R., Speksnijder, J., Jaffe, L. (1998) Patterns of free calcium in zebrafish embryos. J. Cell Science 111, 1613–1622. 2. Lee, K., Webb, S., Miller, A. (1999) A wave of free cytosolic calcium traverses zebrafish eggs on activation. Dev. Biol. 214, 168–180. 3. Reinhard, E., Yokoe, H., Niebling, K., Allbritton, N., Kuhn, M., Meyer, T. (1995) Localized calcium signals in early zebrafish development. Dev. Biol. 170, 50–61. 4. Ahumada, A., Slusarski, D. C., Liu, X., Moon, R. T., Malbon, C. C.,Wang, H. Y. (2002) Signaling of rat Frizzled-2 through phosphodiesterase and cyclic GMP. Science 298, 2006–2010. 5. Slusarski, D., Corces, V. (2000) Calcium imaging in cell–cell signaling. Methods Mol. Biol. 135, 253–261.
6. Lee, K., Webb, S., Miller, A., (2003) Ca2+ released via IP3 receptors is required for furrow deepening during cytokinesis in zebrafish embryos. Dev. Biol. 47, 411–421. 7. Chang, D. C., Meng, L. (1995) A localized elevation of cytosolic free calcium is associated with cytokinesis in the zebrafish embryo. J. Cell. Biol. 131, 1539–1545. 8. Nuccitelli, R., Yim, D., Smart, T. (1993) The sperm-induced Ca2+ wave following fertilization of the Xenopus egg requires the production of Ins(1, 4, 5)P3. Dev. Biol. 158, 200–212. 9. Glahn, D., Mark, S., Behr, R., Nuccitelli, R. (1999) Tyrosine kinase inhibitors block sperm-induced egg activation in Xenopus laevis. Dev. Biol. 205, 171–180.
Chapter 7 Protein Inhibition by Microinjection and RNA-Mediated Interference in Tissue Culture Cells: Complementary Approaches to Study Protein Function Jane R. Stout, Rania S. Rizk, and Claire E. Walczak Abstract A major goal in cell biology is to understand the molecular mechanisms of the biological process under study, which requires functional information about the roles of individual proteins in the cell. For many non-genetic model organisms researchers have relied on the use of inhibitory reagents, such as antibodies that can be microinjected into cells. More recently, the advent of RNA-mediated interference (RNAi) has allowed scientists to knockdown individual proteins and to examine the consequences of the knockdown. In this chapter we present a comparison between microinjection of inhibitory reagents and RNAi for the analysis of protein function in mammalian tissue culture cells, providing both a description of the techniques as well as a discussion of the benefits and drawbacks of each approach. In addition, we present a strategy to employ RNAi for organisms without a sequenced genome. While the focus of our research is on the organization of the mitotic spindle during cell division and thus the examples utilized are from that system, the approaches described here should be readily applicable to multiple experimental models. Key words: Microinjection, RNAi, PtK cells, mitosis, siRNA, antibody, inhibition, protein function.
1. Introduction Understanding the molecular mechanism of any biological process requires the complete description of the role of each protein involved in that process. To achieve this goal, it is necessary to have an experimental means to perturb protein function as well as an assay to determine the functional consequences of that perturbation. The choice of approach will be determined by the reagents available to the protein of interest, the equipment available, David J. Carroll (ed.), Microinjection: Methods and Applications, Vol. 518 Ó 2009 Humana Press, a part of Springer ScienceþBusiness Media, LLC DOI 10.1007/978-1-59745-202-1_7
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whether or not the gene sequence is available, as well as the timecourse of the experimental process being analyzed. Microinjection of inhibitory antibodies or dominant-negative reagents has long been a powerful means to inhibit protein function in many cell types (1). However, it has often been criticized because it is questionable whether true loss of protein function is achieved. It is possible that the antibody is exhibiting cross-recognition of other proteins within the cell or that the antibody is simply binding in situ to the protein of interest and causing nonspecific blocking of other protein interactions. Often these drawbacks can be overcome by examining the effects of multiple antibodies to the same protein or by complementing antibody injection studies with other inhibition methods. RNAi has become an extremely useful tool for looking at protein function in many cell types. RNAi has revolutionized how most scientists view protein function studies, and the importance of this discovery is best highlighted by the awarding of a 2006 Nobel Prize to Andrew Z. Fire and Craig C. Mello, the scientists who first described this process (2). To carry out RNAi in vertebrate cells, short dsRNAs are introduced into the cell by transfection (3). This dsRNA then pairs with the endogenous mRNA and induces its degradation by a series of enzymatic activities. Because RNAi knocks out the mRNA, new protein synthesis is inhibited, and the protein levels decrease over the timecourse of the normal turnover of the protein of interest. In contrast to microinjection, RNAi does not require a purified antibody or dominant-negative reagents, but it does require some information about the individual gene sequence. For organisms in which the genome is sequenced, finding siRNAs to knockout any gene of interest is as easy as searching the website of companies such as Dharmacon or Ambion for their collection of pre-designed RNAs. If a favorite gene is not included in the predesigned collection, then designing a siRNA only requires entering the accession number of a protein into programs such as Block-IT siRNA Designer (http://rnaidesigner.invitrogen.com/rnaiexpress/) or Dharmacon siDesign Center (http:// www.dharmacon.com/sidesign/default.aspx). In the case of organisms without sequenced genomes, it is still possible to use these siRNA design programs by entering a short amount of sequence obtained by RT-PCR or from a cDNA clone. Perturbation of protein function by either microinjection of inhibitory antibodies or RNAi should be considered complementary methods of inhibition. Both methodologies have their own strengths and weaknesses that influence their suitability to answer a particular scientific question. For example, microinjection of inhibitory antibodies is quick and will typically display immediate changes in cell behavior and morphology. This allows the experimenter to time the injection relative to the process being analyzed.
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In contrast, RNAi requires a period of incubation to allow time for the targeted protein to be degraded. With antibody injection the experimenter can inject higher concentrations of the antibody to achieve complete inhibition, whereas with RNAi, sufficient residual protein may remain to carry out all or part of its cellular function. In microinjection, only a small number of cells are often examined, but the exact cell that was injected is known and therefore can be examined phenotypically. In contrast, RNAi is useful to examine a large number of depleted cells. However, since knockdown can vary across a population of cells, it is often difficult to determine if a particular cell shows a phenotypic effect due to depletion unless appropriate antibodies are available. Because of the unique characteristics of each methodology, we use both techniques as complementary approaches to more fully understand the cellular processes we are studying.
2. Materials 2.1. Preparation of Poly-L-Lysine Coated Coverslips
1. Coverslips: We use 12 mm round No. 1 coverslips (Fisher; 12545-80) for fixed analysis by immunofluorescence and 22 22 mm square No. 1½ coverslips (VWR; 48366-227) for live imaging and microinjection. For live imaging, where the sample will be injected on one microscope and imaged on another, it is useful to use photo-etched coverslips (Electron Microscopy sciences; 72264-23), which have a marked grid for ease of relocating the injected/treated cell. 2. Poly-L-lysine: Add 50 mg of poly-L-lysine (Sigma; P1524) to a final volume of 50 ml using sterile ddH2O. Store at –20°C; can be used up to 4 times. 3. Hybridization oven with hybridization bottles, or alternatively a hot plate. 4. Platform rocker. 5. 200 ml 1-M HCl (see Note 1.) 6. 3MM Chromatography paper (Whatman; 3030917), 46 57 cm.
2.2. Cell Culture
1. Maintenance media: PtK2 cells are maintained in complete Dulbecco’s Modified Eagle’s Medium (D-MEM) (Invitrogen; 11965-092) supplemented with 10% fetal bovine serum (Invitrogen; 16140-089), 1% penicillin/streptomycin (Invitrogen; 15140-122), and 1% GlutaMAX (Invitrogen; 35050-061); PtK-T cells are maintained in complete F-12 HAM (Invitrogen;
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11765-054) medium prepared with the same supplements as listed above for D-MEM. Store at 4°C in the dark. 2. 0.25% Trypsin (Invitrogen; 15050-065). Store at 20°C. 3. 70% ethanol. 4. Sterile Pasteur pipettes, pipettes, pipette tips, and 1.5 ml centrifuge tubes. 5. Hemocytometer. 6. Trypan blue, 0.4% w/v in PBS (12 mM phosphate, 137 mM NaCl, and 3 mM KCl, pH 7.4). 7. Tissue culture cells: We use PtK2 cells, which are adherent male marsupial kidney epithelial cells that have a flat morphology and a small number (2n ¼ 12) of large chromosomes. In our studies of mitosis, PtK2 cells provide an ideal system for detailed insight on phenotypes of the spindle components. For live imaging of the microtubule structure using fluorescence microscopy following injection, we use a PtK2 cell line stably expressing GFP-tagged alpha tubulin (PtK-T) (4). 8. Tissue culture plates: For routine passaging, we grow cells in Falcon 100 mm plates (353003). For plating over 22 22 mm coverslips for live imaging or microinjection, we use Corning 60 mm plates (25010). To test antibodies or for phenotypic analysis by immunofluorescence, we use either Corning 35 mm plates (351008) or Corning six-well cell culture plates (3506) (see Note 2). 9. Poly-L-lysine-coated coverslips: For preparation see Section 3.1. 2.3. Basic Immunofluorescence
1. Fixative: The type of fixative depends on the antibody being used for immunofluorescence (see Note 3). 2. Quench: A trace amount of sodium borohydride in 10 ml of TBS (0.15 M NaCl, 0.01 M Tris-HCl pH 7.4). TBS is made as a 10X stock and autoclaved to sterilize. Quench is only necessary if glutaraldehyde is present in the fixative. 3. TBS-Tx (TBS þ 0.1% Triton X-100). 4. Staining dish: We use a 150 mm gridded Petri plate with parafilm covering the bottom chamber and moist paper towels placed around the inside edge to prevent evaporation of antibody dilutions. The lid should be covered in either black electrician tape or aluminum foil to prevent photodamage to the fluorescently labeled secondary antibodies. 5. Abdil-Tx (antibody dilution solution): 2% BSA, 0.1% sodium azide made in TBS-Tx. Filter-sterilize and store at 4°C. 6. Primary antibody diluted in Abdil-Tx (see Note 4). 7. Secondary antibody conjugated to a fluorescent tag diluted in Abdil-Tx.
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8. 2 mg/ml Hoechst in TBS-Tx. Store at 4°C in the dark. 9. Mounting media: 0.5% p-phenylenediamine; 20 mM TrisHCl; 90% glycerol. This is made up by adding 0.1 g pphenylenediamine (Sigma; P6001) to 266 ml of 1.5 M Tris-HCl, pH 8.8 and 18 ml glycerol in a 50 ml tube. The p-phenylenediamine should be brown and flaky. Invert vigorously to make homogenous. Insert two hypodermic needles through a rubber stopper. Place the stopper securely in the 50 ml tube and wrap parafilm around the junction to seal. To one hypodermic needle, attach a hose hooked to a nitrogen tank and flow gas extremely gently over the surface of the liquid (liquid should only dimple slightly). The other needle should be venting excess gas from the tube. The reaction should take several hours and can go overnight. Finished mounting media should be light amber in color with no flecks. Aliquot and keep frozen at 80°C. A small working aliquot can be kept at 20°C, but it should not be used if it turns brown in color. 10. Fingernail polish. 2.4. Microinjection
1. Antibody storage buffer: (10 mM HEPES, pH 7.2; 100 mM KCl): Make up in water and store at 4°C. 2. Glass injection needles: Pre-pulled needles (World Precision Instruments; TW100F-6) provide a reproducible source of needles without the need to invest in a needle puller if one is not readily available. Make sure to use needles with a filament, which allows efficient delivery of the injection sample to the tip of the needle by capillary action. 3. Pipette and pipette tips for loading sample into injection needles: We use an Eppendorf Series 2000 Reference AdjustableVolume pipette and microloader pipette tips, 0.5–20 ml range (Eppendorf North America; 930001007). 4. Inhibitory reagents for injection: Affinity-purified antibodies at a concentration of at least 1 mg/ml (and sometimes as high as 10–15 mg/ml). A control non-immune immunoglobulin (IgG) raised in the same animal as the experimental antibodies should also be used at an equal or higher concentration than the experimental antibodies. 5. Nikon IM300 microinjector with Nikon/Narishige microinjector controls and a Nikon TE-300 inverted microscope (Fig. 7.1A). 6. Two thermoprobes: These are digital thermometers with a remote probe (Acu-Rite; Model # 00890A1), which are used to monitor the temperature of both the microscope stage and the plate warmer.
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Fig. 7.1. The microinjection setup. (A) The microinjection setup includes a Nikon IM300 microinjector with Nikon/ Narishige micromanipulators mounted on a Nikon TE-300 inverted microscope with an attached CCD camera. A slide warmer on the right side is used for short-term incubation (<1 h). (B) A close-up view of the assembled rose chamber mounted on top of the microscope stage secured with tape. A thermoprobe is placed directly under the tape on top of the rose chamber to monitor its temperature during live imaging.
7. Rose chamber: This is a cell imaging chamber, designed for high resolution and long-term live imaging (Fig. 7.2). The chamber consists of three main layers: two metal planchets and one silicon spacer. The rose chamber is designed so that imaging can be carried out through the thin glass of the coverslip while providing enough space for holding up to 1 ml of media on top of the cells and a thin layer of mineral oil that covers the media surface preventing evaporation of media. The rose chamber can be replaced with glass bottom culture dishes (MatTek Corporation; P35GC-1.5-14-C). For a detailed discussion of different viewing chambers refer to (5). 8. Heating tray used as a slide or culture dish warmer.
Fig. 7.2. Assembly of rose chamber for long-term live imaging. (A) A disassembled chamber is shown to illustrate the order of placement of each component of the assembled chamber. (B) An assembled rose chamber.
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9. Air stream incubator (ASI) (Nevtek, Burnsville, VA): This is a device that maintains a flow of air at constant temperature, which is necessary to keep mammalian cultured cells at 37°C. The ASI can be replaced with any available temperature-controlled stage device. 10. Observation/imaging media: For phase contrast imaging of PtK2 cells, we use D-MEM with no phenol red, (Invitrogen; 11039-021), supplemented with 20 mM HEPES (titrated to pH 7.2 with potassium hydroxide) before use to maintain the appropriate pH of the media in the absence of CO2. When imaging fluorescent protein-expressing cells, it is also necessary to add an oxygen scavenging mix such as oxyrase to prevent photo-damage. Oxyrase (Oxyrase Inc.; EC-0050) is used at a final concentration of 0.3 units/ml. 2.5. Oligofectamine Transfection (RNAi)
1. Incomplete D-MEM: D-MEM with no supplements. 2. RNAi media: D-MEM supplemented only with 10% fetal bovine serum and 1% GlutaMAX but without antibiotics. 3. Oligofectamine (Invitrogen; 12252-011) (see Note 5). 4. siRNAs at 20 pmol/ml: siRNAs designed to the genes of interest as well as negative and positive control siRNAs. For a negative control, Dharmacon’s non-targeting siRNA #2 works well in PtK2 cells with no noticeable non-specific effects. For a positive transfection control for mitotic cells, Eg5 siRNA (50 CAAGGAUGAAGUCUAUCAAdTdT) (6) is a good choice because the loss-of-function phenotype (monopolar spindles) is easily identifiable, the phenotype manifests itself after a short incubation time, and because the kinesin-5 Eg5 is found across a wide phylogeny of organisms (7,8) (see Note 6).
3. Methods 3.1. Preparation of Coverslips
Tissue culture cells generally adhere to glass somewhat poorly. Treating glass coverslips with poly-L-lysine helps cells remain attached to the surface keeping them flatter, which improves the ability to follow the intracellular events more clearly. The coverslips are first extensively washed with acid, which etches the glass, and then coated with poly-L-lysine.
3.1.1. Acid Wash
1. Place 1–2 boxes of coverslips in 1-M HCl and heat to 60°C for 4–16 h. A hybridization oven with an internal rotisserie works well for this step, since it can easily be set to the desired temperature. The coverslips can be divided equally between two hybridization bottles containing 50–100 ml of acid. If a
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hybridization oven is not available, a hot plate and glass dish can be used (see Note 1). 2. Let cool to room temperature and carefully decant the acid into a separate container and dispose of appropriately. 3. Rinse coverslips 3X 10 min with dH2O then with 3X 10 min with ddH2O. If the acid wash is performed in a glass container on a hot plate, transfer the glass container to a rocker for the rinses and rock gently to allow the H2O to evenly reach the coverslips. Do not shake or rock too vigorously, or the coverslips will break. 3.1.2. Poly-L-lysine Coating
1. To coat coverslips, replace the ddH2O with 50 ml of 1 mg/ml poly-L-lysine while rocking or rotating gently for at least 30 min. 2. Decant poly-L-lysine and store at 20°C. This solution may be reused up to four times. 3. Rinse coverslips 5X 10 min with dH2O, then 5X 10 min with ddH2O to remove any free poly-L-lysine, which is toxic to the cells. 4. Dry coverslips by laying them out individually on Whatman paper. They must be separated or they will dry stuck together. 5. To sterilize coverslips before plating cells, cover with 100% ethanol, aspirate off, and let dry for a few minutes in the hood.
3.2. Maintenance of Cultured Cells
All of the subsequent protocols should be performed in a sterile tissue culture hood using sterile techniques and solutions to avoid contamination and its potential spread to other cultures (see Note 7.). For more in-depth information on media, cells, and maintenance techniques, please refer to (9,10). 1. Prewarm trypsin and maintenance media in a 37°C water bath. 2. Remove semiconfluent PtK2 plate(s) in log phase from the 37°C, 5% CO2 incubator. Aspirate off media from cells and rinse each plate 2X with 1.5 ml of trypsin. Incubate in 1 ml of trypsin for 5 min in the incubator until the cells detach from the plate. Add 9 ml of maintenance media to each trypsinized plate and combine multiple plates. Pipette cells up and down and across the plate to resuspend the cells. 3. For routine passaging, dilute trypsinized cell solution into prewarmed maintenance media. Return the newly passed cells to the 37°C incubator (see Note 8). 4. For plating for analysis of fixed cells by immunofluorescence, plate diluted cell solution over sterile poly-L-lysine coated coverslips (For preparation of coverslips, see Section 3.1.). The size of the plate will depend on the number of coverslips that need to be analyzed. (For plating volumes and the number of coverslips that will fit into different size dishes, see Note 9.)
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5. To plate cells for microinjection, plate as if for immunofluorescence, except use 60 mm dishes over sterile 22 22 mm coverslips. 6. For siRNA transfections, the number of wells to be plated for RNAi will depend on how the cells will be analyzed (see Note 10). The plating density of the cells also depends on a variety of factors, one of which is incubation time prior to cell analysis. Since this will vary for each protein being knocked down, this will have to be optimized for each oligonucleotide. For our experiments, PtK2 cells typically are plated at 15,000–20,000 cells/ml per 35 mm dish. A 100 mm plate of cells at log phase is sufficient to plate at least 24–35 mm wells. We also typically plate four wells per oligonucleotide: one for immunofluorescence and three for immunoblotting (see Note 11 for further considerations.). 7. To perform cell counts for plating for RNAi, place 90 ml of the trypsinized cell suspension in a 1.5 ml microcentrifuge tube. Add 10 ml of trypan blue, which stains dead cells blue, and let sit for a few minutes (no longer than 10 min). Pipette up and down to break up any cell clumps and to resuspend any settled cells and add 10 ml of cells to each side of a hemocytometer. Count the number of non-blue cells and calculate the cell density. 3.3. Basic Immunofluorescence
In the study of cellular processes, the analysis of fixed cells by fluorescence microscopy is usually the first step to assessing the initial phenotypic effects on the cell by either microinjection or RNAi. Specific steps for these procedures are outlined below for the study of PtK2 cells. For more indepth information, please refer to (11). 1. Pull plates with cells growing on 12 mm coverslips from the 37°C, 5% CO2 incubator. 2. Aspirate off media and rinse cells quickly with PBS. 3. Add fixative and incubate for an appropriate amount of time based on the fixative used. Alcohol-based fixatives, such as cold methanol, are incubated at RT for 5 min. Aldehyde fixatives are incubated at RT for 20 min. 4. If cells were fixed with glutaraldehyde replace fixative with 1 ml of Quench and incubate for 5 min. 5. Rinse cells 2X with TBS-Tx. 6. Move the coverslips to a parafilm-lined staining dish, keeping each coverslip wet with TBS-Tx. Wash 1X with a stream of TBS-Tx. Always finish by leaving a drop of buffer on the coverslip to keep the cells hydrated. 7. Aspirate off all TBS-Tx, and overlay each coverslip with 75 ml Abdil-Tx for minimum of 30 min at RT.
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8. Aspirate off Abdi-Tx and replace with 35 ml of primary antibody diluted in Abdil-Tx, and incubate for 30 min. Wash 3X with TBS-Tx. 9. Aspirate off last wash and incubate coverslip for 30 min in 35 ml of secondary antibody conjugated to a fluorescent tag. Wash 3X with TBS-Tx. 10. The cells can be co-stained with additional antibodies by repeating steps 8 and 9. 11. Stain the DNA by incubating coverslips for 5–15 min in 50 ml of 2 mg/ml Hoechst made in TBS-Tx. Wash coverslips 3X with TBS-Tx. 12. To mount, blot each coverslip briefly by touching its edge to a kimwipe, then place cell-side down on a 2 ml drop of mounting media on a slide. Aspirate the top of the coverslip, seal the circumference with fingernail polish, and let dry. 13. Coverslips must be washed gently with either water or lens cleaner before viewing on the fluorescence microscope. This is particularly important when using oil immersion objectives where any remaining salts on the coverslip will damage the lens. 14. Slides can be stored at 20°C in the dark for several weeks before the fluorescence fades. 3.4. Microinjection
3.4.1. Preparation of Injectate
Microinjection of either antibodies or dominant-negative reagents into vertebrate tissue culture cells is a powerful way to analyze protein function. The cells can be injected during interphase or during mitosis so it is possible to achieve temporal resolution of the experiment. The first part of this section describes the preparation of the injectate (Section 3.4.1.) and the microscope and rose chamber (Section 3.4.2.) followed by an outline of the microinjection steps (Section 3.4.3.). We then present the application of the microinjection for either a fixed time-point experiment (Section 3.4.4.) or for live imaging (Section 3.4.5.). 1. Antibodies are purified by affinity chromatography and eluted using Glycine-HCl, pH 2.0, followed immediately by neutralization in 2 M Tris-HCl, pH 8.5 (12). 2. The antibodies are then dialyzed into antibody storage buffer, quantified by A280, flash frozen in 10 to 30 ml aliquots, and stored at 80°C. 3. For injection, the antibodies and control IgG are thawed on ice and diluted to the desired concentration in antibody storage buffer (see Note 12). 4. Prepare antibody/injection sample by spinning at 14K rpm for 15 min and transfer the supernatant to a clean tube for injection. Place on ice for the duration of the experiment.
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1. Prewarm slide warmer and imaging media (and rose chamber parts, if needed) to 37°C. 2. Place a thermoprobe on the stage beside where either the rose chamber or culture will be mounted (Fig. 7.1B). 3. Turn on ASI and adjust the air flow at the stage. Allow the temperature to stabilize to the desired temperature (see Note 13), readjusting the ASI as necessary. 4. For imaging of live cells, cells grown on a 22 22 mm coverslip are transferred to a 35 mm dish containing 2 ml of prewarmed imaging media and placed on the 37°C warming tray. 5. Assemble the rose chamber (Fig. 7.2). This needs to be done quickly to prevent the cells from cooling too much. Blot to remove excess media, and quickly place the coverslip onto the bottom metal planchet of the chamber, positioning it in the center with the cells facing up. Quickly place the silicon spacer over the coverslip and then the top planchet on the silicon spacer. Keep both in place by pressing down firmly on the top planchet. Using a transfer pipette, add approximately 1 ml of prewarmed media into the chamber on top of the cells. While still pressing firmly, screw down the corners of the chamber. Wipe the bottom of the coverslip with a kimwipe, wet with dH2O, then wipe with a kimwipe wet with 70% EtOH. Using water first will prevent the ethanol from precipitating proteins and salts from the media. Cover the observation media with a layer of mineral oil to prevent evaporation, and place the assembled chamber onto the stage. Tape it down as shown in Fig. 7.1B to prevent it from moving during the subsequent steps. 6. If injecting cells for fixed-time-point analysis, cells grown on a 12 mm coverslip are placed in a 35 mm tissue culture dish with 2 ml of prewarmed observation media. Place the dish on the slide warmer to keep at 37°C until ready to inject. For injection, the dish is simply moved to the microscope stage ring designed for a 35 mm dish. Injection is carried out directly in the 35 mm dish.
3.4.3. Microinjection of Cells
1. Scan the surface of the coverslip to find the cell of interest. For interphase cells find a group of cells to inject (Fig. 7.3A). For mitotic cells find a cell at prophase, right before nuclear envelope breakdown has taken place (Fig. 7.3F). Center the cell in the cross-bars of the eyepiece. 2. Adjust the needle holder, which is attached to the stage to an approximately 45° angle and tighten it in place. 3. For a system with an automated injector, open the N2 tank valve and adjust the output pressure to 60–80 psi.
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Fig. 7.3. Microinjection of interphase and mitotic PtK2 cells are shown in sequence. (A) An interphase cell is shown. (B) The same interphase cell is shown with the microinjection tip just out of focus above the cell. (C) The microinjection tip is in focus above the cell prior to injection. (D) As the cell is injected, the injectate can be seen as a wave traveling through the cell. (E) The cell is shown post-injection. (F) A prophase cell is shown with intact nuclear envelope and condensed chromosomes prior to injection. (G) A prophase cell is shown with the microinjection tip just out of focus above the cell. (H) The microinjection tip is in focus above the cell prior to injection at a thick, organelle-rich area, away from the nucleus. (I) As the cell is injected, the injectate can be seen as a wave traveling through the cell. (J) An image of the cell is shown post-injection.
4. Using the eppendorf microloader tip and pipette, carefully backload the needle with 2 ml of the sample. Avoid touching the needle against anything for it is extremely fragile and will easily break. Attach the filled needle to the needle holder on the stage and secure. 5. Turn off the ASI before injection. Failure to do so will lead to warming the needle to above 37°C and killing the injected cell (see Note 14). 6. By eye, position the needle with the coarse X and Y adjustment knobs to place the needle tip at the center of the transmitted light on the surface of the media. Using the Zaxis coarse knob, carefully lower the tip of the needle so that it breaks the surface of the media, but do not lower it so far that it hits the bottom of the dish and breaks. 7. At this point the needle tip should be in the general vicinity of the cells, and its position is now carefully fine-tuned. To first detect the position of the needle, look through the eyepiece and move the needle using coarse knobs in the X- and Y-axes, to move laterally in relatively large increments. A shadow of the needle moving across the field of view should be detectable. Move the needle using the Y-axis knob until the shadow is at the center of the field of view and adjust with the X-axis knob so that the tip of the needle, which should appear as the narrowest part, is at the center of the field of view. While the
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tip of the needle is in sight, carefully lower the needle in the Zaxis so that the needle is just above the cell surface (Fig. 7.3B and G). 8. Adjust the balance pressure to approximately 1–3 psi, which will create a constant flow of sample from the tip. The sample should now be detected through the eyepiece as a stream of liquid flowing from the tip. 9. To inject in the cytoplasmic area of the cell, center the target cell and aim at a region that is relatively thick. It is usually easiest to inject cells near the periphery of the nucleus because the cell is thickest in this area (Fig. 7.3C and H). 10. Quickly move the tip down in the Z-plane using the fine adjustment of the joystick and then back up (a twisting movement of the joystick back and forth). As the cytoplasm is injected, a fast but gentle movement of sample is seen moving throughout the cell (Fig. 7.3D and I). 11. Before removing the dish/imaging chamber, move the tip up with the Z-plane coarse knob to avoid breaking the needle tip, which can be used for the next set of cells as long as it is intact and functional. 12. For trouble-shooting injections, see Note 15. 3.4.4. Fixed-TimePoint Assay
When first analyzing the function of a protein by microinjection of antibodies, it is easiest to assess the phenotype by injecting the antibody, waiting a set amount of time and then fixing the cells and processing them for immunofluorescence. If it is not known whether the affected process is during interphase or mitosis, it is simpler to start with interphase cells because they are easier to inject and there are more of them on a coverslip. When looking for a mitotic defect, it is easiest to inject only cells at prophase and then allow the cells to incubate at 37°C to progress through mitosis. For interphase cells, we usually start with a 2 h incubation in antibody and have extended that time period up to 24 h post-injection before analysis. For mitotic cells, we usually fix at 30 min post-injection for prometaphase/metaphase defects and 40 min post-injection for anaphase defects. The cellular morphology as well as the cell cycle stage is scored under a fluorescence microscope. 1. Place 35 mm dish with cells growing on 12 mm coverslips onto the stage ring plate designed for 35 mm dishes. 2. Gently inject the cells as instructed in Section 3.4.3. Inject each coverslip for a period of 5 min and return to the 37°C warming tray. Keeping the injection time period short is a way to maintain cell cycle synchrony in the experiment, at least for mitotic cells. 3. Allow the cells to incubate for the desired amount of time. For a short incubation (less than 1 h), it is fine to leave the cells on the warming tray. For longer incubations, return the cells to
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the CO2 incubator, which will better maintain temperature and humidity. 4. Directly following incubation, aspirate off the observation media and add 2 ml of fixative. Fix for the appropriate time period and follow by staining and mounting the coverslip on a microscope slide for observation (see Section 3.3 for basic immunological protocols). 3.4.5. Live Imaging
1. Place assembled rose chamber with live cells on the stage. 2. Select live imaging acquisition settings on the imaging software available with your scope (see Note 16). 3. Inject the cell as instructed in Section 3.4.3. As soon as the cell has been injected, quickly remove the needle and turn the ASI back on. 4. Turn off the microscope light and allow at least one minute for cell to recover, then view live on the screen to center the cell and refocus. 5. Begin time-lapse imaging, and record the time of injection and the time when imaging commenced. 6. Throughout imaging, make sure that the ASI is at 36–38°C. The rose chamber and coverslip with no change of media can be used for a maximum of 5 h. For more detailed discussion of live imaging refer to (5).
3.5. Oligofectamine Transfection (RNAi)
3.5.1. Designing siRNAs for Knockdown
Knocking down protein levels by RNAi is a fairly simple way to address protein function. However it relies on the availability of cDNA sequence to design the siRNA. We describe a fairly straightforward approach to obtain a sufficient amount of sequence for siRNA design to apply this technology to model systems without a sequenced genome. We then describe a basic protocol for transfecting adherent cells grown in culture. All steps should be performed in a sterile tissue culture hood using sterile techniques and solutions. The rat kangaroo genome has yet to be sequenced, therefore no databases exist to search for gene sequences of interest. Traditional methods of cloning cDNAs are laborious and time consuming; however, comparison of homologous sequences across different mammalian species often shows a high degree of DNA identity through portions of the coding region if not the whole sequence (6) that can be used to design primers for RT-PCR, which in turn are used to generate siRNAs. 1. Search databases for homologous sequences to the desired gene. We used homologous sequences from mammalian species since these theoretically should have the most identity to the rat kangaroo genome. 2. Generate sequence alignments using the mammalian homologues on a program such as Sequencher (Gene Codes
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Corporation), which allows hand manipulation to refine the alignments to gain the highest identity possible between the sequences used. 3. Design degenerate primers that (1) minimize the degree of degeneracy needed as much as possible, (2) produce a PCR product between 400 and 1000 bp long, and (3) that overlap, where possible, any published siRNAs of any of the homologous sequences. 4. Use total RNA isolated from PtK2 tissue culture cells as template in an RT-PCR reaction using the designed degenerate primers (see Note 17). 5. Purify the RT-PCR product, either the whole or a partial gene sequence. Sequence both strands of the PCR product and compare the results to the original alignments to verify that the product does indeed have a high degree of identity to the homologous sequences used in the alignment. 6. Use the confirmed sequence to design siRNAs for subsequent RNAi transfections using programs such as the Dharmacon siDesign Center program.
3.5.2. Oligofectamine Transfection
This is a modified version of the Invitrogen protocol ‘‘Transfecting siRNA into HeLa Cells Using Oligofectamine’’ (http://www.invitrogen.com/content/sfs/protocols/sirna_oftsf_proc.pdf) (3,13). 1. Prewarm all media to 37°C. 2. For each 35 mm well to be transfected, prepare the RNAi complexes (see Notes 18–20): a. Before use, gently mix Oligofectamine, then dilute 3 ml of transfection reagent into 12 ml of incomplete D-MEM in a 1.5 ml sterile microcentrifuge tube. Pipette gently up and down to mix and let incubate at room temperature for 5 min. b. Meanwhile, in a second sterile 1.5 ml microcentrifuge tube, add 200 pmol of siRNA to 175 ml of incomplete media. Pipette up and down gently to mix. Add the diluted siRNA to the Oligofectamine complexes after their 5 min incubation and mix gently. c. Let the siRNA:lipid complexes form at room temperature for 20 min. 3. While the complexes are incubating, rinse cells 2X with 1 ml of incomplete D-MEM media, and then cover with 1 ml of RNAi complete media. 4. Add 800 ml of D-MEM complete to the siRNA/lipid complexes. Remove media from wells and replace with the 1 ml of
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siRNA/lipid RNAi media. Incubate at 37°C, 5% CO2 incubator for 24 h. 5. Add 1 ml of D-MEM complete media back to each 35 mm well. Return to 37°C, 5% CO2 incubator. 6. Assay for gene activity at 24–72 h after transfection as is appropriate for the target gene (see Notes 21–22).
4. Notes 1. HCl is extremely corrosive, inhalation of vapor can cause serious injury, and liquid acids can cause severe damage to skin and eyes. Use appropriate precautions when making and handling the hot acid, and dispose of the acid properly. If a hot plate and glass container are used to acidwash the coverslips, the temperature and acid level will have to be monitored much more closely to make sure the acid does not overheat, and the glass container should be loosely covered to prevent excessive evaporation. The use of a hot plate requires that this step be carried out under a ventilated hood. 2. All tissue culture plates are not created equal. We have experienced problems with unhealthy cells when we have attempted to try plates from different manufacturers. When changing any reagents involved with tissue culture, it is always prudent to test the new batch on a subpopulation of cells before completely switching over. 3. The antibodies used will determine the specific fixative needed. To look at microtubule structure with DM1 antitubulin antibody (Sigma), we find that fixation with 4% formaldehyde; 0.1% glutaraldhyde in PHEM buffer (60 mM PIPES, 25 mM HEPES, 10 mM EGTA, and 4 mM MgSO4, pH 7) works well. However, many of our other polyclonal antibodies do not work well with glutaraldehyde fixation, and we typically use 2–4% formaldehyde in PHEM buffer without the glutaraldehyde. Cold (20°C) 100% methanol can also be used to fix cells in which microtubules are to be visualized, and this works for some of our other antibodies as well. It should be noted that methanol fixation can cause some distortion in the condensed chromatin of mitotic cells. Dispose of used fixative in accordance with local and state regulations. 4. Antibody dilutions can be stored at 4°C; however, the concentrations and the lengths of time these dilutions can be
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used will vary from antibody to antibody and will have to be empirically determined by each lab. 5. For siRNA transfection in PtK2 cells, several popular lipidbased reagents were tested, many of which caused vesicularization in cells, which can interfere with subsequent imaging by microscopy. We found that Oligofectamine (Invitrogen) produced the most consistent results with the least degree of cytotoxic effects compared to other transfection reagents tested. 6. siRNAs purchased from Dharmacon have been more successful for us than siRNAs from other companies both in their reliability and efficiency of knockdown. For a negative control, Dharmacon’s non-targeting siRNA #2 designed to Luciferase works well in PtK2 cells, whereas the GFP siRNA (GCAAGCUGACCCUGAAGUUCAU) (14) produced cytotoxic effects in our PtK2 cells. 7. It cannot be stressed enough the importance of an ever-present diligence to avoid contamination. Cultures should be routinely examined under the microscope to assess the health of the culture and inadvertent cross-contamination of other cell lines, or contamination by fungi or bacteria. Cells should also routinely be tested for contaminants such as mycoplasma, either by PCR or immunofluorescent assays. 8. Many cell lines are sensitive to cell density; plating at densities either too low or too high can have an adverse effect on the cells. 9. A 35 mm plate holds 2 ml of media and four 12 mm coverslips; a 60 mm plate holds 4 ml of media and 12 12 mm coverslips or two 22 22 mm coverslips and four 12 mm coverslips; and one 100 mm plate holds 10 ml of media and up to 30 12 mm coverslips. 10. The number of wells to be plated for RNAi will depend on how cells will be analyzed. We typically plate the cells in 35 mm dishes or six-well plates (each well is equivalent to a 35 mm dish). For live imaging, cells are plated on 22 22 mm coverslips in 60 mm dishes. For immunoblots, cells from three wells for each experimental condition are trypsinized, washed, and counted. The number of wells harvested for immunoblots can also be dependent on the antibody used. 11. The density at which cells will be plated for RNAi experiments depends on many factors, such as the growth rate of the cells, the media in which the cells are grown, and the number of days required to achieve knockdown of the protein as determined by either immunofluorescence, immunoblot, or RT-PCR. For Eg5 RNAi in our PtK2 cells, the effect is
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seen in as little as 24 h after transfection though we fix cells for immunofluorescence by 48 h. For kinesin-13 MCAK RNAi, cells are processed after 72 h for efficient knockdown (6). Other factors include cell density at processing; overly confluent cells at processing will retard the number of mitotic cells and cause difficulty in imaging. However, the transfection efficiency is at times better if the cells are more confluent versus less, so sometimes a balance needs to be maintained. The optimal plating density will need to be empirically determined. 12. The concentration of antibody used must be empirically determined. We often start with a needle concentration of 1–2 mg/ml and have rarely needed to go above 5 mg/ml. There are, however, reports in the literature in which people have used much higher concentrations. The buffer in which the antibody is stored must be at physiological pH, cannot have too high a salt concentration and must not contain sodium azide or other preservatives, which will kill cells. 13. While it would be ideal to have a thermoprobe directly monitoring the temperature of the media while imaging, this is not possible due to space restraints and perturbation of the cells. Instead, we attach the thermoprobe to the top-side of the rose chamber. Because the temperature at the top of the chamber will be different than inside the media-filled chamber, it is necessary to predetermine this difference before injections. To do this, place the assembled rose chamber on the stage (Fig. 7.1B). Attach one probe directly on top of the glass coverslip, using tape. Add 1 ml of media and a thin layer of mineral oil on top of the media. Place a second thermoprobe at the top-side of the rose chamber (Fig. 7.1B). Turn the ASI on and allow the temperature reported by the probe inside the rose chamber to reach the desired temperature and remain constant. This probe reflects the closest estimation to the temperature of the cells during imaging. Now record the temperature of the probe placed at the side of the chamber. This is the temperature that will be used to maintain cells at the actual desired temperature. In our setup, we have determined that the temperature of the coverslip within the rose chamber is typically 1°C lower than that of the probe on the top of the rose chamber. 14. As the needle is lowered into the imaging media, it is in a direct flow of the ASI and will heat up to temperatures several degrees warmer than the media. This increased temperature of the glass needle will kill the injected cell, making it imperative that the ASI be turned off right before injections.
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15. Troubleshooting injections: For a clogged needle, press the clear button, or gently scrape the tip of the needle along a cell-free area of the coverslip to unclog it. Be careful because this latter method can also damage the needle tip. A clogged needle can also be due to a high concentration of the injectate or aggregates in the injectate. To alleviate these problems, try diluting the sample or centrifuging at a higher speed before loading the needle. 16. For live imaging of mitotic PtK2 cells under phase contrast microscope we use 100 ms exposures at 30 s intervals for 120 min, which allows us to follow the events of mitosis without damaging the cell. This can be optimized depending on the nature of the experiment. 17. Using degenerate primers is not typically recommended in RT-PCR. However we have had some success if we can limit the degree of degeneracy of the primers. Additionally, in some instances our RT-PCR gave multiple products, which required isolating the different bands and sequencing each band. By comparing the sequences back to the original alignments, we identified the band with the most identity to the homologous gene sequences as the intended product. Additionally, if we could obtain at least a partial sequence of the RT-PCR product, we could further amplify the PCR product by doing nested PCR on the RT-PCR product. 18. If transfecting cells in multiple wells with the same siRNA, cocktail mixes can be used. In this case, multiply each reagent added by a fraction over the number of wells to be transfected. For example, if transfecting three wells with luciferase siRNA, add 3.2-fold more of each reagent to the appropriate tubes: 9.6 ml of Oligofectamine to 38.4 ml of incomplete media and 32 ml of siRNA to 560 ml of incomplete media. The siRNA/lipid complexes can then be added to 2560 ml of RNAi media in a 5 ml sterile tube instead of the 800 ml of RNAi media being added to the complexes. Aliquots of this cocktail are then used to replace the rinse media. 19. The ratio of lipid to siRNA may need to be optimized for each siRNA used to achieve the most efficient knockdown. Using excess lipid or siRNA will increase non-specific cytotoxic effects of the RNAi transfection. Some gene targets may require multiple siRNAs for efficient knockdown or may require that the concentration of oligonucleotide be adjusted to increase the transfection efficiency. Dharmacon has a useful complementary guide book, RNA Interference – Technical Reference & Application Guide (www.dharmacon.com) that has many useful suggestions on how to troubleshoot RNAi.
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20. The length of time a cell line has been subcultured affects the efficiency of the knockdown. This seems to be particularly true for PtK2 cells, and only those cells subcultured for 8 weeks or less should be used for RNAi. 21. The percentage of knockdown that needs to be attained to see a phenotypic effect can vary greatly depending on the gene of interest. For example, we see spindle defects in PtK2 cells when we have reduced MCAK by 70% as shown by western immunoblot. However, CENP-A has to be reduced by >90% to cause mislocalization of other centromere proteins (15). 22. The ideal situation is that both the analysis of the phenotypic effects of knockdown and the assessment of knockdown efficiency by immunofluorescence would be performed on the same cell. We can do that for Eg5 RNAi, since both the Eg5 antibody and the anti-tubulin antibodies used to determine spindle defects work in methanol fix. However, this is not always possible because the antibodies used to assess both the knockdown and the phenotypic effects may require different fixation methods. In this case, we plate four coverslips per well so that two coverslips are used to determine knockdown and use the other two coverslips for the phenotypic analysis in order to reduce as many experimental differences as possible in the treatment of these cells.
Acknowledgments The authors would like to thank Susan Kline for early instruction in microinjection and suggestions on transfections of PtK2 cells. The authors would also like to thank Chantal LeBlanc for editing of the manuscript. Work in the Walczak lab is supported by NIH R01GM059618, an ACS Scholar award RSG CSM-106128, and in part by the Indiana METACyt Initiative of Indiana University, funded in part through a major grant from the Lilly Endowment, Inc. Rania Rizk is supported by a predoctoral fellowship from the American Heart Association.
References 1. Scholey, J. M. (1998). Functions of motor proteins in echinoderm embryos: an argument in support of antibody inhibition experiments. Cell Motil. Cytoskel. 39, 257–260.
2. Fire, A., Xu, S., Montgomery, M. K., Kostas, S. A., Driver, S. E. & Mello, C. C. (1998). Potent and specific genetic interference by double-stranded RNA in Caenorhabditis elegans. Nature 391, 806–811.
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3. Elbashir, S. M., Harborth, J., Lendeckel, W., Yalcin, A., Weber, K. & Tuschl, T. (2001). Duplexes of 21-nucleotide RNAs mediate RNA interference in cultured mammalian cells. Nature 411, 494–498. 4. Khodjakov, A., Copenagle, L., Gordon, M. B., Compton, D. A. & Kapoor, T. M. (2003). Minus-end capture of preformed kinetochore fibers contributes to spindle morphogenesis. J. Cell. Biol. 160, 671–683. 5. Khodjakov, A. & Rieder, C. L. (2006). Imaging the division process in living tissue culture cells. Methods 38, 2–16. 6. Stout, J. R., Rizk, R. S., Kline, S. L. & Walczak, C. E. (2006). Deciphering protein function during mitosis in PtK cells using RNAi. BMC Cell Biol. 7, 26. 7. Mayer, T. U., Kapoor, T. M., Haggarty, S. J., King, R. W., Schreiber, S. L. & Mitchison, T. J. (1999). Small molecule inhibitor of mitotic spindle bipolarity identified in a phenotypebased screen. Science 286, 971–974. 8. Weil, D., Garcon, L., Harper, M., Dumenil, D., Dautry, F. & Kress, M. (2002). Targeting the kinesin Eg5 to monitor siRNA transfection in mammalian cells. Biotechniques 33, 1244–1248. 9. Freshney, R. I. (2005). Culture of Animal Cells: A Manual of Basic Techniques (Freshney, R. I., ed.), Wiley-Liss, New Jersey.
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10. Phelan, M. C. (2003). Basic techniques for mammalian cell tissue culture. In Current Protocols in Cell Biology (Bonifacino, J. S., Dasso, M., Harford, J. B., LippincottSchwartz, J. & Yamada, K. M., eds.). WileyLiss, New Jersey. 11. Spector, D. L., Goldman, R. D. & Leinwand, L. A. (1998). Visualization of Organelles, Proteins, and Gene Expression. Cells: A Laboratory Manual, 3. 3 vols, Cold Spring Harbor Laboratory Press, New York. 12. Harlow, E. & Lane, D. (1988). Antibodies: A Laboratory Manual, Cold Spring Harbor Laboratory, Cold Spring Harbor, NY. 13. Harborth, J., Elbashir, S. M., Bechert, K., Tuschl, T. & Weber, K. (2001). Identification of essential genes in cultured mammalian cells using small interfering RNAs. J Cell Sci. 114, 4557–4565. 14. Caplen, N. J., Parrish, S., Imani, F., Fire, A. & Morgan, R. A. (2001). Specific inhibition of gene expression by small double-stranded RNAs in invertebrate and vertebrate systems. Proc Natl Acad Sci U S A 98, 9742–9747. 15. Liu, S. T., Rattner, J. B., Jablonski, S. A. & Yen, T. J. (2006). Mapping the assembly pathways that specify formation of the trilaminar kinetochore plates in human cells. J Cell Biol. 175, 41–53.
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Chapter 8 DNA Delivery by Microinjection for the Generation of Recombinant Mammalian Cell Lines Sebastien Chenuet, Madiha Derouazi, David Hacker and Florian Wurm Abstract Gene transfer methods for producing recombinant cell lines are often not very efficient. One reason is that the recombinant DNA is delivered into the cell cytoplasm and only a small fraction reaches the nucleus. This chapter describes a method for microinjecting DNA directly into the nucleus. Direct injection has several advantages including the ability to deliver a defined copy number into the nucleus, the avoidance of DNAses that are present in the cell cytoplasm, and the lack of a need for extensive subcloning to find the recombinant cells. The procedure is described for two cell lines, CHO DG44 and BHK-21, using green fluorescent protein as a reporter gene. However, this method could easily be adapted to other cells lines and using other recombinant genes. Key words: Recombinant cell line, microinjection, green fluorescent protein, transfection, gene expression.
1. Introduction Recombinant mammalian cell lines remain one of the most important tools for the production of complex recombinant proteins. The most popular gene transfer methods for establishing recombinant cell lines are based on chemical delivery vehicles such as calcium phosphate (CaPi), polyethylenimine (PEI), and liposomes. However, these methods result in low efficiencies of recombinant cell line generation. This is probably due to the fact that transfected DNA has to bypass several intracellular barriers before reaching the nucleus. Studies have shown that despite the delivery of a large
Sebastien Chenuet and Madiha Derouazi contributed equally to this work. David J. Carroll (ed.), Microinjection: Methods and Applications, Vol. 518 Ó 2009 Humana Press, a part of Springer ScienceþBusiness Media, LLC DOI 10.1007/978-1-59745-202-1_8
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amount of plasmid DNA to the cytoplasm of transfected cells only a small fraction reaches the nucleus (1–3). One explanation for these results is that the nuclear membrane constitutes a physical barrier to DNA entry. This observation is supported by studies based on viral and non-viral gene delivery methods (4–7). Passive transport into the nucleus of DNA molecules greater than 300 bp in length is very limited (8,9). In addition, microinjection of DNA into the cytoplasm results in expression of the reporter protein only after the cells undergo mitosis (10). As the result of this barrier, transfected plasmid DNA remains in the cytoplasm for a variable amount of time depending on the cell’s position in the cell cycle at the time of DNA uptake. Many studies suggest that the shorter the time between transfection and mitosis, the higher the transient recombinant protein expression level (11–13). A high rate of plasmid DNA degradation by cytoplasmic deoxynucleases (DNases) is observed, and it has been shown that single- and double-stranded DNA is degraded in the cytoplasm of HeLa and COS cells with an apparent half-life of about 90 min (3,14). A significant amount of internalized plasmid DNA is also targeted to the lysosomemediated degradation pathway after endocytosis (15). Thus, the amount of transfected plasmid DNA available in the nucleus for integration into the cell’s genome is limited. To overcome the limitations to DNA delivery by transfection, direct microinjection of DNA into the nucleus is an attractive option (16). Plasmid DNA can be injected directly into the nucleus with a defined plasmid copy number. As mentioned above, a high proportion of transfected DNA is targeted by cytoplasmic DNases, meaning that a portion of the DNA that reaches the nucleus may not be transcriptionally active due to cleavage within the recombinant gene. DNA microinjected directly into the nucleus is not affected by DNase degradation and thus a higher percentage of the recombinant genes may be functional. Thus, with chemical transfection methods neither the quantity nor the quality of the plasmid DNA that reaches the nucleus is controlled by the experimenter. In contrast, the number of plasmid molecules microinjected into the nucleus is defined, allowing cell lines to be generated in a reproducible way. Finally, a fundamental requirement for recombinant cell lines is that they are clonal in origin. For this reason, cell lines derived from transfection must be cloned, usually by one or more rounds of limiting dilution, before being considered for recombinant protein production. This long and tedious process can be eliminated with microinjection, since only one cell is injected at a time. Following genetic selection of recombinant cells, the survivors can be assumed to have originated from a single DNA integration event. Therefore the number of cells injected per plate can be maintained at a sufficiently low number to yield a single recombinant cell line per plate. By avoiding the subcloning step, microinjection reduces the development time for recombinant cell lines.
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Here we present the typical procedure to establish mammalian cell lines expressing a recombinant protein. Since plasmid copy number is a determining factor in the generation of cell lines by microinjection (9,10,16–18), this chapter describes the optimization of the different components affecting this parameter. We describe experiments performed with two cell lines, CHO DG44 and BHK-21, which are relevant to the biopharmaceutical industry. As shown here, GFP is a convenient reporter protein for the establishment of the physical parameters of microinjection and recombinant cell line establishment. However, the method can be easily adapted to other recombinant genes and cell lines.
2. Materials 1. Cells: CHO DG44 were routinely grown as a suspension culture in serum-free PROCHO5 medium (Cambrex, Belgium) containing 4-mM glutamine, 0.68 mg/l hypoxanthine, and 0.194 mg/l thymidine in 250-ml square-shaped glass bottles (Schott Glass, Mainz, Germany) (19). BHK-21 cells were routinely cultivated in T25 flask in DMEM/F12 medium (Cambrex) supplemented with 5% fetal calf serum (FCS). The cells were seeded for microinjection on 60-mm petri dish in DMEM/F12 medium containing 10% FCS and penicillin (10,000 units/ml), streptomycin (10 mg/ml), and amphotericin B (25 mg/ml). 2. DNA: The plasmid pMYK-EGFP-puro has the enhanced GFP gene expressed from the mouse cytomegalovirus immediate early promoter/enhancer joined to the first intron of the human elongation factor-1 gene (20). The plasmid also has the puromycin resistance gene expressed from the herpes simplex virus thymidine kinase (TK) promoter. For the assessment of the injection efficiency a solution of DNA at 3 mg/ml containing 30% FITC (150 kDa) was prepared. 3. Microinjection apparatus: A semi-automatic Femtojet (Eppendorf AG, Hamburg, Germany) was connected to an InjectMan NI 2 (Eppendorf AG) mounted on a motorized Axiovert 200 M inverted fluorescence microscope (Carl Zeiss AG, Oberkochen, Germany). External vibrations were avoided by placing the microscope on a Benchtop breadboard (Newport Corporation, Irvine, CA). Injections were performed with Femtotips II (Eppendorf AG) with an inner diameter of 500 nm and an outer diameter of 700 nm.
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3. Methods 3.1. Plasmid Purification
1. Purify plasmid DNA on a Nucleobond AX anion exchange column and linearize it with a single-site restriction endonuclease. 2. Remove proteins and lipids by phenol-chloroform extraction and recover the DNA by ethanol precipitation in the presence of sodium acetate. 3. After centrifugation, resuspend the DNA pellet in ultra high purity (UHP) water at 1 mg/ml. For microinjection, the DNA is diluted with UHP water to the working concentration. This should be less than 400 mg/ml to avoid clogging the microinjection capillary due to the high viscosity of the DNA solution. If clogging is observed at DNA concentrations lower than 400 mg/ml, then traces of salt in the DNA can be eliminated by purifying the DNA on Qiagen plasmid purification membrane according to the manufacturer’s instructions. 4. Before each experiment, centrifuge the DNA solution for 10 min at 8000 rpm to sediment insoluble impurities.
3.2. Cell Preparation
1. On the day before microinjection, the cells are trypsinized and recovered by centrifugation. 2. The cells are then suspended in fresh DMEM/F12 medium with 10% FCS and plated at a density of 25,000 cells per 60mm dish. At this plating density, the contact between adjacent cells is minimal. 3. After microinjection, the medium is removed and replaced with fresh DMEM/F12 medium supplemented with 10% FCS and penicillin (10,000 units/ml), streptomycin (10 mg/ml), and amphotericin B (25 mg/ml).
3.3. Improving Parameters Affecting Injection Efficiency
3.3.1. Capillary Position
The injection of a cell depends on the ability to perforate both the plasma and nuclear membranes and to force the DNA solution into the nucleus. The efficiency of the perforation depends mainly on the angle of injection, whereas the entry of the DNA solution is mainly affected by the pressure of injection. 1. Depending on the cell morphology, determine the appropriate angle of injection. For cells with a flattened morphology, take an injection angle as close as possible to the vertical (see Note 1). 2. Determine the height of injection so as to perforate both the cell and nuclear membrane.
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3. Start the injections at a height of 1 mm above the cell surface and decrease or increase it by increments of 500 nm, if after 2–3 attempts a cell is not injected. 4. Repeat with another cell having the same morphology (see Note 2). 5. Determine the depth of injection so as the capillary should penetrate deep enough to enter the nucleus but without damaging it (see Note 3).
3.3.2. Pressure of Injection
1. To test the effect of injection pressure on the injection efficiency, use a solution of DNA (3 mg/ml) containing 30% 150 kDa FITC (see Note 4). 2. Keep the injection time at 0.1 s and increase the injection pressure incrementally from 20 hPa to 160 hPa. 3. Determine by microscopic observation if the DNA solution is injected into the nucleus. 4. Confirm the presence of FITC in injected cells with the appropriate fluorescence excitation source with emission and excitation wavelengths of 485 nm and 530 nm, respectively (see Note 5). 5. The number of attempts to inject a cell should not exceed 3–4 times. More injections have a negative impact on cell viability. If a cell is not successfully injected after four attempts, then repeat the procedure with another one. However, these unsuccessful events should be included in the total number of cells microinjected when assessing the efficiency of injection. 6. To deliver a constant flow of solution into the cells, change the capillary after injecting 15–20 cells (see Note 6). 7. To assess the efficiency of injection as a function of pressure, determine for each injection pressure tested the number of FITC-positive cells relative to the number of cells that were injected (see Note 7 and Fig. 8.1). 8. To assess the effect of injection pressure on cell damage, determine by microscopic observation the proportion of nuclei damaged after microinjection. The damage is evident as either a bursting of the nuclear compartment during the injection or by the formation of apoptotic vesicles on the surface of the cells a few seconds after injection.
3.4. Improving Parameters Affecting Gene Expression Efficiency
The expression of the recombinant gene depends on the plasmid copy number delivered to the nucleus. The copy number in turn is a function of the concentration of plasmid DNA and the volume of injection. The latter is controlled by the pressure
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Fig. 8.1. The effect of injection pressure on injection efficiency and nuclei damaged. The injection efficiency was assessed by injecting BHK-21 cells with a solution of plasmid DNA at 3 mg/ml containing 0.5 mg/ml FITC dextran. FITCpositive cells were detected by fluorescence microscopy 20 h after injection. Experiments were performed in duplicate and 50–60 cells were injected per experiment.
and duration of injection. Here we present a typical procedure to optimize the volume of injection and the DNA concentration for efficient transient gene expression. 1. On the bottom of 60-mm petri dishes, draw 25–30 circles of approximately 3–4 mm diameter. 2. Plate the cells as described above on the day before microinjection. 3. Regardless of the parameter being tested, only inject one cell per circle (see Note 8). 4. To maintain cell viability, the cells should not remain outside the incubator for more that 40–50 min. 5. After injection, replace the medium with fresh DMEM/ F12 medium containing 10% FCS and penicillin (10,000 units/ml), streptomycin (10 mg/ml), and amphotericin B (25 mg/ml). 6. To determine the optimum injection pressure, use an injection time of 0.1 s and a DNA concentration of 3 mg/ml. For each injection pressure tested, inject about 100–150 cells and change the injection pressure incrementally. 7. Assess the efficiency of gene expression the day after injection by observing the cells under UV light. Score the number of
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Fig. 8.2. The effect of injection pressure on transient gene expression in CHO DG44 (A) and BHK-21 (B) cells. GFP-positive cells were detected by fluorescence microscopy at 20 h after microinjection. Cell division of BHK-21 cells was evaluated by visual inspection of colonies of GFP-positive cells at 5 days after microinjection. Experiments were performed in triplicate and 50–60 cells were injected per experiment.
GFP-positive cells with respect to the number of cells injected (see Note 9 and Fig. 8.2). 8. Assess the proportion of injected cells that are able to grow by observing the cells microscopically over a period of 4–5 days for their ability to generate small colonies composed of GFP-positive cells (see Note 9 and Fig. 8.2B). 9. For each injection pressure yielding a high efficiency of gene expression with good cell viability and growth, determine the optimal duration of injection. 10. Use a DNA solution with a concentration of 3 mg/ml and test different durations of injection by incrementally increasing the time by 0.1 s from 0.1 s to 0.5 s. 11. As in the steps above, determine both the number of GFPpositive cells per the number of cells injected and the number of small GFP-positive colonies per the number of cells injected (see Note 10). 12. To determine the optimal DNA concentration, begin with the optimal conditions for the injection pressure and duration. 13. As a preliminary study, begin with a broad range of DNA concentrations such as 3 mg/ml, 30 mg/ml, and 300 mg/ml. Score the results as in steps 4 and 5. 14. Based on the results, use a narrower window of DNA concentrations around the one yielding the highest efficiency of transient reporter protein expression (see Note 11 and Fig. 8.3).
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Fig. 8.3. The effect of DNA concentration on transient gene expression in CHO DG44 (A) and BHK-21 cells (B). The cells were injected with pMYK-EGFP-puro as described in the text. GFP-positive cells were detected by fluorescence microscopy the day after microinjection. Cell division of BHK-21 cells was evaluated by visual inspection of colonies of GFP-positive cells at 5 days after microinjection.
3.5. Establishment of Cell Lines by Microinjection
3.5.1. Determination of the DNA Concentration to Optimize the Efficiency of Stable Gene Transfer
Here stable gene transfer is defined as the stable integration of the recombinant gene and the selectable gene into the genome of the host cell. The optimal efficiency of stable gene transfer can be assessed for different DNA concentrations, injection pressures, and injection durations. Here we present the procedure to optimize the efficiency of stable gene transfer as a function of the DNA concentration. 1. The day prior to injection, plate the cells at a density of 25,000 cells per 60-mm petri dish with 25 circles drawn on the bottom as described earlier. 2. To inject the cells, use the pressure and duration of injection yielding the highest efficiency of gene expression and cell growth and viability. 3. As a preliminary study, test a few concentrations of DNA, such as 3 mg/ml, 30 mg/ml, and 300 mg/ml, as described above. 4. Inject a limited number of cells, typically two cells per circle, per petri dish (see Note 12). 5. After injection, remove the medium and add DMEM/F12 medium supplemented with 10% FCS. 6. Two days after injection, remove the medium and add a combination of fresh medium (75%) and conditioned medium (25%) containing the appropriate drug for selection of stably transfected cells. In our experiments, we used puromycin at concentrations of 3 mg/ml and 6 mg/ml for selection of CHO DG44 and BHK-21, respectively. 7. Change the medium every two days as in Step 6 for the first six days post-injection to eliminate dead cells.
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8. After 10–14 days, determine the number of large colonies (3–4-mm diameter). The efficiency of stable gene transfer is assessed by the number of colonies with respect to the number of cells injected (see Note 13 and Table 8.1). 3.5.2. Recovery of Recombinant Cell Lines Generated by Microinjection
1. After assessing the efficiency of stable gene transfer, isolate the recombinant cell lines established under the different DNA concentrations by picking the colonies with a micropipette tip. 2. Transfer individual colonies into a 24-well plate. 3. When the cells are 80% confluent (usually after 7–10 days), transfer them to a six-well plate in fresh medium with the selective agent. 4. When the cells reach 70% confluence, use a portion of the cells for establishing a cell bank (usually 4–5 days after transfer to a six-well plate) using standard methods for cell freezing. 5. Transfer the remainder of the cells to a clean six-well plate by keeping the selective agent in the medium.
3.5.3. Assessment of the Productivity and Stability
1. Trypsinize the cells when they reach 70% confluence in a sixwell plate. 2. Centrifuge the cells at 800 rpm for 5 min and resuspend the pellet in PBS containing 10% FCS at a concentration of 200,000 cells/ml. 3. Transfer 2 ml of the suspension into a 5-ml disposable plastic tube. 4. Quantify the level of GFP expression of the clones by analyzing the cells with a flow cytometer (see Note 14 and Fig. 8.4). 5. To study the stability of recombinant protein expression over time, remove the selective agent from the medium and passage the cells every two weeks in six-well plates for 4–5 months.
Table 8.1 Effect of DNA concentration on stable gene transfer efficiency in BHK-21 cells DNA concentration (mg/ml)
3
30
300
Number of large coloniesa
2
11
4
Number of cells injected
1543
952
1221
Efficiency of stable transduction (%)
0.13
1.15
0.32
a The number of puromycin-resistant colonies was determined 2 weeks after puromycin was added to the microinjected cells.
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Fig. 8.4. Flow cytometry profiles for GFP expression of clones established by microinjection of BHK-21 cells with different DNA concentrations. Flow cytometry was performed on clonal GFP-positive cell lines at 25–30 days after the addition of puromycin to the culture. For each concentration of DNA tested, two cell lines were taken as example.
6. Assess the stability of the pattern of GFP expression by analyzing the cells with a flow cytometer every two weeks.
4. Notes 1. For cells with a rounded morphology the efficiency of injection is not that much affected by the angle of injection. However, for cells with a flattened morphology, especially BHK-21 cells, we found that the efficiency of injection slightly increased when the angle of injection was shifted from 45° to 76° (maximal angle for the apparatus described above). 2. Because cell morphology in a population is variable, it is difficult to provide a general rule regarding the height of injection. However, for cells with a rounded and contracted morphology, a low height is not efficient for injection. For
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very flat cells a height of injection that is too great may damage the cells. 3. It is again difficult to propose a general rule regarding the depth of injection into the nucleus. However, to target this compartment, the level of injection should be between 800 nm and 2000 nm below the cell surface, since you go first through the cytoplasm. 4. FITC was mixed with the expression vector pMYK-EGFPpuro. Though this plasmid encodes a protein with the same excitation and emission wavelengths as FITC, this was not a problem since here we assessed the entry of FITC into cells immediately after microinjection. GFP was not yet expressed at this time. 5. The entry of the DNA into the nucleus is seen as a brief flow wave during which the brightness in the immediate neighborhood of the capillary increases. With experience, it becomes possible to distinguish the increase in brightness due to the entry of the DNA solution into the nucleus from the increase of brightness due to the membrane deformation caused by contact of the capillary with the cell surface. The ability to distinguish these two phenomena allows assessment of successful injection when a solution of DNA is injected without any fluorescent marker. This is necessary so as to prevent the under- or overestimation of the efficiency of successful microinjection with plasmid alone. 6. The flow through the capillary might be perturbed by clogging or physical damage. Frequently changing capillaries assures a constant flow and allows better reproducibility of injection. The cleaning function of the Femtojet should also be routinely used for cleaning. 7. The range of possible injection pressures is limited since low pressures do not support DNA delivery while high pressures result in cellular damage. With a DNA concentration of 3 mg/ ml and an injection time of 0.1 s, we incrementally increased the injection pressure from 20 hPa to 160 hPa. For BHK-21 cells, at 20 hPa and 40 hPa more than 60% of the injected cells were FITC-positive (Fig. 8.1). At these pressures, more than 90% of the cells were FITC-positive, the nucleus was not damaged, and the cell surface did not display apoptotic vesicles (Fig. 8.1). With 80 hPa, only 70% of the injected cells remained intact whereas this proportion dropped to 50% for an injection pressure of 160 hPa (Fig. 8.1). 8. Injection of a single cell per circle avoids confusing GFPpositive cells resulting from cell division with those resulting from the initial injection.
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9. With an injection time of 0.1 s the proportion of GFP-positive BHK-21 cells increased with the injection pressure from a low of 30% at 20 hPa to a high of 80% at 80 hPa (Fig. 8.2B). At higher injection pressures the efficiency decreased to 50% (Fig. 8.2B). Although injection pressures of 20–40 hPa did not give rise to the highest proportion of GFP-positive cells, the number of cells injected at 40 hPa that expressed GFP and were able to divide was 1.8 times higher than for the injections performed at 80 hPa. Injection performed in CHO DG44 at the lowest pressure tested (70 hPa) yielded also the highest proportion of GFP-positive cells (Fig. 8.2A). Hence, injections performed at low pressures are more efficient for establishing recombinant cell lines than those performed at high pressures (80 hPa and higher). 10. To evaluate the effect of injection duration on the efficiency of transient GFP expression we performed experiments with injection times of 0.1–0.5 s with BHK-21 cells. Only for the low injection pressures (20–40 hPa) was the transient gene expression increased with an injection time of 0.5 s as compared to 0.1 s. 11. To evaluate the effect of DNA concentration on transient reporter protein expression, we injected BHK-21 cells with pMYK-EGFP-puro at concentrations ranging from 3 mg/ml to 300 mg/ml with an injection duration of 0.5 s and a pressure of 40 hPa. The percentage of GFP-positive cells increased with DNA concentration and eventually reached a plateau of roughly 95% (Fig. 8.3B). Fluorescence microscopy confirmed that the intensity of the GFP signal correlated to the amount of plasmid copies delivered to the nucleus. For injections performed with 3 mg/ml and 30 mg/ml of pMYK-EGFP-puro, more than 85% of the cells were able to divide and form colonies within the first 4 days after DNA transfer (Fig. 8.3B). In contrast, cells injected with a DNA concentration of 300 mg/ml had an extended cell cycle and 40% of the cells were unable to give rise to a colony by 4 days after injection (Fig. 8.3B). 12. By limiting the number of cells injected per plate, the frequency of obtaining multiple stably transfected cells per dish is reduced. Secondly, it limits the time that the plate remains out of the incubator. Ideally, the number of cells injected per dish should be far below the efficiency of stable gene transfer so that a maximum of one clone per petri dish is obtained. This avoids the mixing of cells between different colonies and thus eliminates a subcloning step. To recover a single clone in a dish we typically injected 40–50 cells per dish. Furthermore, this method allows assessing the efficiency of stable gene transfer with a better accuracy.
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13. To assess the efficiency of stable gene transfer as a function of plasmid copy number, BHK-21 cells were injected either with 3 mg/ml, 30 mg/ml, or 300 mg/ml pMYK-EGFP-puro. The percentage of stable recombinants for each DNA concentration is shown in Table 8.1. 14. The clones analyzed by flow cytometry displayed a homogenous expression of GFP under selective pressure (Fig. 8.4). The level of GFP depended on the plasmid copy number delivered to the nucleus by microinjection. Clones obtained by injection of a low DNA concentration (3 mg/ml) had the lowest level of GFP expression (Fig. 8.4). When injections were performed with 30 mg/ml or 300 mg/ml of DNA, the level of GFP expression was 10–50 times higher than at the lowest DNA concentration (Fig. 8.4).
References 1. Coonrod A., Li F.Q., Horwitz M. (1997) On the mechanism of DNA transfection: efficient gene transfer without viruses. Gene Ther. 4, 1313–1321. 2. James M.B., Giorgio T.D. (2000) Nuclearassociated plasmid, but not cell-associated plasmid, is correlated with transgene expression in cultured mammalian cells. Mol. Ther. 1, 339–346. 3. Lechardeur D., Sohn K.J., Haardt M., Joshi P.B., Monck M., Graham R.W., Beatty B., Squire J., O’Brodovich H., Lukacs G.L. (1999) Metabolic instability of plasmid DNA in the cytosol: a potential barrier to gene transfer. Gene Ther. 6, 482–497. 4. Lewis P.F., Emerman M. (1994) Passage through mitosis is required for oncoretroviruses but not for the human immunodeficiency virus. J. Virol. 68, 510–516. 5. Miller D.G., Adam M.A., Miller A.D. (1990) Gene transfer by retrovirus vectors occurs only in cells that are actively replicating at the time of infection. Mol. Cell. Biol. 10, 4239–4242. 6. Mortimer I., Tam P., MacLachlan I., Graham R.W., Saravolac E.G., Joshi P.B. (1999) Cationic lipid-mediated transfection of cells in culture requires mitotic activity. Gene Ther. 6, 403–411. 7. Wilke M., Fortunati E., van den Broek M., Hoogeveen A.T., Scholte B.J. (1996) Efficacy of a peptide-based gene delivery system depends on mitotic activity. Gene Ther. 3, 1133–1142.
8. Ludtke J.J., Sebestyen M.G., Wolff J.A. (2002) The effect of cell division on the cellular dynamics of microinjected DNA and dextran. Mol. Ther. 5, 579–588. 9. Ludtke J.J., Zhang G., Sebestyen M.G., Wolff J.A. (1999) A nuclear localization signal can enhance both the nuclear transport and expression of 1 kb DNA. J Cell Sci. 112, 2033–2041. 10. DerouaziM.,FlactionR.,GirardP.,deJesusM., Jordan M., Wurm F.M. (2006) Generation of recombinant Chinese hamster ovary cell lines by microinjection. Biotechnol. Lett. 28, 373–382. 11. Brunner S., Sauer T. Carotta S., Cotten M., Saltik M., Wagner E. (2000) Cell cycle dependence of gene transfer by lipoplex, polyplex and recombinant adenovirus. Gene Ther. 7, 401–407. 12. Escriou V., Ciolina C., Lacroix F., Byk G., Scherman D., Wils P. (1998) Cationic lipidmediated gene transfer: effect of serum on cellular uptake and intracellular fate of lipopolyamine/DNA complexes. Biochim. Biophys. Acta 1368, 276–288. 13. Grosjean F., Batard P., Jordan M., Wurm F.M. (2002) S-phase synchronized CHO cells show elevated transfection efficiency and expression using CaPi. Cytotechnology 38, 57–62. 14. Pollard H., Toumaniantz G., Amos J.L., AvetLoiseau H., Guihard G., Behr J.P., Escande D. (2001) Ca2+-sensitive cytosolic nucleases prevent efficient delivery to the nucleus of injected plasmids. J. Gene Med. 3, 153–164.
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15. Wolff J.A., Budker V. (2005) The mechanism of naked DNA uptake and expression. Adv. Genet. 54, 3–20. 16. Capecchi M.R. (1981) High efficiency transformation by direct microinjection of DNA into cultured mammalian cells. Cell. 22, 479–488. 17. Folger K.R., Wong E.A., Wahl G., Capecchi M.R. (1982) Patterns of integration of DNA microinjected into cultured mammalian cells: evidence for homologous recombination between injected plasmid DNA molecules. Mol. Cell. Biol. 2, 1372–1387.
18. Dean D.A., Dean B.S., Muller S., Smith L.C. (1999) Sequence requirements for plasmid nuclear import. Exp. Cell. Res. 253, 713–722. 19. Muller N., Girard P., Hacker D.L., Jordan M., Wurm F.M. (2005) Orbital shaker technology for the cultivation of mammalian cells in suspension. Biotechnol. Bioeng. 89, 400–406. 20. Hacker D.L., Derow E., Wurm F.M. (2005) The CELO adenovirus Gam1 protein enhances transient and stable recombinant protein expression in Chinese hamster ovary cells. J Biotechnol. 117, 21–29.
Chapter 9 Bacteriophage fC31 Integrase Mediated Transgenesis in Xenopus laevis for Protein Expression at Endogenous Levels Bryan G. Allen and Daniel L. Weeks Abstract Bacteriophage fC31 inserts its genome into that of its host bacterium via the integrase enzyme which catalyzes recombination between a phage attachment site (attP) and a bacterial attachment site (attB). Integrase requires no accessory factors, has a high efficiency of recombination, and does not need perfect sequence fidelity for recognition and recombination between these attachment sites. These imperfect attachment sites, or pseudo-attachment sites, are present in many organisms and have been used to insert transgenes in a variety of species. Here we describe the fC31 integrase approach to make transgenic Xenopus laevis embryos. Key words: Xenopus, fC31, integrase, transgenesis, fluorescence.
1. Introduction The frog, Xenopus laevis, has a long history of use for studies in embryonic development. Previously described Xenopus integration techniques often insert multiple copies of a transgene at random sites in the embryo’s genome (1–4). Though valuable for many experimental designs, these approaches are problematic for researchers who desire transgene expression to approximate endogenous gene expression levels. Rather, a site-directed integration approach that incorporates a regulated single copy of a transgene into the host genome would more closely match endogenous gene expression. The fC31 integrase approach is one way to accomplish this aim. Bacteriophage fC31 encodes an integrase enzyme that inserts the phage genome into the genome of various Streptomyces bacteria David J. Carroll (ed.), Microinjection: Methods and Applications, Vol. 518 Ó 2009 Humana Press, a part of Springer ScienceþBusiness Media, LLC DOI 10.1007/978-1-59745-202-1_9
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(5, 6). The integrase protein recognizes a 39-bp-long phage attachment site (attP) in its own genome and a 34-bp-long bacterial attachment site (attB) in the bacterium’s genome and then catalyzes an integration event (7, 8). The integrase enzyme does not require perfect sequence fidelity to recognize the attP site (7). Non-perfect attP sites, or pseudo-attP sites, may have as low as 24% sequence homology to endogenous attP sites and still allow recombination (7) although the recombination efficiency may be decreased. Many groups have shown that integrase can insert plasmid DNA sequences that contain an attB site into pseudo-attP sites found in a variety of organisms. Utilizing this approach, transgenes have been inserted into the genomes of plant cells (9), mammalian cells (7,10–18), and Drosophila embryos (19). We used the pseudo-attP sites in the Xenopus genome and an attB site containing reporter plasmid to make transgenic Xenopus embryos (20) (Fig. 9.1). Surprisingly, the reporter genes are expressed in some expected tissues
Fig. 9.1. Representation of fC31 integrase-mediated transgenesis in Xenopus laevis. The reporter plasmid containing an attB site and an insulated reporter gene is injected along with integrase mRNA into single-cell embryos. The chromosomes (thick black lines) contain numerous pseudo-attP sites. Inside the single-cell embryo, the integrase protein catalyzes recombination between the attB site in the reporter plasmid (thin black line) and a pseudo-attP site in the embryo’s genome (thick black line). Recombination results in the formation of two new attachment sites, attR and attL flanking the integrated reporter plasmid.
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Fig. 9.2. fC31 integrase-mediated transgenesis of insulated reporter plasmids generate Xenopus embryos with tissueappropriate expression. In every case, the insert shows a brightfield image of the embryo. (A) Non-injected stage 42 embryo. (B) Stage 42 embryo injected with 5 pg of CMV-EGFP-DI-attB reporter plasmid. (C) Stage 42 embryo injected with 5 pg of CMV-EGFP-DI-attB reporter plasmid and 1 ng of integrase mRNA. (D) Stage 44 embryo injected with 5 pg of CL-EGFP-DI-attB plasmid alone. (E) Stage 44 embryo injected with 5 pg of CL-EGFP-DI-attB plasmid and 1 ng of integrase mRNA. GFP expression is indicated with the white arrow. (F) Southern blot demonstrating single integration events. Lanes 1 and 2 contains DNA harvested from single stage 46 CMV-EGFP-DI-attB transgenic embryos that expressed GFP uniformly. Lanes 3–4 contain DNA harvested from single stage 46 CL-EGFP-DI-attB transgenic embryos that expressed GFP in the lens of the eye. Lane 5 contains stage 46 non-injected DNA. Lane 6 contains DNA from a stage 46 embryo injected with CMV-EGFP-DI-attB plasmid alone. Lane 7 contains DNA from a stage 46 embryo injected with CL-EGFP-DI-attB plasmid alone. Lane 8 contains 10 pg of CMV-DI-EGFP-attB plasmid linearized with BamHI. Markers in kilobase pairs are indicated to the left of the membrane.
but not in others. We recognized this as chromatin position effect and flanked the reporter gene with HS4 insulators. HS4 insulators stop the spread of chromatin silencing and also prevent distant enhancers from acting on a promoter (21, 22). After making transgenic embryos with the new insulated reporter plasmid, we found that the transgenes expressed as expected (20) (Fig. 9.2). The techniques used to generate and recognize fC31 integrase mediated transgenic Xenopus embryos are described below.
2. Materials 2.1. Plasmids and Plasmid Preparation Reagents
1. Plasmids: The integrase plasmid pET11-phiC31poly(A) (23) can be obtained from Dr. Michelle Calos (calos@stanford. edu), and the attB reporter plasmids CMV-EGFP-DI-attB and CL-EGFP-DI-attB (20) can be obtained from Dr. Daniel
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[email protected]). Permission to use HS4 insulator sequences and HS4 insulator constructs needs to be obtained from Dr. Gary Felsenfeld (gary.felsenfeld@nih. gov) (see Note 1). 2. Plate and culture reagents: Bacto-Yeast Extract, Bacto-Tryptone powder, and Bacto-agar (BD Biosciences, Sparks, MD). 3. Antibiotics: Ampicillin and Kanamycin (Sigma Aldrich, St. Louis, MO). 4. Plasmid Purification Kits: Qiaprep Spin Miniprep Kit, HiSpeed Plasmid Maxi Kit (Qiagen, Valencia, CA). 5. In vitro RNA transcription: T7 mMessage machine (Ambion, Austin, TX). 6. Agarose (RPI, Mt. Prospect, IL). 7. 10X TAE: 0.4-M Tris-acetate, 0.01-M EDTA. 8. 10X Non-denaturing DNA loading buffer: 50% glycerol, 60mM EDTA, 1% SDS, and 0.05% bromophenol blue. 9. 1% agarose, 2.2-M formaldehyde MOPS gel. 10. 10X Denaturing RNA loading buffer. 11. Gel electrophoresis apparatus for flat-bed agarose gel electrophoresis (Owl Separation Systems, Portsmouth, NH). 12. Spectrophotometer: Nanodrop ND1000 (Nanodrop Technologies, Wilmington, DE). 13. UV-light transilluminator.
2.2. Xenopus Injection
1. Xenopus laevis: May be obtained from either Xenopus I (Dexter, MI) or Nasco (Fort Atkinson, WI). 2. Human Chorionic Gonadotropin (Sigma Aldrich). 3. Tricaine (3-aminobenzoic acid ethyl ester) (Sigma Aldrich). 4. 10X Marc’s Modified Ringers Solution (MMR): 1-M NaCl, 20-mM KCl, 20-mM CaCl2, 10-mM MgCl2, 50-mM HEPES at pH 7.4. 5. Injection buffer: 88-mM NaCl, 10-mM HEPES. 6. 0.3X MMR with 3% Ficoll Type 400 (Sigma Aldrich). 7. 2% Cysteine (RPI) made in dH2O, pH 7.8–7.9 with NaOH. Should be made on the day of injection. 8. Microinjection needle puller. 9. Micromanipulator: Singer MK-1 (Singer instrument company, Somerset, England) or similar instrument. 10. Microinjector: Inject+Matic (Geneva, Switzerland) or similar instrument.
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11. Glass capillary tubes (Singer). 12. 18°C incubator. 2.3. Microscopy
1. Dissecting microscope: Nikon SMZ (Nikon Instruments, Melville, NY) and Zeiss Stemi SV 11 fluorescent dissecting microscope (Zeiss MicroImaging, Thornwood, NY). 2. Compound microscope with fluorescence: Zeiss Axioplan 2 (Carl Zeiss MicroImaging) or similar instrument. 3. Camera for photo documentation: SPOT camera (Diagnostic Instruments, Sterling Heights, MI), Zeiss Axiocam (Carl Zeiss MicroImaging), or similar instrument.
2.4. Southern Blotting
1. Genomic DNA purification: Qiagen DNeasy (Qiagen). 2. Restriction enzymes: multiple suppliers. 3. Ethidium bromide (Sigma Aldrich). 4. Hybond-N+ nylon membrane (GE Healthcare, Chicago, IL). 5. Rediprime II DNA Labeling System (GE Healthcare). 6. Redivue a-P32 deoxycytidine (GE Healthcare). 7. RapidHyb solution (Ambion). 8. X-ray film, Kodak Biomax XAR (Eastman Kodak Co, Rochester, NY). 9. 20X SSC: 3.0-M NaCl and 0.3-M sodium citrate at pH 7.0. 10. Depurination buffer: 0.25-M HCl. 11. Denaturation buffer: 1.5-M NaCl with 0.5-M NaOH. 12. Neutralization buffer: 1-M Tris-HCl, pH 8.0, 1.5-M NaCl. 13. Hybridization Oven: Hybaid (Thermo Corporation, Waltham, MA).
3. Methods 3.1. Preparation of Plasmids and fC31 Integrase mRNA
1. Transform the pET11-phiC31poly(A) plasmid and the attB reporter plasmids (CMV-EGFP-DI-attB and CL-EGFP-DIattB) into Escherichia coli and plate onto antibiotic selective plates using standard techniques. The pET11-phiC31poly(A) plasmid contains an ampicillin resistance gene and the attB reporter plasmids contain a kanamycin resistance gene. We have found that insulated attB reporter plasmids frequently undergo rearrangements and thus recommend using Stbl2 cells (Invitrogen) which are designed to prevent recombination.
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2. Select individual colonies on each plate and grow 3-ml cultures at 37°C overnight with agitation. Use 1 ml for mini-preps (Qiaprep mini-prep kit) and save the other 2 ml of culture at 4°C. Using the mini-prep-generated DNA, confirm plasmid size and insulator orientation by restriction enzyme digestion and gel analysis. Once plasmid size and insulator orientation are confirmed, inoculate 1 l of LB broth containing the appropriate antibiotic with the remaining 2 ml of culture. Grow at 37°C with agitation overnight. 3. Perform maxi-preps on the 1-l cultures following the manufacturer’s instructions (Hi Speed Plasmid Maxi Kit). Do not add RNase to buffer P1. 4. Confirm maxi-prep-generated plasmid sequences by restriction enzyme digestion and gel analysis. Store the maxi-prepgenerated DNAs at 20°C. 5. Linearize 5 mg of pET11-phiC31poly(A) maxi-prep DNA with either BamHI or EcoRI restriction enzymes. Run approximately one-tenth of the digestion on a 1% agarose gel to ensure linearization. Heat inactivate the restriction enzyme in the remaining portion of digestion by heating reaction to 65°C for 20 min. Precipitate the DNA by adding one-tenth the volume of 5-M NH4 acetate and 2 volumes of ethanol. Place solution at 20°C for 15 min and centrifuge at 10,000 g for 15 min. Remove the supernatant and resuspend the pellet in 10 ml of RNase-free TE buffer. 6. Synthesize the fC31 integrase mRNA using the T7 mMessage machine following the manufacturer’s instructions. Because the protocol includes a DNase treatment step, the DNase needs to be removed or inactivated. We routinely follow the LiCl precipitation protocol described in the manufacturer’s instructions. Resuspend the integrase mRNA in RNase-free water at a concentration of 1 mg/ml. 7. Run 1 mg of fC31 integrase mRNA on a formaldehyde MOPS gel in 1X MOPS buffer to ensure that the transcript is approximately 1.9 kilobases long. 8. Store fC31 integrase mRNA (for up to 1 month ) at 80°C until it is needed.
3.2. Injection of Xenopus Embryos
1. Obtain approval from host institution to house and care for Xenopus adult frogs and embryos. If Xenopus experience is minimal, the book, Early Development of Xenopus laevis – A Laboratory Manual (24), may be a useful reference to consult. 2. Induce Xenopus females to lay eggs by injecting 1 ml (1000 IU) of human chorionic gonadotropin into the dorsal lymph sac the night before desired day of egg collection.
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3. The next morning, the cloaca on the injected females should be swollen and there may be eggs in the water tank holding the frogs. 4. After eggs are present in the tank, inject a lethal dose of Tricaine (1 ml of a 10% solution) into the dorsal lymph sac of a Xenopus male and then surgically remove the testes. Store the testes in 1X MMR. 5. Induce egg laying into a dry petri dish by spreading apart the female’s legs and gently squeezing and rubbing the pelvic region. Immediately fertilize the eggs by rubbing a small piece of testis (one-sixth of a testis) through the eggs. Then crush the testis in 1 ml of 0.3X MMR and spread this solution over the eggs. Let the eggs sit in the sperm solution for about 1 min and then flood the eggs with 0.1X MMR. 6. Successful fertilization can be determined if the eggs align with the animal pole (pigmented half) facing up. Embryos should be spherical, uniform size, and have smooth even pigmentation of the animal hemisphere. At 30 min after fertilization, remove the 0.1X MMR and place the embryos in 2% cysteine, pH 7.8–7.9 for 2 min to remove the jelly coats. 7. Remove the cysteine and wash the embryos in three 5-ml washes of 0.3X MMR. 8. Place the embryos in 0.3X MMR, 0.3% Ficoll. 9. Inject single-cell embryos into the center of the animal hemisphere with either 10 nl of injection solution, 10 nl containing 5 pg of reporter plasmid resuspended in injection solution, or 10 nl containing 5 pg of reporter plasmid + 1 ng of fC31 integrase mRNA also resuspended in injection solution. 3.3. Monitoring Developing Embryos for Transgenesis
1. Allow injected embryos to develop at 18°C in 0.3X MMR, 0.3% Ficoll for approximately 6–8 h after injection and then transfer to 0.3X MMR. 2. Remove delaminating or dead embryos and provide fresh 0.3X MMR at least twice a day for the first 3 days. 3. On the second day after fertilization, begin to monitor the embryos for GFP expression using a fluorescent microscope optimized to detect green fluorescence (see Note 2). Embryos injected with 1 ng integrase mRNA and 5 pg of CMV-EGFPDI-attB should express GFP uniformly (Fig. 9.2C) while embryos injected with 1-ng integrase mRNA and 5 pg of CL-EGFP-DI-attB should express GFP only in the lens of the eye (Fig. 9.2E). Embryos injected with 5 pg of reporter plasmid alone rarely give GFP expression (Fig. 9.2B). Digital photography with long exposure times may detect fluorescence before it is visible to the eye.
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4. Once fluorescence is detected, photograph the embryos using a fluorescent microscope with a digital microscope. If embryo movement prevents photography, consider anesthetizing the embryos with 0.02% Tricaine. Embryos may be exposed to the Tricaine solution for approximately 5 min; longer exposure times may lead to the demise of the embryos. Embryos should be placed back into 0.3X MMR after photography. 3.4. Southern Blot Analysis to Confirm Integration
1. Isolate DNA from single-stage 46 embryos using the Qiagen DNeasy kit following the manufacturer’s instructions. Onestage 46 embryo should yield approximately 50 mg of DNA. DNA should be collected from non-injected embryos, embryos injected with reporter plasmid alone, and transgenic embryos determined by green fluorescence. 2. Digest 5–10 mg of harvested DNA and 10–100 pg of reporter plasmid with a restriction enzyme that cuts the reporter plasmid at a single site overnight (for CL-EGFP-DI-attB or CMV-DI-EGFP attB, BamHI is a suitable enzyme). 3. Run digested DNA on a 0.6% agarose gel. To allow for good separation, use a gel that is 10–15-cm long and run the gel until the bromophenol blue dye front is approximately 2 cm from the bottom of the gel. Stain the gel with ethidium bromide and then photograph the gel while on a UV transilluminator. Place a UV light visible ruler next to the gel while taking the photograph as it will be useful to approximate the size of the insertions. 4. Transfer the DNA onto a positively charged nylon membrane using standard southern blot protocols. 5. Following the transfer, rinse the membrane in 5X SSC and prehybridize in Ambion RapidHyb supplemented with 0.1 mg/ml herring sperm DNA at 45°C for 1–2 h with agitation. 6. Generate a P32-labeled EGFP probe. We digest the CLEGFP-DI-attB plasmid or CMV-EGFP-DI-attB plasmid with BamHI and AgeI, run the digested DNA on a 1% agarose gel, and then isolate the GFP fragment using the Qiagen Gel Extraction Kit. We then label the EGFP fragment using Rediprime II Random Prime Labeling System following the manufacturer’s instructions. Free nucleotides are removed using the Qiagen nucleotide removal kit. 7. After prehybridizing the membrane in Ambion RapidHyb for 1–2 h, boil the labeled probe for 5 min and then place the probe on ice for 5 min. Add denatured probe to prehybridization solution and hybridize at 45°C overnight. 8. After hybridization, rinse the blot with 2X SSC, 0.1% SDS at room temperature and then wash the blot with two 68°C 100-ml washes of 2X SSC, 0.1% SDS (5 min each), two 100-
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ml washes using 1X SSC, 0.1% SDS (10 min each), and four 100-ml washes using 0.1X SSC, 0.1% SDS (30–60 min each). 9. Sandwich washed blot in plastic wrap and expose to Kodak MS film. We recommend using film cassettes with intensifying screens at 80°C. Develop the first exposure 12 h after placing the film. If the signal is weak, additional exposure time may be needed (48–120 h). 10. Analyze the southern blot for evidence of genomic integration. Southern blot insertion sites have ranged in size from 10 kb to 50 kb in size (Fig. 9.2F).
4. Notes 1. MTA agreements are required from Dr. Gary Felsenfeld (NIH) to use the HS4 insulator sequences and Dr. Daniel Weeks (Iowa) for the CMV-EGFP-DI-attB and CL-EGFP-DI-attB plasmids. An MTA is also required to obtain the plasmid pET11phiC31poly(A) from Dr. Michele Calos (Stanford). 2. The amount of fluorescence produced from a single-copy transgene (fC31 integrase approach) may be significantly less than an embryo containing multiple copies of a transgene generated from the restriction enzyme mediated insertion (1, 2) approach or the meganuclease (3, 4) approach.
Acknowledgements We would like to thank Professor Michele Calos for providing the pET11phiC31poly(A) plasmid, Professor Gary Felsenfeld for providing the HS4 insulator sequences, and Paul Kreig for providing the gamma crystallin lens promoter. This work was supported by funding from the NIH (GM069944 and DC007481). Bryan Allen is a student in the Medical Scientist Training Program at the Roy J. and Lucille A Carver College of Medicine, University of Iowa.
References 1. Kroll K.L., Amaya E. (1996) Transgenic Xenopus embryos from sperm nuclear transplantations reveal FGF signaling requirements during gastrulation. Development 122, 3173–83. 2. Sparrow D.B., Latinkic B., Mohun T.J. (2000) A simplified method of generating
transgenic Xenopus. Nucleic Acids Res. 28, E12. 3. Ogino H., McConnell W.B., Grainger R.M. (2006) Highly efficient transgenesis in Xenopus tropicalis using I-SceI meganuclease. Mech. Dev. 123, 103–13.
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4. Pan F.C., Chen Y., Loeber J., Henningfeld K., Pieler T. (2006) I-SceI meganucleasemediated transgenesis in Xenopus. Dev. Dyn. 235, 247–52. 5. Liu J., Jeppesen I., Nielsen K., Jensen T.G. (2006) Phi c31 integrase induces chromosomal aberrations in primary human fibroblasts. Gene Ther. 13, 1188–90. 6. Kuhstoss S., Rao R.N. (1991) Analysis of the integration function of the streptomycete bacteriophage phi C31. J Mol Biol. 222, 897–908. 7. Thyagarajan B., Olivares E.C., Hollis R.P., Ginsburg D.S., Calos M.P. (2001) Site-specific genomic integration in mammalian cells mediated by phage phiC31 integrase. Mol. & Cell. Biol. 21, 3926–34. 8. Thorpe H.M., Smith M.C. (1998) In vitro site-specific integration of bacteriophage DNA catalyzed by a recombinase of the resolvase/invertase family. Proc. Nat. Acad. Sci. USA 95, 5505–10. 9. Lutz K.A., Corneille S., Azhagiri A.K., Svab Z., Maliga P. (2004) A novel approach to plastid transformation utilizes the phiC31 phage integrase. Plant J. 37, 906–13. 10. Groth A.C., Olivares E.C., Thyagarajan B., Calos M.P. (2000) A phage integrase directs efficient site-specific integration in human cells. Proc. Nat. Acad. Sci. USA 97, 5995–6000. 11. Chalberg T.W., Genise H.L., Vollrath D., Calos M.P. (2005) phiC31 integrase confers genomic integration and long-term transgene expression in rat retina. Invest. Ophthalmol. Vis. Sci. 46, 2140–6. 12. Thomason L.C., Calendar R., Ow D.W. (2001) Gene insertion and replacement in Schizosaccharomyces pombe mediated by the Streptomyces bacteriophage phiC31 site-specific recombination system. Mol. Gen. & Genomics: MGG. 265, 1031–8. 13. Olivares E.C., Hollis R.P., Chalberg T.W., Meuse L., Kay M.A., Calos M.P. (2002) Site-specific genomic integration produces therapeutic Factor IX levels in mice. Nat. Biotechnol. 20, 1124–8. 14. Held P.K., Olivares E.C., Aguilar C.P., Finegold M., Calos M.P., Grompe M. (2005) In vivo correction of murine hereditary
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tyrosinemia type I by phiC31 integrasemediated gene delivery. Mol. Ther. 11, 399–408. Belteki G., Gertsenstein M., Ow D.W., Nagy A. (2003) Site-specific cassette exchange and germline transmission with mouse ES cells expressing phiC31 integrase. Nat. Biotechnol. 21, 321–4. Keravala A., Portlock J.L., Nash J.A., Vitrant D.G., Robbins P.D., Calos M.P. (2006) PhiC31 integrase mediates integration in cultured synovial cells and enhances gene expression in rabbit joints. J. Gene Med. 8, 1008–17. Ishikawa Y., Tanaka N., Murakami K., Uchiyama T., Kumaki S., Tsuchiya S., Kugoh H., Oshimura M., Calos M.P., Sugamura K. (2006) Phage phiC31 integrasemediated genomic integration of the common cytokine receptor gamma chain in human T-cell lines. J. Gene Med. 8, 646–53. Bertoni C., Jarrahian S., Wheeler T.M., Li Y., Olivares E.C., Calos M.P., Rando T.A. (2006) Enhancement of plasmid-mediated gene therapy for muscular dystrophy by directed plasmid integration. Proc. Natl. Acad. Sci. USA 103, 419–24. Groth A.C., Fish M., Nusse R., Calos M.P. (2004) Construction of transgenic Drosophila by using the site-specific integrase from phage phiC31. Genetics 166, 1775–82. Allen B.G., Weeks D.L. (2005) Transgenic Xenopus laevis embryos can be generated using phiC31 integrase. Nat. Methods 2, 897–8. Kuhn E.J., Geyer P.K. (2003) Genomic insulators: connecting properties to mechanism. Curr. Opin. Cell Biol. 15, 259–65. West A.G., Gaszner M., Felsenfeld G. (2002) Insulators: many functions, many mechanisms. Genes Dev. 16, 271–88. Hollis R.P., Stoll S.M., Sclimenti C.R., Lin J., Chen-Tsai Y., Calos M.P. (2003) Phage integrases for the construction and manipulation of transgenic mammals. Reprod. Biol. Endocrinol. 1, 79. Sive H.L., Grainger R.M., Harland R.M. (2000) Early Development of Xenopus laevi: A Laboratory Manual. Cold Spring Harbor Laboratory Press. Cold Spring Harbor, N.Y.
Chapter 10 Germline Transformation of Caenorhabditis elegans by Injection Pavan Kadandale, Indrani Chatterjee and Andrew Singson Abstract Microinjection is a commonly used technique for DNA transformation in Caenorhabditis elegans. It is a powerful tool that links genetic and molecular analysis to phenotypic analysis. In this chapter we shall provide an overview of microinjection for germline transformation in worms. Our discussion will emphasize C. elegans reproductive biology, applications and protocols for carrying out microinjection in order to successfully obtain transgenic worms. Key words: Microinjection, C. elegans, Caenorhabditis elegans, transformation, reproductive biology, transgenics.
1. Introduction 1.1. Reproduction in Caenorhabditis elegans
The Caenorhabditis elegans hermaphrodite reproductive tract shows bilobed symmetry. Each lobe or gonad arm (one anterior and one posterior) is U-shaped and consists of a distal ovary, proximal oviduct, spermatheca, and uterus (Fig. 10.1). The distal region (with respect to the uterus) of each gonad arm is a syncitium of nuclei that are eventually surrounded by membranes, and the cells thus formed give rise to the germline. Oocytes in the oviduct mature in an assembly-line fashion and enter the spermatheca, the site of fertilization in C. elegans. Oocytes can be fertilized by hermaphrodite-derived sperm or male-derived sperm introduced by mating. Upon fertilization, partial embryonic development occurs in the uterus, and eggs are laid before hatching. In C. elegans, genetic transformation is achieved by microinjection of DNA into the syncitial gonad (1,2). This
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Fig. 10.1. A schematic overview of C. elegans transformation by injection and the hermaphrodite reproductive tract. Plasmids containing marker and target DNA are injected into the syncytial gonad arm. Injected DNA then forms an extrachromosomal array or minichromosome. Progeny that inherit the array display the marker phenotype.
ensures that the injected DNA gets incorporated into the nuclei of oocytes, and can eventually be inherited via the fertilized egg. 1.2. Microinjection, Method and Outcome
Kimble et al. (3) first employed transformation via microinjection in C. elegans where sup-7(ts5) (amber suppressor mutant) tRNA was injected into the gonad of mutant hermaphrodites. Following this, several key studies standardized injection of foreign nucleic acid sequences into the worm gonad (4–6). DNA microinjection is typically accompanied by coinjection of an easily scorable marker (Fig. 10.1). The choice of marker often depends on the background of the worm strain. Usually a marker whose expression will not be interfered with is chosen. One such marker used is rol-6, a dominant marker encoding a cuticle collagen. The mutant form causes animals to roll when they move and hence animals
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that are transformed can be identified easily under the dissecting scope. Another example is a strongly expressed green fluorescent protein (GFP) marker, such as myo-3::GFP (Fire lab vector kit) (2) which is expressed in muscle tissue. Although this requires the use of a fluorescence microscope it can be used reliably for detecting the presence of transgenes by detection of GFP in body wall muscles. Injected exogenous DNA typically assembles through nonhomologous recombination to form large concatemers or extrachromosomal arrays with numerous copies of the injected DNA (Fig. 10.1) (5). The extrachromosomal arrays are heritable and show non-Mendelian segregation patterns. The transmission rate is positively influenced by array size (5) and can also be improved by integration into the genome (see below). In the germline, the repetitive structure of extrachromosomal arrays can cause them to be silenced (7). This problem can often be overcome by increasing the complexity of the injection mixture by including C. elegans or yeast genomic DNA so that tandem repeats of the extrachromosomal DNA are not formed. Injected DNA can also occasionally insert into the genome through homologous and nonhomologous integration (2). Irradiation/ mutagenesis that induces chromosomal breaks in animals carrying extrachromosomal arrays (1) or microparticle bombardment techniques can be used when integration is required (8). 1.3. Applications for Microinjection
In C. elegans, transformation using microinjection has been used frequently for cloning by mutant rescue. A commonly used procedure for sterile mutants is to introduce the exogenous DNA into the wild-type background and then cross the transgenic worms to the mutant strains. Rescue is scored as recovery of fertility in the mutant strain due to the presence of a transgene. This method can also be used for analysis of gene expression through reporter gene assays that include using markers like GFP, lacZ, or tissue-specific, ectopic and overexpression assays. Reporter genes introduced via microinjection can be used not only for the analysis of temporal and spatial expression patterns of genes but also to ascertain the conditions required for various expression patterns. Microinjection can also be used for structure–function analysis, expression of tagged proteins, and RNA interference. Once transgenic lines are established, often there is a random loss of extrachromosomal DNA during mitotic division so that animals are mosaic for a given transgene. Although this may be a problem, sometimes the analysis of such mosaic animals can be used to determine the site of action for a gene (9).
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2. Materials 1. Worm Pick: Platinum wire, Pasteur pipette. 2. Injection pads: 2% agarose, glass coverslip (24 40 mm). 3. Injection needles: borosilicate capillary tubing is commonly used (1.0 mm o.d. 0.75 mm i.d./Fiber). 4. Needle puller: Sutter Instruments, Model P-97 micropipette puller for injection needles. 5. Microscope for injection: An inverted compound microscope such as the Zeiss Axiovert equipped for Nomarski DIC microscopy or equivalent along with a micromanipulator for the needle. 6. Halocarbon oil 700 for covering worms during injection. 7. Young adult hermaphrodites (see protocol). 8. DNA for the injection mix. 9. A nitrogen tank connected to the injection rig with valves for controlling needle pressure for injection. 10. Seeded NGM plates, dissecting microscope, and 1X M-9 buffer for recovery (22-mM KH2PO4, 22-mM Na2HPO4, 85-mM NaCl, 1-mM MgSO4).
3. Methods 3.1. Growing and Cultivating Worms
1. Standard techniques and recipes for growing worms and cultivating them are used. In short, worms are grown on NGM plates seeded with the OP50 strain of bacteria (10).
3.2. Make a Worm Pick
1. Use thin-gauge platinum wire to make a worm pick. The platinum can be flamed between uses to sterilize it, and cools rapidly. 2. Break off the long end of a Pasteur pipette to make the pick ‘‘handle’’. The length of the handle depends upon individual preference. 3. Place one end of the platinum wire into the pipette and then flame this till the glass seals over the pore of the pipette, fusing the platinum wire in. Make sure to keep the pipette vertical over the flame, so that you get a straight pick, instead of a bent one. 4. Trim the platinum wire to the desired length (again, the length depends on personal choice).
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5. Using a coin, flatten the free end of the pick so that it is pretty flexible. 6. Trim the flattened end of the pick using a blade so that it is quite narrow. The trimming is best carried out under a dissecting microscope. 7. The pick is now ready for use. 3.3. Prepare Pads
1. Make a solution of 2% agarose in water. 2. Using a Pasteur pipette, place two separate drops of the 2% agarose (see Fig. 10.2) onto a glass coverslip (24 40 mm coverslips are best). Repeat for as many coverslips as the prepared solution will make. 3. Allow the agarose solution to solidify. Do not stack the coverslips. 4. Trace a circle around the agarose drop on the back surface (the surface behind the side upon which the agarose drop was placed) of the coverslip using a thin-tipped marker, and then divide this circle into quarters (see Fig. 10.2). Repeat for all the coverslips. 5. Mark the front surface of the coverslip with a letter so that the front surface can be distinguished from the back. 6. Allow the coverslips to dry for a day. 7. Store the coverslips in a sealed container (so that they do not re-moisturize) on a sheet of filter paper.
3.4. Get Worms Ready
1. The ideal worms to inject are young adults that have already completed their first few ovulations. 2. Pick hermaphrodites at the L4 larval stage (which show a typical clear zone around the developing vulva – see Fig. 10.3) and allow them to grow overnight at 20°C. 3. These worms are ready to inject the next day.
Fig. 10.2. Injection pads. The injection pad consists of two drops of dried agarose. The letter on the pad (‘‘P’’) helps to indicate which side of the pad is up, since it can be hard to visualize the dried out agarose drops.
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Fig. 10.3. Picking the right worms for injections. (A) Hermaphrodites should be picked as L4 larvae and allowed to age overnight. The arrowhead indicates the vulva clearing that characterizes L4 hermaphrodites. (B) The overnight aged worms are young adults that can be used for injections.
3.5. Prepare Injection Mix
1. Make sure that the total concentration of all the DNA is greater than 100 mg/ml. Typically, the injection mix consists of 100 mg/ml of a marker (such as a muscle-expressed GFP, or a dominant mutation that causes the worms to roll) and 10 mg/ml of the DNA of interest. 2. Centrifuge the injection mix for 10 min at 14,000 rpm. Carefully pipette 2–3 ml from the top into a fresh eppendorf tube and use this for injections. Any unused DNA can be pooled back for later use. This step ensures that any fine particulate matter – which can clog the injection needle – that may be present in the injection mix is eliminated from the sample that is used to load the needle.
3.6. Clean Worms
1. The hermaphrodites that are going to be used for injections must be cleansed of bacteria that can clog the injection needle. 2. To do this, place the young adults from the ‘‘Get worms ready’’ step onto a NGM plate that has not been seeded with bacteria. 3. As the worms crawl around on this plate, they will lose most of the bacteria that might clog the needle. 4. Keep in mind, though, that these worms are now being starved, so put the worms on these unseeded NGM plates in batches of 8–10, as they are used up for injections.
3.7. Load Needle
1. There are three alternative methods to load the injection needle. A. Invert the needle and place it into the DNA solution, so that the DNA is pulled up into the needle by capillary action. This process usually takes a few minutes. This method is not ideal for larger-bore (0.75-mm inner diameter) needles.
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B. Heat the middle of the needle for 1–2 s over a flame and then place a drop of the injection mix on the non-tip end of the needle. As the heated air in the needle cools and contracts, the drop on injection mix will get sucked into the needle. C. Pull a Pasteur pipette and break off the thin, pulled part in the middle. Using appropriate tubing and adapters, connect the non-pulled end of this pulled pipette to a 1-ml syringe (Fig. 10.4). Pull the plunger of the syringe back about half-way and then dip the tip of the pulled pipette into the injection mix. Some of the injection mix will be pulled into the pipette tip by capillary action. Once a sufficient quantity of liquid has entered the pulled pipette tip, remove it from the injection mix. Carefully thread the pipette tip into the injection needle and depress the plunger to squeeze the injection mix into the needle. Pull the pipette tip out of the injection needle. This method is a little more complicated than the other two, but yields very good results.
Fig. 10.4. Loading the injection needle. (A) A syringe (grey arrowhead) is attached to the end of a pulled Pasteur pipette tip (white arrowheads – see text) using an appropriately sized adaptor (white arrow). (B) The injection solution is drawn into the pulled Pasteur pipette tip, which is then inserted into the injection needle bore. The injection solution can now be pushed into the injection needle.
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2. Once the needle has been loaded, place it into the needle holder of the injection apparatus on the microscope. 3. Adjust the needle so that it makes as flat an angle as possible with the microscope stage. 3.8. Break Needle
1. Break off a small part of the pulled pipette tip and place it on a 24 40 mm glass coverslip. 2. Overlay the pipette tip with a drop of Halocarbon 700 oil. 3. Now, focus on the edge of the pipette tip and then adjust the needle so that its tip is in the same focal plane as the edge of the pipette. 4. At this stage, ensure that the tip is intact (there should very little, or no flow of liquid when the nitrogen pump is turned on and gas is forced into the needle). 5. Very gently, move the pipette tip so that it just taps the needle. Check the flow of liquid by turning on the pump. 6. Repeat the tapping of the pipette tip onto the injection needle till it has been broken sufficiently to allow a good flow rate. 7. However, if the opening of the needle at this point is too large, it cannot be used for injections, and a new needle must be loaded and broken. 8. Once broken, prevent the injection mix from drying out by always keeping the needle dipped in a drop of oil (Halocarbon oil 700 on a glass coverslip) when it is not being used to inject a worm (see Note 1).
3.9. Lay Worms Down on Pad
1. Place a drop of Halocarbon oil 700 on each of the two agar pads of one coverslip, so that the pad is completely covered by the oil. 2. Now, with the help of a dissecting microscope and the flat worm pick, pick a single worm from the unseeded NGM plate and place it on one of the unused quadrants of the agar pad (see Note 2). 3. Immobilize the worm by gently pushing it down onto the pad with the pick. It is sufficient for only a substantial portion of the worm to be stuck to the pad. Often, you will see about ½–3/4 of the worm immobilized on the pad, but the remainder will be moving. As long as the worm cannot completely come off the pad, it can be injected. 4. Make a mental note of which quadrant the worm has been placed in. 5. Transfer the coverslip with the worm onto the microscope to be used for injection.
3.10. Inject
1. Make sure that the injection needle is above the worm, to avoid the possibility that it will inadvertently poke or kill the worm.
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Fig. 10.5. Injecting the worm. (A) The needle is aligned with the immobilized worm. (B) The stage is now moved (not the needle) so that the worm is impaled on the needle and the needle enters the gonad. (C) The injection solution is now pushed into the gonad. Correctly done, the injection will result in the gonad swelling up a little bit, much like a long balloon being filled with air. (D) The stage is now moved backwards so that the needle is withdrawn from the worm, allowing the injected animal to be recovered.
2. Orient the worm so that its gonad (the gonad looks like a large sac filled with grapes) is almost parallel to the injection needle (see Fig. 10.5). 3. Lower the injection needle so that it is in the same plane as the gonad. 4. Now, carefully move the worm towards the needle so that the needle finally breaks the cuticle and enters the gonad (see Fig. 10.5). 5. Turn on the pump, so that the DNA solution in the needle is forced into the gonad. You should see the gonad filling up, like a balloon being filled with water. 6. Make sure not to inject too much DNA into the gonad, or too quickly. You do not want to see the DNA ‘‘explode’’ into the gonad, but rather to fill it up slowly. 7. Once you have injected the gonad, gently move the worm off the needle. 8. Raise the needle again and transfer the coverslip back to the dissecting microscope. 3.11. Recover
1. While watching under the dissecting microscope, using a 20-ml pipette, carefully pipette one small drop of M9 onto the immobilized, recently injected worm.
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2. The worm should begin rehydrating and will begin to move again. 3. Use the worm pick to carefully transfer the worm from the injection pad to a seeded NGM worm cultivation plate. 4. Make sure that each injected worm is recovered individually, so that at least three independent lines of transformants can be obtained later. 3.12. Score for Transformants
1. The injected worms, if they successfully recover, will lay eggs that will hatch and be ready to score in a couple of days. 2. Score the progeny of the injected worms for the phenotype of the injection marker (e.g., GFP or rolling worms). 3. Place the transformed worms from each successfully injected worm onto a fresh, seeded NGM plate. 4. The pooled, transformed progeny from each injected animal can potentially generate an independent line of transformants. 5. Keep in mind that not all of the transformed F1 progeny from the injected worm will pass on the transgene to its progeny. Further, even the worms that do transmit the transgene through their germline will do so with differing efficiencies. The process of transformation involves a complex rearrangement of the injected DNA into a single, large, multimer of the various DNA species injected (Fig. 10.1). Transformation efficiency depends on the specific gene being used for transformation and the size of the recombined transgene. We have observed transmission efficiencies ranging from almost 90% to as low as 1–2%. 6. Allow the transformed F1 progeny to self-fertilize and in 3 days, score for stably transmitting lines. 7. For all experimental data, at least three independently derived transformed lines must be examined, to negate any artifacts from the recombination events required to form the transgene.
4. Notes 1. If the tip of the needle is always too large when it is broken sufficiently for a good flow rate, there might be problem with the injection mix. Too concentrated an injection mix will often result in this problem. Try diluting the mix. 2. The agar pads hold the worm in place and start to cause desiccation. A small amount of desiccation allows for the volume of the injection without bursting the worm. However, the injection and recovery described below should be done
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relatively quickly to avoid lethal levels of dehydration. Overdesiccation can also make injection more difficult.
Acknowledgments We would like to thank members of the Singson lab for helpful comments and discussion. Research in the Singson lab has been supported by grants from the National Institutes of Health, the National Science Foundation, the Busch Biomedical Fund, and Johnson and Johnson Discovery Awards.
References 1. Jin Y. (1999). Transformation. In: C. elegans: A Practical Approach. Hope I.A, ed. New York: Oxford University Press, 69–96. 2. Mello C., Fire A. (1995) DNA transformation. Meth. Cell Biol. 48, 451–82. 3. Kimble J., Hodgkin J., Smith T., Smith J. (1982) Suppression of an amber mutation by microinjection of suppressor tRNA in C. elegans. Nature 299, 456–8. 4. Stinchcomb D.T., Shaw J.E., Carr S.H., Hirsh D. (1985) Extrachromosomal DNA transformation of Caenorhabditis elegans. Mol. Cell Biol. 5, 3484–96. 5. Mello C.C., Kramer J.M., Stinchcomb D., Ambros V. (1991) Efficient gene transfer in C. elegans: extrachromosomal maintenance and integration of transforming sequences. EMBO J. 10, 3959–70.
6. Fire A. (1986). Integrative transformation of Caenorhabditis elegans. EMBO J. 5(10), 2673–80. 7. Kelly W.G., Xu S., Montgomery M.K., Fire A. (1997) Distinct requirements for somatic and germline expression of a generally expressed Caenorhabditis elegans gene. Genetics. 146, 227–38. 8. Praitis V., Casey E., Collar D., Austin J. (2001) Creation of low-copy integrated transgenic lines in Caenorhabditis elegans. Genetics. 157(3), 1217–26. 9. Yochem J., Herman R.K. (2003) Investigating C. elegans development through mosaic analysis. Development. 130(20), 4761–8. 10. Brenner S. (1974) The genetics of Caenorhabditis elegans. Genetics. 77, 71–94.
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Chapter 11 Quantitative Microinjection of Mouse Oocytes and Eggs Douglas Kline Abstract Quantitative microinjection is used to introduce known quantities of molecules or probes into single cells to examine cellular function. The relatively large mammalian oocyte or egg is easily manipulated and can be injected with impermeant reagents including a variety of signaling molecules and fluorescent probes. Techniques have been developed to inject picoliter quantities of solution into oocytes and eggs with precision and reliability. The methods described here outline the quantitative injection procedures as they are used to inject mouse oocytes and eggs in a culture dish on the stage on an inverted microscope. The techniques are applicable to the oocytes, eggs, and early embryos of most mammalian species. Included are some general instructions on fabrication of transfer pipettes, holding pipettes, beveled injection pipettes, and equipment for quantitative injection. Key words: Microinjection, egg, oocyte, mouse, pipette, beveler, microforge.
1. Introduction The mouse oocyte is arrested at prophase I of meiosis and the cell contains a large nucleus (germinal vesicle). Following hormonal stimulation, the oocyte resumes meiosis and matures to form the egg or ovum, which can be fertilized and produce an embryo (Fig. 11.1A). A number of studies have utilized microinjection to examine cellular processes in these cells. Microinjection of oocytes has been used to examine the processes that regulate prophase arrest and meiosis (e.g., ref. (1). Our understanding of fertilization has relied on the injection of calcium indicators and other factors to determine the mechanism by which the sperm activates the egg to begin development (e.g., refs. (2,3). Microinjection of exogenous DNA into the pronucleus of the early embryo is used in the production of transgenic mice (for David J. Carroll (ed.), Microinjection: Methods and Applications, Vol. 518 Ó 2009 Humana Press, a part of Springer ScienceþBusiness Media, LLC DOI 10.1007/978-1-59745-202-1_11
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Fig. 11.1. (A) Mammalian oocyte maturation and fertilization (GV, germinal vesicle; PB, polar body; PN, pronuclei). (B) Transfer pipette and aspirator assembly. Shown here, in a plastic container, are the mouthpiece and an in-line filter. (C) Photographs of a holding pipette and an injection pipette. Left: Holding pipette after fire polishing. The inner diameter is approximately 10 mm. Right: The injection pipette at the same magnification. Inset: The injection pipette at higher magnification. The tip is beveled and has an outer diameter of approximately 2 mm. (D) The microinjection system for quantitative microinjection consists of a micropipette inserted into the micropipette holder, which is connected by tubing to a screw-controlled syringe. (E) Microforge. A microscope is configured with a heating element to forge micropipettes. Current is regulated by a DC power supply (right). In this case, the microscope is positioned on its back and the pipette is lowered vertically. (F) Microforge close-up view. The pipette is lowered vertically down toward the filament using a small manipulator.
methods see ref. (4) and injection of protein inhibitors or activators may be used to examine a variety of signaling or metabolic pathways. For some of experiments (such as the injection of DNA), knowing the approximate amount of material injected is sufficient; however, it is preferable to use a quantitative microinjection method whenever possible so that a reasonable statement can be made about the intracellular concentration of injected materials, thus ensuring that the experiments provide physiologically relevant data. Injection volumes range from 1 pl to 20 pl. It is possible to rather precisely and reliably inject such small volumes using a
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quantitative injection method that relies on the injection of solutions through a micropipette by the application of hydraulic pressure. A micropipette is attached to a fluid-filled tubing system connected to a screw-controlled syringe. One of two methods is employed to permit fine control on the fluid in the tip of the micropipette as pressure is applied to the fluid in the tubing. The first method utilizes a small drop of mercury that is backfilled into the micropipette tip. The mercury drop serves to regulate flow at the tip of the pipette (5–7). The second method uses a micropipette that has a constriction near the tip that modulates the flow at the tip (8–10). Calibration of the injection volume for both methods is achieved by injecting and measuring the volume of an oil drop that is equivalent to the volume of injected solution. The following methods describe the quantitative injection procedures as they are used to inject mouse oocytes and eggs in a culture dish on the stage of an inverted microscope. Oocytes and eggs are collected and manipulated with small transfer pipettes. The cells are held in position in culture dishes with holding pipettes and injected using beveled injection pipettes. The injection pipette and injection apparatus permit fine control of the injection of picoliter amounts of injection solution.
2. Materials 2.1. Supplies for Oocyte or Egg Collection
1. Mice (3–10 weeks old) of specific strains can be obtained from a number of suppliers (e.g., The Jackson Laboratory, Harlan Sprague Dawley, Charles River Laboratories). 2. Equine chorionic gonadotropin (eCG) also known as pregnant mare’s serum gonadotropin (PMSG) (e.g., Sigma #G4877). Store powder at –20°C and reconstituted solution aliquots (200 IU/ml) at 20°C or refrigerated for several days. This hormone may also be obtained from the National Hormone and Peptide Program (http://www.humc.edu/hormones). 3. Human chorionic gonadotropin (hCG; e.g., Sigma, # C1063). Store powder at 2–8°C and reconstituted solution aliquots (200 IU/ml) at 20°C or refrigerated for several days. 4. CO2 tank and low flow regulator for euthanizing animals. 5. Surgical instruments including fine scissors and forceps (such as Dumont #5, Fine Science Tools). 6. Tuberculin syringe and 26-gauge needle for oocyte and egg collection. 7. Petri dishes (35mm, e.g., Falcon #1008).
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8. Cell collection medium such as MEM (Sigma, #M0643). Media can be made in smaller volumes from powder. Supplement with 20-mM HEPES (pH 7.4), 75 mg/ml penicillin G, 50 mg/ml streptomycin sulfate, 0.1% PVA, or 0.4% BSA. Filter and store at 4°C. 9. HEPES (Sigma #H6147), store at 20°C. 10. Penicillin G (Sigma #P4687), store at room temperature. 11. Streptomycin sulfate (Sigma #S1277), store at 4°C. 12. Poly(vinyl alcohol) average Mw 9000–10,000 (Sigma-Aldrich, #360627), store at room temperature. 13. Bovine serum albumin (BSA) (e.g., Cohn Fraction V Powder; Equitech-Bio #BAC62). 14. Dibutyryl cyclic-AMP (Sigma #D0627) for maintaining oocytes in meiotic arrest. Store at 20°C and add to the media the day of use at 0.1 mg/ml. 15. Hyaluronidase (type IV-S; Sigma #H3884) prepare as 10 mg/ml stock and store at 20°C. 16. 0.22-mm syringe filters or other filter units. 17. Cell culture tested light mineral oil (Sigma #M8410) filtered through a 0.8-mm filter (Nalgene #127-0080) or prefiltered oil (Sigma #M5310). 18. Stereo microscope preferably with 20x eyepieces, transmitted light base as well as fiber optic illumination. 19. Warm tray for holding culture dishes near 37°C (e.g., Fisher slide warmer #12-594Q). Surface thermometer (e.g., Fisher #15-170-10A) and temperature probe (e.g., BAT 12 with IT-18 probe from Physitemp Instruments). 20. Cell culture incubator for long-term culture (37°C). If the culture medium contains a bicarbonate buffer, incubate in an atmosphere of 5% CO2, 95% air, or 5% CO2, 5% O2, 90% N2. 2.2. Oocyte and Egg Transfer Pipettes
1. Borosilicate glass pipettes (100 ml, Drummond Scientific, #2-000-100). 2. SigmacoteTM (Sigma #SL2). Sigmacote is a chlorinated organopolysiloxane in heptane; it is flammable and corrosive. Use it in the hood and take precautions. Store at 2–8°C. 3. Aspirator assembly (Sigma-Aldrich, #A5177 or Drummond Scientific #2-000-000). 4. Diamond pencil (Thomas Scientific, glass marker or Wale Apparatus Company, diamond scribe).
2.3. Microinjection Platform
1. Inverted compound microscope which may be equipped with epifluorescence, differential interference contrast (Nomarksi)
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or modulation contrast (Hoffman), and 10x–40x objective lenses. A long working distance condenser is preferred. 2. The microscope must be equipped with an ocular micrometer reticle. A reticle with a 10-mm line divided into 100 parts works well. Consult the microscope dealer to find the correct diameter of the reticle for the eyepiece. Use a stage micrometer to calibrate the ocular reticle. 3. Glass-bottom petri dishes may be purchased (e.g., Bioptechs) or made in the lab (see 4,5,6 below). 4. Roper Whitney XX Hand Punch (Roper Whitney #135010001, with bench mounting base #139010001) for glass-bottom dishes produced in the lab (see Section 3.3.3). 5. Hand deburring tool (Small Parts Inc., #DBR-HT). 6. Sylgard (World Precision Instruments, #SYLG184) for attaching coverslips to petri dishes. Store the two components at room temperature. Prepare small portions immediately before use in a small weigh boat. 7. A warming stage that can accommodate the injection dish (35-mm petri dish) and that can maintain the cells at 37°C (e.g., Bioptechs, Warner Instruments). If a bicarbonatebuffered medium is used, the warming stage must be equipped with a means to blow a warm gas mixture of 5% CO2/95% air over the surface of the dish (see also ref. (11). 8. The microscope must be placed on vibration isolation surface (e.g., isolation tables from Newport Corporation, Technical Manufacturing Corporation) 9. Micromanipulators (e.g., Narishige International USA, Warner Instruments, Leica Microsystems, World Precision Instruments, Applied Scientific Instrumentation (see Section 3.3.2). 2.4. Microinjection System and Supplies
1. Micropipette puller (e.g., Narishige, Sutter Instruments, Kopf, Warner Instruments, and Harvard Apparatus). 2. Microforge (e.g., Sutter instruments, World Precision Instruments, Harvard Apparatus, Micro Data Instruments, Warner Instruments; see Section 3.4.4). 3. Prepared holding pipettes (if desired; e.g., Conception Technologies, Humagen Fertility Diagnostics/Medicult). 4. Micropipette beveler (e.g., Narishige International USA, Sutter Instruments, World Precision Instruments; see Section 3.4.3). 5. Turntable beveler: Record player with tonearm, phono cartridge, and stylus (needle). 3 M 0.3-mm aluminum oxide lapping film (Ted Pella, Inc. #815-342). Mini audio amplifier (RadioShack #277-1008), 9 V battery or AC adaptor (RadioShack #273-1767), 24–30 AWG wire (or tonearm wire from
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an audio parts supplier), 1/800 mono phone plug (RadioShack #274-286), soldering tool and solder. 6. Borosilcate glass for holding and injection pipettes. The outer diameter should be 1 mm and the inner diameter 0.5 mm (thick-walled) (e.g., Drummond, World Precision Instruments, Sutter Instruments; a length of 10 cm is convenient to pull two pipettes). 7. Injection system, which includes micrometer syringe, tubing, and pipette holder (e.g., Narishige International USA, #IM5B). The system may also be assembled in parts using a Gilmont 2-ml syringe (Cole-Parmer #K-07845) connected by Teflon tubing (Narishige # CT-1 tubing and CI-1 tube connector) to a pipette holder (e.g., Narishge #HI-7). A second system is required for the holding pipette (see also ref. (7). 8. Fluorinert1 FC-70 (Sigma # F9880), liquid mixture of completely fluorinated aliphatic compounds used in the microinjection system. Store at room temperature. 9. Dimethylpolysiloxane (viscosity 20 centistokes; Sigma DMPS-2X). Store at room temperature. 10. Fixed-needle syringe (e.g., Hamilton 701 N, Hamilton # 80300). 11. Mercury (Sigma-Aldrich #215457). Store at room temperature in a tightly closed container (may be stored under nitrogen gas; see Note 7 for additional information and precautions). 12. Mercury clean-up kits and mercury vapor absorbent wipes using Mercon TM, a mercury vapor adsorbent, are produced by EPS Chemical, Inc. and sold by Fisher Scientific. Adsorbent tray (Fisher # 17-975), wipes (Fisher #17-976-8), and clean-up kits (e.g., Fisher # 17-976-2). 13. Long-tipped plastic pipette for backfilling holding pipettes (e.g., Eppendorf Microloader #930001007) or hand-pulled pipette (see Note 8).
3. Methods 3.1. Oocyte and Egg Collection
Mice of specific strains (3–10 weeks old) are housed following institutional animal care and use procedures. An approved method for euthanizing mice is exposure to 100% CO2. This is best done by placing the animals in a loosely sealed container supplied with flow of CO2 from a regulated gas tank.
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3.1.1. Culture Media
Culture media is best prepared using sterile plastic pipettes and containers. Media should be made only with, endotoxin–free, ultra pure water and quality reagents. Some companies (e.g., Sigma) sell reagents that have been embryo-tested to ensure quality. Media should be sterile-filtered using syringe filter assemblies or larger sterilization units. A HEPES-buffered medium is often used during cell collection and injection to avoid changes in media and cellular pH that would occur with bicarbonate-buffered media exposed to air. A HEPES-buffered medium can be prepared by substituting HEPES at pH 7.4 for bicarbonate in the formulation of most media. HEPES-buffered MEM (see Section 2.1) may be used with oocytes and eggs anytime cells are outside of the incubator. Mammalian oocytes and eggs are traditionally cultured in microdrops under light mineral oil in plastic petri dishes equilibrated to 37°C. Following collection and microinjection the oocytes or eggs can be transferred into a bicarbonate buffered medium made from laboratory reagents for long-term culture. Complete media for egg and embryo culture may also be purchased.
3.1.2. Obtaining Immature Mouse Oocytes
Administer 5–10 IU eCG to female mice by intraperitoneal injection. eCG mimics follicle stimulating hormone (FSH) causing follicles to grow. At 44–48 h later, euthanize the animal and remove the ovaries, rinse and trim away excess tissue in 2-ml media. Transfer each ovary to 2–3 ml media in a 35-mm petri dish on the stage of a stereoscope and repeatedly puncture with a 26-gauge needle to liberate oocytes. To prevent spontaneous maturation of isolated oocytes, the medium should include 0.1 mg/ml dibutyryl cyclic-AMP, which is used to maintain meiotic arrest (see Note 1). Collect only oocytes that have an intact nucleus (germinal vesicle) and are enclosed loosely by cumulus cells. Dissociate the cumulus cells from the oocytes by vigorous pipetting with a transfer pipette (see Section 3.2). Wash oocytes through 3–4 200-ml drops of medium under mineral oil using a transfer pipette. Incubate at 37°C. Oocyte collections and transfer requires a good-quality stereoscope with transmitted light. Oocytes will mature into fertilizable eggs overnight if transferred to a medium without dibutyryl cyclic-AMP.
3.1.3. Obtaining Unfertilized Eggs
Mature mouse eggs can be obtained from most strains following a hormone treatment to induce superovulation. 5–10 IU eCG is introduced by intraperitoneal injection and 48 h later, the mice are injected with 5 IU hCG. Mouse eggs are best collected 12–14 h after hCG injection, since the eggs tend to undergo more spontaneous activation and are more sensitive to artificial activation if collected at later times. Euthanize the animal and remove the ovary and oviducts, rinse in fresh medium, and remove extra tissue and fat with fine
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forceps and scissors. Then, transfer the ovary/oviduct to 2-ml fresh medium. Using reflected (e.g., fiber optic) rather than transmitted illumination for this step, isolate the swollen ampulla of the oviduct to the side of the rest of the tissue, then gently puncture the enlarged portion of the ampulla with a 26-gauge needle and tease the egg-containing cumulus mass with the needle until it oozes out. Using a Pasteur pipette that has been treated with Sigmacote (see Section 3.2) or a standard pipetter and 200-ml pipette tip, transfer the cumulus mass to a 200-ml drop of hyaluronidase solution (0.3 mg/ml in media) under oil for up to 5 min, then transfer individual, cumulus-free eggs through four drops of media (100–200 ml under oil) using a transfer pipette (see Section 3.2). For both oocytes and eggs, if the incubator is not near the microscope, it is convenient to keep the dishes close to 37°C by placing them on a warm tray (see Section 2.1). For further details on superovulation and egg collection, see ref. (12). 3.2. Oocyte and Egg Transfer Pipettes
It is necessary to prepare small-bore glass transfer pipettes to transfer individual oocytes and eggs. Oocytes and eggs are traditionally transferred by mouth pipetting using an aspirator apparatus (see Note 2). Transfer pipettes for mouth pipetting can be prepared from Pasteur pipettes, which are pulled in a Bunsen burner, scored, broken to size, and attached to flexible tubing; however, because a micropipette puller is available in a microinjection lab, the following procedure is best. Borosilicate glass pipettes (100 ml; see Section 2.1) fit in the pipette holder of the aspirator assembly described below. To prevent cells from sticking to the glass pipette, the pipettes can be treated with Sigmacote. In the hood, Sigmacote is drawn into the pipette and then expelled. After drying overnight, a thin film of silicone is left on the glass. The glass tubing is pulled into the micropipette puller at settings similar to those used for the other pipettes (see Section 3.4.2). After the glass is pulled to a small tip, the tip can be scored with a diamond pencil and broken off to give an outer diameter of about 100 mm. This produces an inner diameter just larger than the oocyte and egg, so that cells can be collected and transferred with minimal transfer of media. A slightly larger-diameter tip may be used to strip cumulus cells from oocytes. A good-quality diamond pencil is necessary; some are too coarse and do not produce a good score. The diamond pencil should be reserved only for this use. The trick is to lightly score the tip, then rotate the pipette about 90° and finally tap off the tip end using the tip of the diamond pencil (see ref. (11) for an alternative method of fabricating transfer pipettes). The aspirator assembly consists of a mouthpiece, tubing, and pipette holder. A syringe filter unit is inserted into the tubing to
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maintain some sterility inside the aspirator. The mouthpiece end can be kept in a clean plastic container next to the work area. The container can be covered after use and labeled with the name of the operator (see Fig. 11.1B). 3.3. Microinjection Platform
3.3.1. Microscope
3.3.2. Manipulators
Mouse oocytes and eggs are most easily injected in a petri dish in a drop of medium under oil. An inverted microscope with a warming stage is preferable. The inverted microscope allows easy access to the cells in the dish. Contrast enhancement is necessary if nuclei are to be injected. Phase contrast is not generally as useful as differential interference contrast (Nomarksi) or modulation contrast (Hoffman). The microscope must be equipped with an ocular micrometer reticle, which is needed for volume calibrations. For best imaging, and for differential interference contrast microscopy, the bottom of the petri dish should be glass (see Section 3.3.3). Temperature control is important. A warming stage can maintain the cells at 37°C. The temperature in the center of the dish should be monitored with a small temperature probe, since the dish may not be warmed evenly. Cells can be injected in a HEPES-buffered medium. If a bicarbonate-buffered medium is used, the warming stage must be equipped with a means to blow a warm gas mixture of 5% CO2/95% air over the surface of the dish. The microinjection platform must be isolated from environmental vibrations. If not isolated, the injection pipette may vibrate too much and damage the cell during injection (see Note 3). In most cases, a commercial vibration isolation system is well worth the moderate cost. The micropipettes must be positioned carefully during injection. Two manipulators are required, one for the holding pipette and one for the injection pipette. Manipulators are used to position the pipette in each horizontal direction (X and Y ) and in the vertical direction (Z). The manipulator may be capable of moving the pipette straight forward along its axis; however, this is usually not required. Hydraulic manipulators with remote joystick controls may be used. Other styles include piezoelectric (also with a remote control) or mechanical. The remotely controlled manipulators allow positioning of the micropipette without touching the microscope, manipulator, or vibration isolation table (if the controls are placed off of the table). Mechanical manipulators can be used if they can be attached to the microscope stage or small platforms attached to the microscope or vibration isolation surface. A mechanical manipulator is very stable, though a little more care must be taken because the operator must touch the manipulator to position the pipette and may introduce unwanted vibration.
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3.3.3. Production of Glass-Bottom Dishes
Glass-bottom culture dishes are best for fluorescence microscopy, and glass is required for differential interference microscopy and for lenses with short working distances. Glass-bottom dishes may be purchased from commercial vendors. Alternatively, No. 1½ coverslips can be mounted on petri dishes in which a hole is punched in the dish using a hand punch. A suitable punch is manufactured by Roper Whitney (XX Hand Punch with the 17/3200 round punch and die). After punching, a hand deburring tool is used to remove any rough edges from hole so that the coverslip can be glued to a flat surface without burrs. The best way to attach the coverslip is with Sylgard, a two-part silicone elastomer which, after curing, withstands cold, heat, and moisture and is nontoxic to cells. A small ring of Sylgard is applied to the bottom of the dish around the hole and then the coverslip is lightly pressed on. Curing takes several days and can be done in a dry oven (37–40° C). After the Sylgard has cured, the dishes can be rinsed with purified water or washed with a suitable cell-safe detergent, rinsed, dried, and used immediately (see Note 4).
3.4. Microinjection System and Equipment
Two micropipettes are needed for injection of oocytes and eggs. One pipette holds the cell in position, while the second is used for the injection. Both are made from glass tubing using a micropipette puller. To prepare long tips, a wide heating coil or filament is best (6 mm). Standard heating elements are usually 2–3 mm. Wider filaments should be available from the puller vendors.
3.4.1. Micropipette Puller 3.4.2. Injection Pipettes
3.4.3. Beveling the Injection Pipette
Borosilcate glass tubing is usually used because it is hard, but easy to pull. The outer diameter (o.d.) should be 1 mm and the inner diameter (i.d.) may range from 0.5 mm (thick-walled) to 0.8 mm (thin-walled). The thick-walled glass, which is easier to bevel and less likely to break, is well suited for the quantitative injection method, although thin-walled glass can also be used. The length of the tip from the shoulder to the tip should be about 1 cm, which gives a shank near the tip that is long and small in diameter. This type of pipette minimizes the potential damage resulting from inserting the pipette deep into the interior of the cell. It is best to bevel the tip to increase the cross-sectional area of the tip (see the following section; Fig. 11.1C). See reference (13) for the history, theory, and practice of fabricating micropipettes. Beveling increases the cross-sectional surface area of the opening at the tip, making it possible to front-load the injection pipette with viscous solutions. Beveling also assists in cell penetration. See reference (13) on techniques for beveling. Commercial bevelers consist of a small turntable coated with a fine abrasive. These bevelers are adequate for producing large tips such as for sperm injection (14) or pronuclear transfer (12); however, it takes extra skill to bevel a very fine 1–2-mm tip because one is not able to see the tip to know when it is touching the turntable.
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A method of dry beveling micropipettes described by Kaila and Voipio (15) is quick and simple. The method uses a phonograph pick-up for acoustic monitoring of the beveling process. The grinding surface consists of a rotating piece of 0.3-mm aluminum oxide lapping film. The micropipette is mounted so that the shank touches the stylus (needle) which is wired to an audio amplifier. The pipette is lowered at an angle (such as 35° or 45°) to a distance of about 2 mm from the surface, rotating at approximately 33 rpm. This process is guided by watching the shadow of the tip as the pipette is lowered. Then the pipette is lowered very slowly while listening for the sound indicating the pipette has touched the grinding surface. A ‘‘whooshing’’ sound occurs as the tip hits the surface. Because there will always be some wobble in the grinding surface, the tip will usually hit once per revolution. Listening to the number of hits determines the beveling time and tip diameter. With some practice, it is easy to make beveled tips to the same diameter of about 1 mm (see Note 5). An old record player is easily modified to bevel tips. We made a beveler from a late 1960s-era turntable (Fig. 11.2) that works quite well. The lapping film is attached to the turntable after removing the rubber pad. The tonearm is removed and the
Fig. 11.2. Acoustic beveler. An old phonograph turntable is modified by placing a circular piece of 3 M lapping film on the surface. The tonearm is removed from the turntable and mounted on a manipulator. The tonearm is rewired to plug into an audio amplifier (lower right). The pipette is clamped into position with a spring clip on a plexiglass plate that has a groove in it which is aligned with the stylus. As the pipette is lowered to the surface, the sound of the pipette hitting the surface is amplified. Inset: Close-up of the beveler surface and tonearm. The pipette is positioned so that the shaft of the pipette touches the stylus. The tip is then lowered to the turntable surface (see Section 3.4.3).
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leads, which leave the back of the tonearm from the phono cartridge, are connected by a convenient length of wire to a 1/8’’ phono plug which plugs into a mini audio amplifier. Most systems have four color wires (white = L, red = R, blue = L-ground, and green = R-ground) matching the pins on the phono cartridge which are usually labeled L, R, LG, and RG. The two live white and red wires (L and R) are attached to the central connection of the phono plug and the blue and green wires (LF and RG) are connected to the outer terminal of the phono plug. In some tonearms the ground wires are combined to give a three-wire arrangement. The connections are not difficult to solder although the tonearm wire is very thin (33 AWG). Wire that is 24–30 AWG is suitable for extending the length of the tonearm wires. Tonearm wire might also be purchased from an audio parts supplier. Once the wiring is complete the tonearm needs to be fitted with a pipette holder that holds the pipette so that the shank of the pulled pipette touches the stylus (Fig. 11.2). A machinist may be of help with this modification. 3.4.4. The Microforge
The microforge is used to produce the holding pipette and can be used in a variety of ways to make and modify pipettes. A microforge is equipped with a pipette holder, heating filament and control, and a microscope oriented to view the forging process. The primary use for the microforge is to make holding pipettes. The forge can also be used to break the pipette, to fire polish it, and to bend the tip at a 30–35° angle (see refs (12,14)). If a microforge is not available, it is relatively easily to assemble one with an old microscope and a DC power supply. A small platinum wire (0.127-mm diam.; Sigma-Aldrich # 267 63) is connected to two leads which are in turn connected to a regulated DC power supply (e.g., Epsco Incorporated model EC-2 light purpose power supply; 1–16 VDC, 0–5 A). The wire is mounted on a small block and placed on the microscope stage. The pipette is positioned with the aide of a manipulator. Modifying the microforge using a wire loop and reconfiguring the setup permits the production of constriction pipettes (see Section 3.5.2).
3.4.5. Holding Pipettes
The holding pipette is made from borosilicate glass (1.0 mm o.d., 0.5 mm i.d.) and pulled in the micropipette puller at the same setting used for preparation of the injection pipette. The tip is then scored with a diamond pencil near the tip and then broken off to give an outer diameter of about 80–100 mm with the diamond pencil (see Section 3.2; Fig. 11.1C). Some researchers use a microforge to break the tip of the holding pipette (see refs (12,14)). Whether the tip is scored and broken off by hand or broken in the microforge, it must be fire-polished in a microforge until the inner diameter is approximately 10–15 mm (Fig. 11.1C). The tip
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of the holding pipette is then bent at an angle of approximately 35° by using a microforge or the flame of a small Bunsen burner (e.g., Fisher Scientific # 04-067 N). The pipette is positioned to the side of the flame and, at the same time, a light pressure is applied with a dissecting probe to bend the tip as it just softens near the flame. A bend near the tip permits the holding pipette to be positioned at an angle with the tip more parallel to the dish bottom providing better support during the injection (see Fig. 11.3). 3.4.6. Injection System
The quantitative injection method can be done very effectively with a simple micrometer syringe (see Section 2.4). The same apparatus is used for the holding pipette. The syringe is connected by plastic tubing to a microinstrument or pipette holder which holds the injection pipette (Section 2.4; Fig. 11.1D). The holder, tubing, and syringe are filled with oil (Fluorinert FC-70).
3.5. Injection Procedures
1. Pull the injection pipette and bevel to an outside diameter of about 1 mm.
3.5.1. Pipette with a Mercury Brake
2. Prepare the injection dish, which contains cells in a 50-ml drop of media. In addition, position one or several 1–2 ml loading drops nearby, but not touching the media drop. These loading drops contain the solution to be injected into the cell (see Note 6). Mineral oil should not be used for an injection dish. Both drops are carefully covered by dimethylpolysiloxane
Fig. 11.3. Loading the injection pipette. (A) The injection dish with loading drop (not to scale). In the center is a drop of media containing oocytes or eggs which will be positioned with the holding pipette. To the side is a small drop of injection solution, which will be used to fill the injection pipette. (B) Detailed view of the loading procedure: (1) Mercury is forced to the tip. (2) Oil is drawn into the pipette. (3) The tip is inserted into the loading drop and the injection solution is drawn into the tip. (4) The tip is placed back in the oil and an equivalent amount of oil is drawn into the tip to form the oil cap (see Section 3.5.1, steps 11–12).
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(viscosity 20 centistokes), which I refer to below as oil. Place the dish on the stage of the microscope, preferably on a warm stage. 3. Expel any air that may be trapped in the pipette holder or tubing of the injection system by turning the syringe until a little Fluorinert drips out. Loosen, but do not remove, the collar on the pipette holder. 4. Backfill the injection pipette with approximately 1-ml mercury using a 10 ml, fixed-needle syringe (see Note 7). The mercury can be placed deep in the pipette close to the tip, but it will not flow into the tip. It is not necessary to backfill the pipette with oil. The air and mercury serve as a pressure transducer, permitting the small displacement of a solution in the tip with relatively large pressure increases behind it. 5. Carefully insert the pipette into the holder and tighten the cap. The pipette is held in place by the cap, a brass collar, and silicon tubing. This may be a snug fit if the silicon tubing in the holder is new. Push in very slowly with firm pressure. Keep the pipette straight in line with the holder. Sometimes a small twist on the holder while inserting helps. Tighten the cap on the holder. Check the tightness with a little pull on the pipette. If the glass breaks, put the tip in a mercury waste container in the hood. It may be necessary to take the holder apart and remove glass from the silicon tubing. Be careful and do not loose the brass collar inside the holder if it is removable. If the pipette goes through the silicon too easily and the pipette seems to slip if pressure is applied by turning the syringe, the silicon tubing should be replaced using a similar-sized piece. 6. A drop of mercury, having a high surface tension, usually takes on a spherical shape, but it can be forced to the tip of the micropipette by applying pressure. With the tip of the pipette point away from the user and everyone else, slowly apply pressure to the system by turning the screw on the micrometer syringe. Be patient here. The mercury will move toward the tip slowly at first. After a few seconds, the mercury will move rather quickly to the tip of the pipette. With good lighting on a white background, it is easy to see the mercury fill the tip. If the mercury does not go to the tip, increase the pressure slightly and wait another 10–60 s. If mercury does not flow to the tip within a minute, it may be that the tip is too small to use. If the pipette shows any sign of slipping under higher pressure and the tip does not fill, it should not be used. 7. Mount the pipette holder containing the injection pipette in the manipulator in a position above the stage of the microscope, but up out of the way.
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8. At low magnification (5x or 10x) find the cells in the media drop and focus on them to find the bottom of the dish. 9. Completely backfill a holding pipette with media using a long-tipped plastic pipette or a standard tip pulled to a finer point (see Note 8). Insert the holding pipette into the pipette holder of the second injection system. Position the holder in the manipulator and lower the pipette down toward the media drop, but not in it. 10. Move the injection dish to the injection drop, position the edge of the drop in the center of the field, and bring the edge into focus. Focus up slightly. Bring the injection pipette down into the oil and near the injection drop (while looking at the stage, not through the oculars). Be careful not to go too fast. When the tip is in the oil, close to the center of the light path, view it through the oculars. Bring the tip down slowly using the coarse controls of the manipulator. Focus up to find the tip. Move the tip around using the manipulator controls if it cannot be seen. A shadow may be visible as it moves. Lower the tip down and refocus as needed. If the tip is first brought into focus above the dish bottom, the operator is far less likely to break the tip. With some practice and familiarity with the equipment, the positioning of pipettes can be done rather quickly, although the novice user generally spends more time locating the pipette tips. 11. If the mercury is not at the tip, increase the pressure on the syringe slightly. Switch to a 20X objective lens and begin loading the pipette (Fig. 11.3). Draw up some oil by turning the syringe to the left to apply a negative pressure behind the mercury drop. Only a small amount of oil is needed to separate the injection solution from the mercury. Expel excess. 12. Position the tip of the pipette in the injection solution so that it is conveniently lined up with a calibration mark in the ocular micrometer reticule. Draw up the injection solution, noting the linear movement with respect to the ocular micrometer reticle. The pipette is entering the dish at an angle, so it will be necessary to focus up to find the meniscus (particularly for large volumes). Position the tip out of the injection solution and into the oil by moving the injection drop to the side of the field of view using the stage controls. Draw up the same linear amount of oil to cap the injection pipette. Because the same linear amount was drawn into the tip, the volume of the injection solution and the oil cap are the same. Patience is needed at this step. In some cases, it may take a minute or two for the oil to move into the tip to form the cap. Usually when it starts to move, it does so rapidly. Observe the tip and be ready to apply some pressure to prevent too much
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oil from going in. A slight tap to the manipulator sometimes helps to draw up the oil for the cap if it does not move in after a minute or so. If the cap does not form after loading a relatively large volume, expel the solution. Draw up a small amount of the solution, an oil cap and then expel the cap and draw up the larger volume and then the cap. Often oil can be drawn in to form a cap after the pipette has been capped once. 13. The pipette can be calibrated at this time by injecting the oil cap into the injection solution or injecting the injection solution into the surrounding oil. Volume is determined by measuring the diameter of either drop using the ocular micrometer and calculating the volume (V = 4/3pr3). Repeat the loading procedure and the injection pipette is ready for injection. After several injections, the operator will have an idea what the corresponding volume is for a linear amount of solution. Then the calibration may be done in the cell rather than at this step (see step 18). A short videotape of the pipetteloading process can be found at http://biology.kent.edu/ FacultyPages/kline. 14. After the known volume of injection solution and oil cap are drawn into the pipette, adjust the pressure slightly to prevent more oil from entering. Keep the tip of the pipette in the field of view and move the stage controls to bring the media drop and cells into the field. Watch the pipette tip. Typically, it is necessary to slightly increase the pressure to prevent media from entering the tip. 15. Select a cell and bring the holding pipette down into the media drop near the cell. A slight negative pressure will draw a cell onto the holding pipette. Position the cell in the center of the field. The holding pipette should be resting on the dish bottom so that both the holding pipette and the dish bottom act as a backstop during the injection. If the cell is held above the bottom of the dish, it will likely roll during the injection and be damaged. 16. Position the tip of the injection pipette near the zona pellucida of the cell. Without jarring the microscope stage, switch to a 32x or 40x objective. It is best to position the tip next to the zona and just slightly above the optical equator of the cell. Unless the manipulator has an axial control, the tip will be inserted using either the vertical or horizontal controls of the manipulator. In fact, the horizontal entry is better. As the tip is moved horizontally into the cell, it will depress the membrane and then ‘‘pop’’ through the membrane and into the cytoplasm. Typically, the pipette has to be inserted quite far into the cell before it penetrates. Once in, it is best to move the tip back, releasing some of the tension on the membrane (see Note 9).
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17. Inject the oil cap and injection solution. Applying pressure to the syringe forces the oil cap out of the tip, then the injection solution, which is usually less viscous, flows out quickly leaving only oil in the pipette (see Note 10). A short videotape of the injection process can be found at http://biology.kent. edu/FacultyPages/kline. 18. Remove the pipette (with care to avoid damaging the cell). The oil drop inside the cell will not affect the cell and serves as a marker indicating that the cell was injected. The oil drop is also used to calibrate the injection volume. Measure the diameter of the oil drop inside the cell. From the diameter of the oil drop, calculate the volume injected, which must be the same volume as the injection solution (see Note 11). The amount of material injected is determined from the concentration and volume of the injection solution. The final concentration in the cell is the amount injection divided by the cell volume (an average of volume of 200 pl for the mouse oocyte, for example). Of course, this assumes an even distribution in the cell, which is not necessarily so. Repeat injections with other cells (see Note 12). It is possible to do double injections with different solutions using the same pipette (see Note 13). Some solutions may require additional pipette treatment (see Note 14). 3.5.2. Pipette with a Constriction Brake
The second quantitative injection method uses a constriction in the pipette near the tip to control the flow of solution. Mercury is not used; however, the same injection equipment is used and most of the procedure is similar (see refs (8–10)). 1. Pull an injection pipette and bevel it to about a 1-mm tip. 2. Using a microforge, place a small constriction in the pipette. The microforge consists of a small loop of platinum wire through which the pipette tip is inserted. Current is applied to heat the wire and produce a constriction in the inner diameter (see Section 3.4.4). The constriction should be made about 200 mm from the tip. The inner diameter of the constriction should be approximately the same as the inner diameter of the tip. The platinum wire can be attached to a block that can be positioned on the microscope stage or it can be mounted on an arm of a mechanical manipulator. The injection pipette must also be mounted on a manipulator. It may be easiest to mount the coil and pipette on separate manipulators so that they can be adjusted and are independent of the microscope stage and focus (see ref. (10)). 3. Prepare the injection dish as described in Section 3.5.1 step 2. 4. Expel any air that may be trapped in the pipette holder or tubing of the injection system by turning the syringe until Fluorinert drips out. It is important for this method that
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there is no air in the syringe, tubing, or micropipette holder. Loosen, but do not remove, the collar on the pipette holder. 5. Carefully insert the pipette into the holder and tighten the cap. With the tip of the pipette pointed away from the user and everyone else, slowly apply pressure to the system by turning the screw on the micrometer syringe. Backfilling the pipette in this manner fills the entire pipette with Fluorinert FC-70. A small amount will flow out of the tip. The pipette is now ready to be front-loaded with oil and the injection solution. 6. Mount the pipette holder in the manipulator and position the pipette tip near the loading drop following the procedures described in Section 3.5.1 step 10. 7. The pipette is first loaded with dimethylpolysiloxane, which should be drawn up beyond the constriction in the pipette. Draw up a known amount of the injection solution and then the same amount of oil as the cap. Calibrate or proceed directly to an injection (Section 3.5.1 steps 12–18). The constriction pipette is not difficult to make; however, it is an extra step which requires a small microforge and manipulators. Some practice is required to form the constriction. In other respects, the two quantitative methods are similar. It may be worthwhile to try both methods to determine which is best suited for a particular laboratory, experiment, and injection conditions.
4. Notes 1. For some studies, particularly those involving the regulation of meiosis and the role of somatic cells in that process, it is preferable to study oocytes within the ovarian follicle. Jaffe and her colleagues (11) have developed a well-designed alternative method for the injection of oocytes within the normal physiological environment of the follicle. 2. This is certainly the only case in which mouth pipetting is permitted in the lab. Other methods may work if mouth pipetting is not possible. For example, one might use a 5–10 ml Wiretrol (glass tube and plunger; Drummond Scientific) to transfer cells. 3. Only the microscope should rest on the vibration isolation surface. For best results, the microscope should not rest on rubber feet. If there is any instability in the microscope base, it can be attached to the table with a glue gun, the glue does not harm the microscope should it need to be removed. If mechanical manipulators are used, they should be firmly
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attached to the microscope stage, a shelf attached to the microscope, or to solid platforms on the table surface. Other equipment (such as remotely controlled manipulators) or cables that might cause vibration when touched should not be placed on the surface, but to the side of the table or perhaps on shelves over the table. 4. The desired dishes can be sterilized by placing the open dishes and accompanying lids in a clean culture hood with a UV lamp for several hours or overnight. While UV-treatment is usually sufficient, it is not effective in preventing contamination from spores and other sterilization methods may be needed. 5. The pipettes may be used immediately after dry beveling. Fine dust from grinding is occasionally left on the pipette tip; this comes off during the loading procedure and is usually never a problem. If desired, the pipettes can also be washed by rinsing several times in purified water, and ethanol. Pipettes are also very effectively cleaned (and thinned or sharpened) by a brief dip in 10–25% hydrofluoric acid, and then rinsed in purified water. Hydrofluoric acid must be handled with care and a special hydrofluoric acid spill clean-up kit must be on hand (normal acid spill clean-up kits cannot be used). We prefer to use the beveling system because it produces a sharp tip with a large opening; however, the tip can be also be broken to size by touching it to a flat surface (see refs (7,11)). 6. Some care must be taken with the injection buffer. Especially for larger-volume injections, the buffer should be similar in composition to the intracellular environment, for example,75-mM KCl, 20-mM HEPES, pH 7.0. As in all experiments the appropriate control injection of the same volume must be included in the experimental design. 7. It is best to use fresh mercury; it should be shiny and bright. Because only small amounts are needed, one good source of mercury is a glass thermometer. The bulb end of the thermometer is scored and broken off in a small, clean beaker. The mercury can then be stored in a 1-ml screw-top vial. The risks associated with mercury use are reduced if proper precautions are taken. All manipulations with mercury, except for the final injection, should be done in the hood over a tray lined with a mercury vapor adsorbent. A clean-up kit and mercury vapor absorbent towels should be on hand in case of a spill (see Section 2.4). 8. A backfilling pipette tip can be produced by heating a 200-ml pipette tip near the flame from a small Bunsen burner. While holding the pipette at each end, the middle portion of the tip is heated at the edge of the flame. The pipette is turned to heat both sides. When the plastic has softened, it is removed from the flame, allowed to cool for a few seconds and then pulled.
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After the pull is started, it best to hold the pipette vertically and let the tip fall to the floor. A very fine tip is formed that can be cut to about 5 cm with a razor blade. For backfilling holding pipettes, it is best to attach the tip to a 1-ml syringe filled with media. The holding pipette should be completely filled with media. This is best done by withdrawing the plastic tip as fluid is introduced into the pipette (true for any backfilling operation). 9. The cell membrane is very elastic and can wrap around the pipette without actually breaking. To avoid this problem, it is best to view the injection. After the oil cap enters the cytoplasm, the cytoplasm should be displaced forward slightly as the injection solution enters. If not, the pipette may not have penetrated the membrane. If this occurs in a batch of cells, it can also be avoided by a slight aspiration of the cytoplasm into the tip before the injection. 10. It is possible to inject the solution from the pipette without directly observing the injection. This is particularly useful when real-time fluorescence measurements need to be made during and just after the injection. After the pipette is inserted into the cell, the recording system can be turned on. The injection can be made ‘‘blindly’’ by applying pressure to the system by turning the screw syringe. With a given pressure the oil cap and injection solution will flow into the cell. The injection solution invariably enters the cell after injection of the oil cap since an aqueous solution flows readily out of the pipette once enough pressure is applied to move the oil cap. A larger pressure is required to introduce the oil behind the injection solution and this is not applied. With some practice, the operator can get a good feel for how much pressure to use. Some care should be taken with the calibration because it is possible that a little extra oil may enter. If the oil drop in the cell is larger than expected for a series of injections, the pipette should be calibrated visually in another cell or in oil, rather than relying on the oil drop in the cell. 11. For large-volume injections it is not always necessary to inject the equivalent volume of oil. A smaller oil drop may be better tolerated by the cell. Calibration of the pipette can be done in two ways. The aqueous solution can be injected into oil and the volume calculated as described in Section 3.5.1 step 13. The same volume may then be capped with a smaller volume of oil that serves not for calibration, but just to prevent the loss of injection solution until the injection and to mark the injected cell. Alternatively, if several cells are injected, one might be injected with the full volume of solution and the equivalent oil drop volume to calibrate the system. Thereafter, the same pipette with the same volume can be used in other cells, with a small oil cap.
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12. One pipette may be used for a number of injections. A pipette can be refilled and used several times. However, after several injections pipettes may become more difficult to fill and the chance of cell damage may increase as the outside of the pipette becomes coated with proteins from the cytoplasm. The pipette should be replaced if loading is difficult or if it damages a cell. Pipettes, pulled using the same settings on a puller have nearly the same shape. If the loading sequence is the same, the calibration is usually very similar from pipette to pipette. 13. To inject two solutions using the same pipette, two loading drops are used. After drawing up the first solution and some oil, the pipette is moved into the second drop, a known amount is drawn in, and then the pipette is capped with oil. The two solutions have oil separating them. The injections can then be injected in sequence, with a variable wait period between the injections. However, the longer the pipette remains in the cell, the more likely the cell might be damaged. 14. Some solutions are very viscous and may stick to the inside or outside of the pipette, making it difficult to load the pipette. Some proteins appear to coat the outside of a pipette, causing poor sealing on insertion and withdrawal of the pipette, thereby damaging the cell. Poor sealing may be overcome by silanizing the pipette after it is pulled (16). Silanes form a monomolecular hydrophobic coating on glass surfaces. Beveled pipettes are placed on a metal rack in a large glass petri dish and baked at 150–200°C for 15 min with the cover slightly ajar. A volume of 40-ml N,N Dimethyltrimethylsilylamine (Sigma-Fluka # 41716) is added and the dish is immediately covered and heated for 20 min. The lid is removed and the pipettes heated for a few more minutes. After cooling they are ready for use. N,N Dimethyltrimethylsilylamine is flammable and toxic and hence should be handled in fumehood.
Acknowledgments I thank Laurinda Jaffe for teaching me some of the fine points in the art of microinjection and I remember Ray Kado for his sage advice.
References 1. Mehlmann, L.M. (2005) Oocyte-specific expression of Gpr3 is required for the maintenance of meiotic arrest in mouse oocytes,Dev. Biol. 288, 397–404.
2. Kline, D., Mehlmann, L., Fox, C., and Terasaki, M. (1992) The cortical endoplasmic reticulum (ER) of the mouse egg: localization of ER clusters in relation to the
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generation of repetitive calcium waves. Dev. Biol. 215, 431–442. Shirakawa, H., Ito, M., Sato, M., Umezawa, Y., and Miyazaki, S. (2006) Measurement of intracellular IP3 during Ca2+ oscillations in mouse eggs with GFP-based FRET probe. Biochem. Biophys. Res. Com. 345, 781–788. Brown, G.A.J. and Corbin, T.J. (2002) Oocyte injection in the mouse. Methods Mol Biol. 180, 39–70. Hiramoto, Y. (1962) Microinjection of the live spermatozoa into sea urchin eggs. Exp. Cell. Res. 77, 416–426. de Fonbrone, P. (1949) Technique de Micromanipulation. Monographies de L’institut Pasteur. Paris. Jaffe, L.A. and Terasaki, M. (2004) Quantitative microinjection of oocytes, eggs, and embryos. Methods Cell. Biol. 74, 219–242. Hiramoto, Y. (1974) A method of microinjection. Exp. Cell. Res. 87, 403–406. Kishimoto, T. (1986) Microinjection and cytoplasmic transfer in starfish oocytes. Methods Cell. Biol.. 27, 379–394. Spector D.L., Goldman, R.D. and Leinwand, L.A. (eds.) (1998) Cells: A Laboratory Manual. Vol. 2, Cold Spring Harbor Laboratory Press, Woodbury, NY, pp. 83.1–83.23.
11. Jaffe, L.A., Norris, R.P., Freudzon, M., Ratzan, W.J. and Mehlmann, M. (2009) Microinjection of follicle-enclosed mouse oocytes. Methods Mol. Biol. 12. Nagy, A., Gertsenstein, M., Vintersten, K., and Behringer, R. (2003) Manipulating the Mouse Embryo: A Laboratory Manual (3rd edition). Cold Spring Harbor Laboratory Press, Woodbury, NY. 13. Brown, K.T. and Flaming, D.G. (1986). Advanced Micropipet Techniques for Cell Physiology. Methods in the Neurosciences, Vol. 9. John Wiley and Sons, Chichester. 14. Joris, H., Nagy, Z., Van de Velde, H., De Vos, A., and Van Steirteghem, A. (1998). Intracytoplasmic sperm injection: laboratory set-up and injection procedure. Hum. Reprod. 13 Suppl. 1, 76–86. 15. Kaila, K., and Voipio, J. (1985). A simple method for dry beveling of micropipettes used in the construction of ion-selective micro-electrodes. J. Physiol. 369, 8p. 16. Voipio, J., Pasternack, M., Macleod, K. (1994) Ion-sensitive microelectrodes, in Microelectrode Techniques. The Plymouth Workshop Handbook (Ogden, D, ed.). The Company of Biologists Limited, Cambridge, pp. 275–316.
Chapter 12 Microinjection of Follicle-Enclosed Mouse Oocytes Laurinda A. Jaffe, Rachael P. Norris, Marina Freudzon, William J. Ratzan, and Lisa M. Mehlmann Abstract The mammalian oocyte develops within a complex of somatic cells known as a follicle, within which signals from the somatic cells regulate the oocyte, and signals from the oocyte regulate the somatic cells. Because isolation of the oocyte from the follicle disrupts these communication pathways, oocyte physiology is best studied within an intact follicle. Here we describe methods for quantitative microinjection of follicleenclosed mouse oocytes, thus allowing the introduction of signaling molecules as well as optical probes into the oocyte within its physiological environment. Key words: Microinjection, follicle-enclosed oocyte, mouse.
1. Introduction Intracellular microinjection allows the study of biochemical events within intact cells, but since most cells are interconnected to form complex tissues, many physiological processes could most ideally be investigated by microinjection of cells within intact tissues. This chapter describes methods for injecting mouse oocytes within the complex of surrounding somatic cells known as the follicle (Fig. 12.1). These methods allow the introduction of molecules into the oocyte specifically, without also applying them to the somatic cells, and without disassembling the structure that is needed for investigating physiological responses. These approaches can be used to introduce antibodies, signaling proteins, dominantnegative proteins, buffers, or toxins that interfere with or mimic regulatory pathways (1–4), as well as optical probes to monitor signaling (3,5–7). Because follicles can be cultured for multiple days, these methods also allow the time needed for protein turnover following injection of short interfering RNAs (8). David J. Carroll (ed.), Microinjection: Methods and Applications, Vol. 518 Ó 2009 Humana Press, a part of Springer ScienceþBusiness Media, LLC DOI 10.1007/978-1-59745-202-1_12
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Fig. 12.1. Developmental stages of mouse ovarian follicles. From ref. (10), # Society for Reproduction and Fertility (2005). Reproduced by permission.
The mouse ovarian follicle forms as a single layer of somatic cells around the oocyte; the somatic cells then proliferate to form multiple layers (Fig. 12.1). Follicles containing an oocyte that is surrounded by 2–3 layers of somatic cells are referred to as ‘‘preantral’’. As the follicle continues to grow, spaces form between the somatic cells, and the spaces fuse to form a single antrum. The antrum separates the 1–3 layers of cumulus cells that directly surround the oocyte from the peripheral layers of mural granulosa cells; the cumulus mass is connected on one side to the mural cells. In response to follicle stimulating hormone (FSH), which stimulates follicle growth, receptors for luteinizing hormone (LH) are synthesized in the mural granulosa cells. Up to this point, the oocyte is arrested in meiotic prophase. Then, in response to a surge of LH from the pituitary, meiosis resumes and progresses to metaphase II, and ovulation occurs. This chapter describes methods for microinjecting follicle-enclosed oocytes at both preantral and antral stages. In our lab, we have used these methods to identify mechanisms by which meiotic arrest is maintained prior to the LH surge, and to investigate mechanisms by which LH stimulates meiotic resumption. Although cAMP in the oocyte had long been recognized as an inhibitor of meiotic progression (see refs (9,10)), it had been uncertain where and how this cAMP is generated. Microinjection of follicle-enclosed oocytes using the methods described here has contributed essential evidence that a constitutively active G-protein coupled receptor (GPR3) activates Gs in the oocyte, leading to cAMP production (1, 2, 4, 6, 7). Studies using these methods have also provided evidence that LH does not cause meiotic resumption by terminating GPR3/Gs signaling (7), or by stimulating Gi signaling (3). The recent development of optical probes for monitoring cAMP dynamics in living cells (11) should,
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in combination with these microinjection techniques, allow further studies of the role of cAMP in regulating meiotic progression within an intact follicle. More generally, these techniques provide a new approach that could be used to investigate many other aspects of the complex bidirectional communication between the oocyte and somatic cells (12,13). The following methods for injection of follicle-enclosed mouse oocytes are based on methods that were previously developed for microinjection of echinoderm oocytes (14–16). The follicle is placed between coverslips that compress it slightly, allowing the oocyte to be clearly visualized with a compound microscope (Fig. 12.2). The microinjection pipette is brought in horizontally, and contains mercury to allow precise control when pressure is applied through a screw-driven syringe. The pipette is front-loaded, and the volume injected is quantified by drawing up an equivalent volume of oil and then measuring the diameter of the expelled oil drop. These general methods have been described in detail in a recent chapter (16), which should be referred to for setting up the equipment, and for the basic procedures for injection. The present chapter explains how these techniques have been adapted for follicle-enclosed mouse oocytes. Videos demonstrating these procedures can be viewed at http://www.sciencedirect.com/science/MiamiMultiMediaURL/ B6WDG-4P8B0X8-6/B6WDG-4P8B0X8-6-C/6766/63fd3555 30400573e073a9091ba406e0/video1.mov (supplementary material for ref. 7), and at http://www.jcb.org/cgi/content/full/jcb. 200506194/DC1 (supplementary material for ref. 6).
Fig. 12.2. Microinjection of follicle-enclosed mouse oocytes. (A) Diagram of the microinjection chamber and micropipette. (B) Photograph of a plastic slide for assembling the injection chamber. (C) Photograph of an antral follicle-enclosed oocyte as it appears in the injection chamber.
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2. Materials 2.1. Supplies for Dissection and Culture of Follicles
1. Mice. Follicles are most easily dissected from ovaries of prepubertal mice, 22–25 days old. We use BL/6 SJL F1 mice (The Jackson Laboratory, #100012). 2. Equine chorionic gonadotropin (pregnant mare serum gonadotropin). From the National Hormone and Peptide Program (http://www.humc.edu/hormones/), or various commercial sources. Store single-use aliquots (50 IU/ml, 500 ml) at –80°C. 3. MEM (Invitrogen, #12000-022). We make this medium from powder, adding 25-mM NaHCO3, 75 mg/ml penicillin G, and 50 mg/ml streptomycin sulfate, and store at 4°C for up to 2 weeks. Add serum and other supplements on day of use. 4. Penicillin G (Sigma, #PEN-K). Store powder at room temperature. 5. Streptomycin sulfate (Sigma, #S-6501). Store powder at 4°C. 6. Fetal bovine serum (Invitrogen, #16000-044, or other suppliers). Store aliquots at –80°C, and then at 4°C for up to 2 weeks. 7. Insulin-transferrin-sodium selenite (Sigma, #I1884), dissolved according to the manufacturer’s instructions to make a 100X stock. Store aliquots at –80°C, and then at 4°C for up to 2 weeks. 8. FSH (ovine), from the National Hormone and Peptide Program. Store single-use aliquots (10 mg/ml, 25 ml) at –80°C. 9. Steriflip sterile disposable vacuum filter units, 0.2-mm pore size, 50-ml volume (Millipore, #SCGP 005 25). 10. Miniforceps (Fine Science Tools, #11200-14). 11. Millicell culture plate inserts (Millipore, #PICMORG50). 12. Petri dishes (Falcon, #1008, 35 mm).
2.2. Equipment for Microinjection
1. Stereoscope with an eyepiece micrometer (for dissection of follicles and assembly of injection chambers). 2. Vibration free table (a table with a steel plate supported by rubber spacers, or an air table). 3. Compound microscope with a 20x objective and focusable eyepieces with a micrometer reticle. 4. Micromanipulator with X, Y, and Z controls. 5. Horizontal micropipette puller. 6. Screw-driven syringe connected to a micropipette holder by a piece of narrow tubing containing fluorocarbon oil (Sigma #F9880, Fluorinert FC-70).
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7. Machined plastic slides for assembling the injection chamber (see Fig. 12.2B). 8. For further details, see ref. (16). 2.3. Equipment for Temperature, Humidity, and CO2 Control
1. Warm air blower (Nevtek air stream incubator, #ASI400) 2. Electric thermometer (Physitemp Instruments, #BAT-12R, with an IT-18 temperature probe) 3. Fritted glass cylinder (Corning, #31770-500C, distributed by VWR Scientific Products) 4. Flowmeter (Gilmont, #GF-8321-2410, distributed by Barnant Company) 5. Translucent silicon rubber sheet (Reiss Manufacturing, Inc., 1/800 thick) 6. Magnetic rubber sheet (Custom-Magnets.com)
2.4. Supplies for Construction of Mouth Pipettes for Follicle Transfer
1. Diamond knife (Fine Science Tools, #10100-00), or ceramic knife (Fine Science Tools, #10025-45) 2. Translucent silicon rubber sheet (Reiss Manufacturing, Inc., 1/1600 thick) 3. Polyethylene tubing (Clay Adams, #PE-60, internal diameter 0.76 mm) 4. Mouth pipette assembly (Sigma, #A5177-5EA), see Note 1. 5. Glass capillaries (Drummond Scientific, #9-000-1061, o.d. ¼ 0.8 mm, i.d. ¼ 0.6 mm) 6. Syringe filter (Fisher, #09-719C, 25 mm, 0.2 mm) 7. Glass capillaries (Drummond Scientific, #2-000-100, 100 ml calibrated pipettes) 8. Sigmacote (Sigma, # SL2)
2.5. Supplies for Construction and Assembly of Injection Chambers
1. Coverslips, 22-mm square, #1.5. These should be cleaned before use (see ref. (16)). 2. Diamond pencil. This is an ordinary diamond-tipped glass marker, not the ‘‘diamond knife’’ referred to in Section 2.4.1. 3. Clear flexible plastic ruler, to be used as a guide for cutting coverslips. 4. Black Plexiglas work surface (from a machinist). 5. Double-sided tape (Scotch #137, previously called ‘‘doublestick’’ tape, office supply store). 6. Double-coated tape (Scotch #667, office supply store). 7. Small sharp scissors (Fine Science Tools, #14370-22, Moria 10.5 cm straight).
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8. Silicone grease (Dow Corning high-vacuum grease). 9. Silicon oil (Sigma, #DMPS-2X, dimethylpolysiloxane, 20 centistokes viscosity). 10. Valap (a 1:1:1 mixture of Vaseline, lanolin, and beeswax). 2.6. Supplies for Fabrication of Micropipettes
1. Glass capillaries (Drummond Scientific #1-000-0500, 50 ml ‘‘microcaps’’) 2. Mercury (Sigma-Aldrich, #215457) 3. 10-ml syringe (Hamilton, #701 N)
3. Methods 3.1. Dissection and Culture of Follicles
1. The procedures we use for follicle isolation and culture are based on those used in the lab of John Eppig (see ref. (17)). B6SJLF1 mice 22–25-day old are either primed with 3–5 IU equine chorionic gonadotropin, 40–46 h prior to use, or are unprimed, depending on the particular experiment. 2. The culture medium used for follicle culture is MEM (with NaHCO3 and antibiotics indicated above), supplemented with 5–10% fetal bovine serum. For experiments that require extended culture periods (overnight or longer), the medium is also supplemented with 50 mg/ml insulin, 5 mg/ml transferrin, and 5 ng/ml selenium (from the 100X stock referred to in Section 2.1.7), and 10 ng/ml ovine FSH (from the 1000X stock referred to in Section 2.1.8), and is filtered through a 0.2-mm filter. All media are equilibrated with 5% CO2 and 95% air. 3. To dissect follicles, the ovaries are removed and the bursa broken by grasping the ovary on either side using watchmaker’s forceps, one grasping the fat pad adjacent to the ovary, and the other the oviduct. The ovary pops out of the bursa as the forceps are gently pulled apart. Excess fat is trimmed from the ovaries, and the ovaries are placed into MEM with supplements (2–3 ml in a 35-mm plastic petri dish). Ovaries from primed animals are examined prior to follicle isolation to select only those that are well-primed (see Note 2). 4. Follicles are dissected from ovaries using a stereomicroscope and illuminated using transmitted light. The tools used are miniforceps and/or 30-gauge syringe needles. The combination used depends on personal preference; for example, some people prefer to use one forceps to hold the ovary while cutting the follicles using a needle, while others prefer to use two sets of forceps or two sets of needles. We first slice the ovary in half
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cross-wise; this yields two concave halves. A half-ovary is rotated such that the rounded part is at the top; many follicles, particularly large antral follicles, can be observed when the ovary half is in this orientation. By pushing down slightly on this top (trying to avoid popping follicles of the desired size), the ovary flattens out somewhat and some of the follicles become exposed at the periphery; these can be gently teased away from the ovary. Excess tissue remaining on the follicles can be removed gently with forceps or needles; squeezing the follicles should be avoided as much as possible, as large antral follicles are easily broken. The ovary can be cut again into several smaller pieces to expose more follicles. The number of follicles obtained per ovary depends on the individual ovary as well as the number of follicles that break during the dissection. On average, we obtain 30 antral follicles from a primed mouse. 5. The best looking follicles are selected from the general pool, and 4–5 follicles are placed into the microinjection chamber. Criteria for selection are a relatively round shape without a slit through the granulosa cells or a break in the theca layer that surrounds the follicle, a visible cumulus–oocyte complex, optical clarity, and size. Follicles of equivalent appearance are set aside in a separate dish to be used for uninjected controls. Only follicles in which the nucleus is clearly discernible when the follicle is in the injection chamber are used for experiments. In our experience, almost all follicles contain such oocytes. 6. Following microinjection, the follicles are removed from the chamber and placed into fresh medium and incubated. The injected follicles are placed onto Millicell culture plate inserts in 35-mm petri dishes containing 1.6-ml medium in the reservoir below the filter; we have found that this volume is ideal because it provides the follicles plenty of medium exchange from below the membrane, but not too much so that the Millicell floats, which can result in spillover onto the membrane. Follicles are placed onto the filters using a mouth-controlled pipette. Up to 10 follicles are placed on a single filter, by drawing them up in a minimal volume of medium and spotting them onto the filter. If follicles are to be cultured for several days, the medium is replaced daily by removing 1.2 ml of medium from the culture dish and replacing it with 1.2 ml of fresh medium. Follicles are incubated in a humidified incubator with 5% CO2 and 95% air, at 37°C. 3.2. Assembly of Equipment for Microinjection
1. Sources and assembly of this equipment are described in detail in ref. (16). See Note 3. 2. The microscope to be used for injection can be upright (Fig. 12.3A) or inverted (Fig. 12.3B). The microscopes we use have a focusable stage; this is convenient, but a fixed-stage
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Fig. 12.3. Equipment for microinjecting follicle-enclosed mouse oocytes. (A) Upright microscope. (B) Inverted microscope.
microscope can also be used. With the manipulator positioned on the right, the stage controls should be on the left (a ‘‘lefthanded’’ stage), such that one hand can be used to operate the manipulator, and the other to operate the stage. We mount the manipulator on a magnetic base next to the microscope; a microscope with a narrow base is best, such that the manipulator is close to the microscope objective. A system with an upright microscope is easiest to set up; a Zeiss Axioskop works well (Fig. 12.3A). 3. An inverted microscope can also be used, although to bring in the horizontally held micropipette holder, it may be necessary to elevate the objective slightly such that the pipette holder does not contact the stage. We have used a Zeiss Axiovert 200, with a motorized stage (Fig. 12.3B); for this microscope, we needed to insert a threaded plastic spacer between the
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nosepiece and objective, to elevate the objective 6 mm. This spacer was made by a machinist, but some microscope manufacturers sell spacers for their objectives. Because the Axiovert stage is higher than that of a small upright microscope, we also needed to mount the micromanipulator on top of two 8-cmhigh magnetic bases (see Fig. 12.3B), rather than on the single magnetic base used for the Axioskop (see Fig. 12.3A). The manipulator can also be mounted on the microscope body, as long as it is arranged to bring in the micropipette horizontally. 3.3. Assembly of Equipment for Temperature, Humidity, and CO2 Control
1. We have not found any deleterious effects of keeping the oocytes in a microinjection chamber at room temperature in air (vs. a 5% CO2 environment) for a period of up to 30 min, and microinjection works equally well at room temperature or at 35°C. 2. For microinjection during imaging, we maintain the stage at 35°C and perfuse the slide with humidified 5% CO2 in air. Heating is provided by a warm air blower and monitored with an electric thermometer. Air containing 5% CO2 from a compressed gas cylinder is bubbled through a fritted glass cylinder in water, and flowed over the edge of the injection chamber through a hole in a cut-out 100-mm plastic petri dish ‘‘incubator’’ that is placed around the injection chamber (Fig. 12.4). The injection chamber is held firmly in place by gluing on magnetic strips that match corresponding strips on the microscope stage. The flow rate is maintained at 1 l/min by use of a flowmeter. Baffles cut from a silicon rubber sheet are positioned on the slide to direct the gas over the open edge of the injection chamber, and wet kimwipes inside the petri dish maintain a
Fig. 12.4. Petri dish incubator for perfusing the chamber with 5% CO2/air during injection. The micropipette enters the chamber from the right, through a cut-out section of the petri dish. Silicon rubber baffles direct the gas flow from the tube entering the dish from the upper right over the edge of the chamber.
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moist environment (see Fig. 12.4). Under these conditions, the pH of the medium is maintained at a normal level, and evaporation is minimal during a 30-min period. 3.4. Construction of Mouth Pipettes for Follicle Transfer
1. Follicles can be transferred between dishes using a Pasteur pipette that has been fire-polished at the end. For transfer into and out of microinjection chambers and onto Millicell membranes, we use mouth-controlled pipettes attached to Pyrex capillaries. We often silanize the pipettes and capillaries, to minimize sticking of the follicles to the glass during transfer, but this is not essential (see Note 4). 2. In some cases the capillary is pulled with a micropipette puller, then scored using a diamond knife to a diameter that is slightly larger than the follicles. An ordinary diamond pencil is difficult to use for this purpose; we recommend an excellent, but fragile and expensive, diamond knife (see Section 2.4.1). A ceramic knife is an inexpensive alternative that also works quite well (see Section 2.4.1). When using these knives, lay the glass to be cut on a small silicon rubber ‘‘cutting board’’ (see Section 2.4.2), on the stage of a stereomicroscope with an eyepiece reticle. Score the glass, then break it by finger pressure or a tap with a forceps. 3. Mouth pipettes can be constructed from polyethylene tubing (see Section 2.4.3), connected at one end to a plastic mouthpiece (see Section 2.4.4 and Note 1) and at the other end to a glass capillary (see Section 2.4.5). For a detailed description, see ref.(16). 4. An alternative is a preassembled mouth pipette consisting of latex tubing connected to a mouthpiece at one end, and at the other end to a silicone rubber piece into which the glass capillary is inserted (see Section 2.4.4 and Note 1). A syringe filter (see Section 2.4.6) may be attached between the latex tubing and the rubber piece, to help maintain a sterile environment; this also provides resistance so that finer control may be achieved. With this assembly, we use either the capillaries described above, or larger diameter, thicker wall capillaries (see Section 2.4.7) that are pulled out and cut to the desired diameter.
3.5. Construction and Assembly of Injection Chambers
1. The chamber for holding the follicle during injection (Fig. 12.2A and B) consists of a machined plastic slide, of the same width and length as a standard microscope slide (1" 3"), but 6 mm in thickness, with a U-shaped cutout (Fig. 12.2B; see ref.(16)). It is convenient to have a machinist make 10 or more of these reusable slides. Coverslips are attached to the upper and lower surfaces of the U-shaped cut-out, using silicon grease, forming a space 2 mm in thickness that will be filled with 400 ml of culture medium.
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2. Prior to assembly of the chamber, double-sided tape is used to attach a smaller piece of coverslip to the full-sized coverslip that will be closest to the objective (the top coverslip in the case of an upright microscope). This small piece of coverslip is positioned on the double-sided tape to form a ledge into which the follicles are placed using a mouth pipette (see below). The edge of the tape forms the back wall of the ledge, against which the follicles can be pushed during injection. For detailed diagrams of the injection chamber assembly, see ref. (16). 3. For preantral follicles (140–180 mm in diameter), we use one layer of double-sided tape, resulting in a space between the coverslips that is 100 mm in thickness; the compression of the follicle holds it firmly in place during injection, and improves visibility. 4. For antral follicles from unprimed mice (300–400 mm in diameter), we use two layers of this tape, to form an 200mm thick space. For antral follicles from primed mice (400–500 mm in diameter), two layers of double-sided tape can be used if the follicles are at the smaller end of the diameter range. For follicles that are at the larger end of the diameter range, we use two layers of double-sided tape, with one layer of ‘‘double-coated’’ tape sandwiched in between (see Note 5). Double-coated tape is slightly thinner than double-sided tape, so the combination results in a space of 240 mm in thickness. The multilayer tape assembly is formed by stacking two or three layers of tape, and then slicing them into small pieces with a small scissor. The scissor should be sharp in order to produce a clean edge. 5. If the injections are to be done at 35°C, some evaporation will inevitably occur. To minimize this problem, it is useful to position the front of the coverslip ledge 4–5 mm back from the front of the full-size coverslip. If the ledge is too far back, it is difficult to bring in the injection pipette without the pipette contacting the coverslips, but with finely tapered micropipettes, a 4–5 mm distance is workable (see Note 6). 6. The coverslip/double-stick tape chamber can also be used to hold isolated mouse oocytes for injection (see refs (6,7,18), and Note 7). 7. For use with an upright microscope, the coverslip assembly is mounted on the top surface of the plastic slide; for use with an inverted microscope, it is mounted on the bottom surface of the U-shaped cut-out (Fig. 12.2B). In either case, the ledge holding the follicles is positioned to face the reservoir of medium (Fig. 12.2A). 8. To assemble the chamber, the follicles are placed in the ledge before placing the coverslip assembly on the plastic support
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slide. For inserting preantral follicles in the ledge, the transfer pipette can be drawn out in a pipette puller and then cut with a diamond knife (see Section 3.4.2) to 200 mm internal diameter. The follicles are deposited near the edge of the ledge. The pipette is used to apply suction at the far end of ledge, which helps to pull the follicles into the gap, and also to nudge the follicles into this space. For antral follicles, the internal diameter of the drawn-out capillary should be 400–500 mm. This slight taper makes it easier to apply suction and to push the follicles into the gap. It is not necessary to move the follicle completely into the gap at this stage; halfway is enough. Once the injection slide is assembled and placed on the compound microscope stage, the micropipette can be used to pull the follicle fully under the coverslip. Likewise, the micropipette can be used to remove the follicles from the ledge after injection. 9. The last step in assembling the chamber for microinjection is to make a loading capillary, containing the solution to be injected between columns of silicon oil (see ref. (16)). Various buffers can be used for the injection solution; ionic strength does not appear to matter. For microinjection of mouse oocytes, where the pipette tips are generally smaller than for echinoderm oocytes, we use a somewhat less viscous silicon oil (20 centistokes) than previously described. The loading capillary is mounted on the injection slide using ‘‘Valap’’ (see Section 2.5.10). 3.6. Fabrication of Micropipettes
1. Micropipettes are fabricated from glass capillaries (see Section 2.6.1, and ref. (16)). The capillaries are often silanized (see Note 4), to prevent sticking of solutions to the inside wall of the micropipettes; however, silanizing is not necessary when injecting solutions of small molecules. The glass is pulled in a horizontal puller to form micropipettes, and an 1-mm column of mercury is placed in the back of the pipette using a Hamilton syringe; pipettes made in this way can be stored in a covered container for months (see ref. (16)). 2. Just prior to use, each individual micropipette tip is broken slightly, such that its external diameter is 1 mm. This is accomplished by gently tapping the micropipette tip against the face of the loading capillary. The tip should be broken to the smallest diameter that still allows the injection solutions to be drawn up. The major cause of oocyte damage is pipette tips that are too large! If possible, try to break off the tip such that it has a slight bevel; the sharper the tip, the easier it is to insert it into the follicle and oocyte.
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3. Bevelling of the pipette tip by touching it to a rotating surface covered with a fine abrasive paper (19,20) is an alternative way to produce a sharp point with maximal surface area of the tip opening. 3.7. Injection Procedure
1. The microinjection procedure described here is a modification of methods that have been developed for echinoderm oocytes (see ref. (16)). Here, we provide details of the procedure as we have adapted it to be used for injecting follicle-enclosed mouse oocytes. 2. Because the basic setup is the same as that used for echinoderm or small frog oocytes, we recommend first practising on these cells if they are available. If not, we suggest beginning with follicle-free mouse oocytes, or mouse oocytes within preantral follicles, as these follicles have fewer layers of surrounding cells and the oocyte is more optically clear. In addition, the survival rate for preantral follicle-enclosed oocytes is higher (100%) than for oocytes within antral follicles (50%). Once the basic method is mastered, then it is easier to move up to injecting oocytes within larger antral follicles. 3. This microinjection procedure is highly quantitative; see ref. (16) for details on calibration. The final concentration of an injected substance is based on an oocyte volume of 200 pl (70–75-mm diameter), and the volumes injected typically range from 4 to 14 pl (2–7% of the cell volume), though volumes of up to 20 pl (10%) can be used. 4. Start by positioning the follicles within the injection chamber, using a micropipette. If the portion of the follicle containing the oocyte is not fully in the chamber, the follicle can be moved further in by using the injection pipette to either roll or drag it until the oocyte is completely under the coverslip. When antral follicles are used, it is often helpful to orient the follicle such that the antrum is closest to the injection pipette, with the oocyte near the tape at the back of the chamber; this helps the oocyte stay in place when the pipette pushes against it. It is also useful to position the follicle such that the oocyte nucleus is not near the site where the pipette will enter the oocyte. 5. After positioning the follicles, insert a new micropipette into the pipette holder, and break the tip slightly (see Section 3.6.2). To load the pipette, the mercury is first pushed to the tip. A column of silicon oil from the loading capillary (see ref. (16)) is drawn into the pipette to separate the mercury from the injection solution. A measured amount of injection solution is drawn up next, followed by an oil cap that is used to prevent mixing of the injection solution and culture
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medium (see Fig. 12.2A). Avoid drawing up too much backing oil, and make a small oil cap. Drawing the oil cap into the pipette can be difficult, and is in large part dependent on the size of the pipette tip. Several tricks can be used to produce an oil cap if one does not easily form the first time (see Note 8). 6. Once the pipette has been loaded, it is moved into the chamber in front of the follicle (see Note 6). The focus should be adjusted to obtain a sharp image of the oocyte edge, and the nucleus and nucleolus should be readily visible. The plane of the injection pipette should be adjusted with the manipulator so that the tip is in focus with the edge of the oocyte. 7. The pipette is then pushed through the wall of the follicle. If the pipette moves out of focus as it enters the follicle, it should be moved with the manipulator until it is back in focus. Once the tip pops through the granulosa cell layers, it is inserted into the oocyte. 8. It is not necessary to push the pipette far into the oocyte, and the nucleus should be avoided. Once the pipette pops through the plasma membrane, the solution can be expelled. The oil cap will enter the oocyte first, followed by the injection solution, which will visibly disperse throughout the cytoplasm. Once the solution is expelled, the pipette can be removed slowly from the oocyte and follicle. 9. Pipettes can be reused several times for non-sticky solutions. 10. For safety, used pipettes should immediately be placed in a covered waste container.
4. Notes 1. The Sigma mouth pipette assembly provides a round mouthpiece. Flat mouthpieces are more comfortable, but are not currently available. The round mouthpieces that come with the Sigma mouth pipette assembly can be flattened by warming the plastic in hot water and pinching gently with a pliers, or you can make a mouthpiece from the outer barrel of a 1-ml plastic syringe; cut off an 3-cm piece including the tip, and flatten the wide end with a pliers. 2. A well-primed mouse has swollen, dark pink uteri and enlarged, dark pink ovaries that are not excessively bloody. Antral follicles from these ovaries are generally >360 mm in diameter, and the mural granulosa cells appear dark, but with large antra and easily discernible oocytes in the interior. In contrast, the uteri and ovaries from overprimed mice are larger and dark red, due to more blood vessels. Follicles from these
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ovaries often contain oocytes that undergo meiotic resumption spontaneously after a few hours in culture, and such ovaries often contain corpora lutea (large structures without an antrum that develop from follicles that have ovulated). In addition, these ovaries often appear to be ‘‘spongy’’ and the follicles fragile. Unprimed mice have uteri and ovaries that are lighter pink, and the follicles obtained from these mice are smaller. 3. Place the injection setup in an undisturbed environment where there is little traffic. For your well-being, provide comfortable eye pieces and chair. Good music is also recommended – in our experience, oocytes like classic rock. 4. Silanizing should be performed in a fume hood. We use Sigmacote (see Section 2.4.8), by dipping a small bundle (20) of capillaries directly into the bottle and then inverting the capillaries several times to coat the inside of the glass. Pipettes are then blotted onto kimwipes and left for 2 weeks in the hood to completely dry. Pasteur pipettes used for transferring follicles can be silanized in the same way. 5. Double-coated tape is made to be removable, while doublesided tape is made to be permanent; the double-sided tape is the most common type. A chamber made with double-coated tape alone does not hold together reliably, since this type of tape is less adherent. Thus, double-coated tape is only useful for chamber construction when it is sandwiched between double-sided tape. 6. If the pipette contacts the coverslips and fails to move into the space between them without losing focus, remove the pipette and carefully tip the front of the chamber upwards slightly (a few hundred microns); upwards applies to an upright microscope. On an inverted microscope, the front of the chamber should be tipped slightly downwards. On our inverted microscope, the magnetic strip on the stage that matches the magnetic strip on the plastic slide (see Fig. 12.4) is permanently slightly elevated in the back, for this reason. 7. Isolated oocytes are most commonly injected using methods like those described in ref. (20), but the methods we describe here can also be used. Since the diameter of an isolated oocyte is 70–75 mm, it is somewhat loose in a chamber formed by one layer of double-sided tape, and the oocyte often comes out of the chamber when the micropipette is withdrawn after injection. If it is desirable to keep the isolated oocyte in the chamber after injection, it helps to withdraw the pipette quickly, by quickly moving either the pipette or the microscope stage. Another solution is to use the serrated plastic edge of the tape dispenser to produce a serrated edge on the side of
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the tape that will face the front of the chamber. Tearing the tape against the plastic edge results in irregular ‘‘hooks’’ that can be used to hold the oocyte in place as the pipette is withdrawn. 8. Several tricks can be used to produce an oil cap if one does not form easily the first time: (1) tapping on the manipulator, or flicking the micropipette holder, (2) quickly moving the pipette in and out of the interface between the loading capillary oil and air, and (3) loading the injection solution until the amount is stable, then breaking the pipette tip on the capillary again. This must be done with care to avoid breaking the tip too large. It is also useful to replace the silicone gasket in the injection pipette holder (see ref. (16)). Occasionally, the solution is too sticky for the pipette to cap despite trying all of the above. In these cases, it is sometimes possible to omit the oil cap. For this to work, it is important to ensure that the solution is stable such that no culture medium is drawn into the pipette when it is placed into the chamber.
Acknowledgments We thank John Eppig, Marilyn O’Brien, and Karen Wigglesworth for showing us how to obtain and culture mouse follicles, and Melina Schuh for critical reading of the manuscript. This work was supported by grants from the NIH.
References 1. Mehlmann, L.M., Jones, T.L.Z., and Jaffe, L.A. (2002) Meiotic arrest in the mouse follicle maintained by a Gs protein in the oocyte. Science 297, 1343–1345. 2. Mehlmann, L.M., Saeki, Y., Tanaka, S., Brennan, T.J., Evsikov, A.V., Pendola, F.L., Knowles, B.B., Eppig, J.J., and Jaffe, L.A. (2004) The Gs-linked receptor GPR3 maintains meiotic arrest in mammalian oocytes. Science 306, 1947–1950. 3. Mehlmann, L.M., Kalinowski, R.R., Ross, L.F., Hewlett, E.L., and Jaffe, L.A (2006). Meiotic resumption in response to luteinizing hormone is independent of a Gi family G protein or calcium in the mouse oocyte. Dev. Biol. 299, 345–355. 4. Kalinowski, R.R., Berlot, C.H., Jones, T.L.Z., Ross, L.F., Jaffe, L.A., and Mehlmann, L.M.
(2004) Maintenance of meiotic prophase arrest in vertebrate oocytes by a Gs proteinmediated pathway. Dev. Biol. 267, 1–13. 5. Simon, A.M., Goodenough, D.A., Li, E., and Paul, D.L. (1997) Female infertility in mice lacking connexin 37. Nature 385, 525–529. 6. Freudzon, L., Norris, R.P., Hand, A.R., Tanaka, S., Saeki, Y., Jones, T.L.Z., Rasenick, M.M., Berlot, C.H., Mehlmann, L.M., and Jaffe, L.A. (2005) Regulation of meiotic prophase arrest in mouse oocytes by GPR3, a constitutive activator of the Gs G protein. J. Cell Biol. 171, 255–265. 7. Norris, R.P., Freudzon, L., Freudzon, M., Hand, A.R., Mehlmann, L.M., and Jaffe, L.A. (2007) A Gs-linked receptor maintains meiotic arrest in mouse oocytes, but
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luteinizing hormone does not cause meiotic resumption by terminating receptor-Gs signaling. Dev. Biol. 310, 240–249. Mehlmann, L.M. (2005) Oocyte-specific expression of Gpr3 is required for the maintenance of meiotic arrest in mouse oocytes. Dev. Biol. 288, 397–404. Eppig, J.J., Viveiros, M.M., Marin-Bivens, C., De La Fuente, R. (2004) Regulation of mammalian oocyte maturation, in The Ovary, 2nd edition (Leung, P.C.K. and Adashi, E.Y., ed.), Elsevier/Academic Press, San Diego, CA, pp. 113–129. Mehlmann, L.M. (2005) Stops and starts in mammalian oocytes: recent advances in understanding the regulation of meiotic arrest and oocyte maturation. Reproduction 130, 791–799. Nikolaev, V.O., and Lohse, M.J. (2006) Monitoring of cAMP synthesis and degradation in living cells. Physiology 21, 86–92. Matzuk, M., Burns, K.H., Viveiros, M.M., and Eppig, J.J. (2002) Intercellular communication in the mammalian ovary: oocytes carry the conversation. Science 296, 2178–2180. Park, J.Y., Su, Y.Q., Ariga, M., Law, E., Jin, S.L.C., and Conti, M. (2004) EGF-like growth factors as mediators of LH action in the ovulatory follicle. Science 303, 682–684.
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14. Hiramoto, Y. (1962) Microinjection of the live spermatozoa into sea urchin eggs. Exp. Cell Res. 27, 416–426. 15. Kiehart, D.P. (1982) Microinjection of echinoderm eggs: apparatus and procedures. Meth. Cell Biol. 25, 13–31. 16. Jaffe, L.A., and Terasaki, M. (2004) Quantitative microinjection of oocytes, eggs, and embryos. Meth. Cell Biol. 74, 219–242. 17. Su, Y.-Q., Denegre, J.M., Wigglesworth, K., Pendola, F.L., O’Brien, M.J., and Eppig, J.J. (2003) Oocyte-dependent activation of mitogen-activated protein kinase (ERK1/2) in cumulus cells is required for the maturation of the mouse oocyte–cumulus cell complex. Dev. Biol. 263, 126–138. 18. Schuh, M., and Ellenberg, J. (2007) Selforganization of MTOCs replaces centrosome function during acentrosomal spindle assembly in live mouse oocytes. Cell 130, 484–498. 19. Kaila, K., and Voipio, J. (1985) A simple method for dry beveling of micropipettes used in the construction of ion-selective micro-electrodes. J. Physiol. 369, 8p. 20. Kline, D. (2009) Quantitative microinjection of mouse oocytes and eggs. Methods Mol. Biol. 518.
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Chapter 13 Functional Studies of Regulatory Genes in the Sea Urchin Embryo Vincenzo Cavalieri, Maria Di Bernardo, and Giovanni Spinelli Abstract Sea urchin embryos are characterized by an extremely simple mode of development, rapid cleavage, high transparency, and well-defined cell lineage. Although they are not suitable for genetic studies, other approaches are successfully used to unravel mechanisms and molecules involved in cell fate specification and morphogenesis. Microinjection is the elective method to study gene function in sea urchin embryos. It is used to deliver precise amounts of DNA, RNA, oligonucleotides, peptides, or antibodies into the eggs or even into blastomeres. Here we describe microinjection as it is currently applied in our laboratory and show how it has been used in gene perturbation analyses and dissection of cis-regulatory DNA elements. Key words: Microinjection, sea urchin embryo, gene function, cis-regulatory analysis, PCRamplified transgenes, green fluorescent protein, Orthopedia homeobox-containing gene.
1. Introduction Regulatory proteins act as switches of development (1,2). By regulating their downstream effectors, either regulatory or structural genes, they mediate and amplify the large flow of information encoded in the genome. The net result of such a complex project is the balanced and controlled construction of a new organism. Species-specific programs of development are encoded in the cis-regulatory apparatus of the genome which, through progressive steps, establishes local combination of transcription factors, thus defining cell identity (3–5). Realization of these events relies on multiple interactions among gene products, whose expression is highly regulated in response to signals and to the different combinatorial environment of each group of cells. David J. Carroll (ed.), Microinjection: Methods and Applications, Vol. 518 Ó 2009 Humana Press, a part of Springer ScienceþBusiness Media, LLC DOI 10.1007/978-1-59745-202-1_13
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Besides these mechanisms, additional regulatory levels in the cell, dictated by local chromatin structure remodeling and by posttranscriptional regulation events, make the picture much more complicated than that represented by the architecture of gene regulatory networks alone. Functional studies of given genes are usually preceded by the analysis of the temporal activation and spatial distribution of their transcripts. Basic information on gene expression timing, size, and number of the transcripts come from a combination of widely used methods: Northern blot hybridization (6), RNAse protection assays (7), Quantitative reverse transcription-PCR (RT-PCR) (8), and in situ hybridization (9). Looking at the activity of single genes, these methodologies tell us when and where they are expressed. However, the unravelling of the cis-regulatory elements, and the role played by regulatory genes in development or differentiation, requires an inverse genetic methodology, such as that given by DNA and RNA microinjection in living embryos. Gene function is currently approached by gain-of-function or loss-of-function studies conducted by extensive or clonal ectopic expression, functional knock down or in the case of transcription factors, by binding site competition. The effects of this interfering action can be then directly assayed on candidate targets or on a large panel of genes, by high-throughput approaches. Gene regulation of single gene units is primarily approached by promoter function analysis, in order to identify cis-regulatory elements involved in modulating gene activity in time and space. DNA constructs, containing large and sequentially deleted gene-specific regulatory elements, are injected and tested by the ability to activate downstream reporter genes (10–14). Finally, gene databases (available for multiple organisms and updated almost daily) permit in silico analyses and the identification of sites of binding for specific trans-regulatory factors. In this chapter we will illustrate in detail the experimental procedures used to study gene function and regulation during the embryonic development of the sea urchin Paracentrotus lividus. In particular, we will describe evidence obtained by applying these techniques for the study of Orthopedia (Otp), a regulatory gene encoding a homeodomain transcription factor involved in the synthesis of the embryonic skeleton (15,16).
2. Materials 2.1. Microinjection Apparatus and Needles
1. Binocular inverted microscope (Leitz Fluovert). 2. Micromanipulator provided with joystick (Leitz). 3. Microinjector (Eppendorf 5242).
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4. Pressurized Nitrogen Tank. 5. Injection needles (Femtotips I or II, Eppendorf). 6. Pipette tips to fill in microinjection needles: Microloader (Eppendorf). 2.2. Microinjection Solutions
1. UltraPure water (Invitrogen). Store at room temperature (see Note 1). 2. UltraPure 100% (w/v) DNAse, RNAse, and protease-free glycerol (Invitrogen). Store at room temperature. 3. Texas Red-conjugated dextran (TRCD, Molecular Probes, Eugene, OR) is dissolved in water at 15% (w/v), stored in aliquots wrapped in aluminium foil at 20°C. 4. Antisense morpholino-substituted (AMS) oligonucleotides (Gene Tools LLC, Corvallis, OR) are resuspended in water to obtain 5 mM stock solutions. The optimal concentration of AMS oligonucleotides in the final injection solution range from 100 to 600 mM. Each oligonucleotide should be tested and the minimal concentration which gives rise to a significant phenotype should be used. The AMS oligonucleotide injection solutions are highly stable and, if stored at 20°C, can be used repeatedly for several months.
2.3. Microinjection Plates and Egg Handling Buffers
1. Cell culture petri dishes (60 mm, Orange Scientific). 2. Protamine sulphate (Sigma) is dissolved in distilled water at 1% (w/v) and stored at 4°C (see Note 2). 3. The mouth pipette is assembled by attaching a mouthpiece to one end of a piece of rubber latex tubing and a Pasteur pipette with a hand-pulled tip to the opposite end, as shown in Fig. 13.1D. 4. Millipore filtered sea water (MFSW) is obtained by filtering sea water firstly with a pleated paper filter to remove large particulate matter and then with a 0.45 mm membrane filter (Millipore). Store at 18°C. 5. Citric acid monohydrate (J.T. Baker) is dissolved in distilled water at 0.5 M. Store at room temperature. 6. Tris-HCl pH 8.0 (1 M). Store at room temperature.
2.4. Preparation of Synthetic mRNAs
1. Plasmid expression vectors of the pCS2 series are digested with the appropriate restriction enzymes (see Note 3). 2. In vitro transcription kit. We use mMESSAGE mMACHINE High Yield Capped RNA Transcription Kit with SP6 polymerase, Ambion. 3. Stock solutions of Proteinase K (Sigma) at 20 mg/ml are maintained as small volume frozen aliquots at 20° C.
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Fig. 13.1. Equipment for microinjection. (A) The microinjection apparatus. (B) The injection plate: a track marks the line where unfertilized dejellied eggs will placed down, as shown in (C), by the use of a mouth pipette. (D) The mouth pipette is assembled by attaching a mouthpiece to one end of a rubber latex tube and a hand-pulled Pasteur pipette with a blue tip to the opposite end.
4. Phenol/Chloroform (5:1 v/v) pH 8.0 (Sigma). Store at 4°C, shielding from light. 5. Phenol/chloroform/isoamyl Alcohol (125:24:1 v/v/v) pH 4.7 (Sigma). Store at 4°C, shielding from light. 6. TE buffer (pH 7.6): 10 mM Tris-HCl pH 7.6, 1 mM EDTA. Store at room temperature. 7. EDTA pH 7.0 (0.5 M). Store at room temperature. 8. SDS 20% (w/v). Store at room temperature. 9. Isopropyl alcohol (abs., Merck). Store at room temperature. 2.5. PCR Amplification of DNA Transgene Constructs
1. Template DNA transgene construct and specific forward/ reverse PCR primer pairs. 2. High-fidelity Pfu DNA polymerase (Promega). 3. 10X PCR Buffer: 200 mM Tris-HCl (pH 8.8 at 25°C), 100 mM KCl, 100 mM (NH4)2SO4, 20 mM MgSO4, 1% Triton X100, and 1 mg/ml BSA.
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4. dNTPs mix (20 mM each of dATP, dCTP, dGTP, and dTTP in water, Amersham). Store at 20°C. 5. Agarose, electrophoresis buffers and tanks, power packs, gel viewing system.
3. Methods Pressure-based microinjection of sea urchin eggs, early embryos, or single blastomeres constitute the most commonly used approach to introduce almost any kind of test molecule that can potentially interfere with gene function, including DNA transgene constructs, synthetic mRNAs, AMS oligonucleotides, antibodies, peptides, and fluorescent probes (17–22). We use a binocular inverted microscope with transmitted light (Leitz Fluovert) equipped with a micromanipulator which holds the microinjection needle and a joystick which moves the needle in the three dimensions. Accuracy and reproducibility of the injection parameters – pressure, time, and compensation pressure – can be easily ensured by the use of a microinjector which is connected to a tank of pressurized nitrogen (see Fig. 13.1A). In a typical session lasting 3–4 h, a practiced hand can inject several hundred specimens. Here we describe methods currently used in our laboratory for microinjecting zygotes of the Mediterranean species of sea urchin P. lividus. 3.1. Preparation of Plates and Eggs for Microinjection
1. Lids of 60 mm cell culture petri dishes constitute the best choice for microinjection. By using a fine-point marker, two straight parallel lines are drawn on the outer side of each lid, as shown in Fig. 13.1B. The lane between the two lines will be used as a guide when rowing the eggs. Inner surfaces of the lids are then coated with a 1% protamine sulphate solution approximately for 2 min, rinsed with deionized water, and finally placed down on a sheet of absorbent paper to air dry. Treated lids can be stored at room temperature for several weeks. 2. Male gametes from adult animals are collected and stored at 4°C for up to 1 week. The eggs are harvested in MFSW and are viable for microinjection only for few hours. Small amounts of eggs are dejellied simply by allowing incubation in acidified MFSW (pH 5.0, obtained by adding few drops of 0.5 M citric acid) for approximately 1 min, with gentle swirling. Then, pH is quickly brought back to the normal value by adding TrisHCl pH 8.0 and eggs are collected and resuspended in fresh MFSW.
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3. To increase rapid and successful microinjection, dejellied eggs are arranged in neat rows of 180–240 units on the treated inner surfaces of the injection plates filled with fresh MFSW, by using a mouth pipette of correct size and shape, as shown in Fig. 13.1C. The eggs will adhere to the surface of the dishes immediately upon contact. Fertilization is obtained by placing few drops of a sperm suspension diluted in MFSW just above the row and zygotes are microinjected with 1–2 pl of the appropriate sample. After injections have been completed, the embryos are allowed to develop at 18°C until reaching the desired stage, mounted on glass slides, and examined under an upright microscope using brightfield illumination and epifluorescence. 3.2. Preparation of Microinjection Samples
1. The injection sample solution may contain either AMS oligonucleotides, synthetic mRNAs or DNA transgene constructs as test molecules at the appropriate concentrations (further details are given below); glycerol and TRCD are invariably added at final concentrations, respectively, of 30% and 5% in a total volume of 5 ml (see Note 4). 2. Immediately prior to injection, samples are centrifuged for 10 min at 13,000 g at 4°C, to pellet any particulate material that might clog the injection needle. By using a microloader pipette tip, approximately 0.8 ml of injection solution are loaded into the needle and the tip is filled within few seconds by capillary action. The remaining sample aliquot should be kept on ice during the microinjection session.
3.3. Unravelling Gene Function
The most widely used approaches to decrypt the function of genes during embryonic development involve mis/overexpression and loss-of-function assays. Disruption of gene function can be accomplished by microinjection of AMS oligonucleotides (23–26). Annealing of such kind of a oligonucleotide to the 50 UTR, encompassing the AUG codon and the first bases of coding sequence, guarantees an efficient block of translation initiation (an exhaustive discussion and technical details can be found at http:// www.gene-tools.com). Loss of function can be also obtained by injection of transcripts encoding for modified variant proteins that specifically compete with the endogenous gene product (27–29). A transcriptional activator, for instance, could be converted into an obligate negative regulator simply by fusing the DNA-binding domain encoding sequences to those of the well-characterized dEngrailed repressor domain (dominant-negative construct, see ref. (30,31)). Conversely, gain of gene function can be achieved by microinjection of synthetic exogenous mRNA molecules, encoding for the wild-type protein (32–34). Moreover, mRNA injection approach efficiently complements experiments with AMS
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oligonucleotides and is useful to assess their specificity by the rescue of gene function (35–37). The AMS oligonucleotidesmediated loss of function assay coupled to the overexpression of exogenous mRNAs encoding either the wild-type or the dominant-negative variant of the protein, has been crucial in order to assess the involvement of the Otp homeobox-containing gene in patterning the sea urchin embryo (Fig. 13.2). A detailed protocol to prepare ‘‘microinjection grade’’ synthetic mRNA is described below (see Note 5): 1. Insert the cDNA fragment encoding for the protein of interest into a transcription vector (pCS2 is preferred) and obtain
Fig. 13.2. Orthopedia (Otp) gene function studies in sea urchin. Otp encodes for a transcriptional activator. (A, B) Embryos injected with 2 pl of 500 mM Otp AMS oligonucleotide (AMSo) lack any skeletal element. The inhibition of Otp translation in the oral ectoderm abolishes the signaling cascade that induces the underlying primary mesenchyme to initiate skeletal synthesis and growth. (C) Embryo injected with 1 pg of in vitro transcribed capped mRNA encoding for an En-Otp chimera with a dominantnegative effect. Skeletal synthesis is blocked as well as in embryos injected with the AMSo. (D, D0 ) Otp misexpression. Brightfield and darkfield images of an embryo injected with 1 pg of full-length Otp mRNA causes the growth of abnormal spicule elements and the formation of a radial-shaped skeleton. (E) Function was fully rescued when Otp AMSo and synthetic myc tag-Otp mRNA were co-injected. The presence of an excess of a translatable Otp mRNA caused the appearance of well-defined gain-of-function phenotypes. (F, F0 ) Clonal expression of the Otp transgene. Fluorescent and brightfield images of an embryo injected with the HE-OTP-GFP DNA construct. The mosaic expression of the Otp gene was guided in the ectoderm by the Hatching Enzyme (HE) promoter and visualized by the presence of the GFP reporter. In correspondence to the Otp ectopic expression (F), primary mesenchyme cells synthesized an extra spicule (arrow in F0 ). (G) A glycerol-injected embryo at pluteus stage. All embryos are observed at 48 h after fertilization.
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highly pure plasmid DNA by your favorite method or commercial kit. 2. Linearize 10 mg of plasmid DNA with an appropriate restriction enzyme that cuts at the 30 -side of the protein-coding region and downstream of the poly-adenylation signal (see Note 6). 3. Check a small aliquot on an agarose gel to confirm that cleavage is complete. 4. Incubate the linearized plasmid DNA with 100–200 mg/ml of proteinase K and 0.5% SDS for 30 min at 50°C. 5. Extract the DNA twice by using an equal volume of phenol/ chloroform. 6. Remove any trace of phenol by a further chloroform extraction and precipitate DNA with 2.5 volumes of ethanol in presence of 0.3 M Na-Acetate. Chill at 20°C for at least 15 min. 7. Resuspend the DNA in RNAse-free water or TE buffer (pH 7.6) at 0.5–1 mg/ml. 8. Assemble the in vitro transcription reaction by using 1 mg of purified DNA as template, following the manufacturer’s kit instructions. Incubate for 2 h at 37°C. 9. Add 1–2 units of RNAse-free DNAse I and incubate at 37°C for 15 min, to remove template DNA (see Note 7). 10. Stop the reaction by adding 1 ml of 0.5 M EDTA (pH 7.0) and purify transcripts by extracting with an equal volume of phenol/chloroform/isoamyl alcohol (pH 4.7) and then with an equal volume of chloroform. 11. Precipitate the RNA by adding 1 volume of isopropyl alcohol and chill at 20°C for at least 15 min. 12. Centrifuge at 13,000 g for 10 min at 4° C and resuspend the RNA in water. 13. Estimate concentration by optical density at 260/280 nm and run an aliquot on a denaturing agarose gel. Keep in small aliquots at 80°C until use. 14. Prepare the injection solution at a final mRNA concentration of 0.1–2 mg/ml. Following microinjection into zygotes, synthetic mRNA molecules are distributed among all the blastomeres of the early embryo and translated in their protein products, leading to a global perturbation of gene function. Nevertheless, mis-expression in a restricted subset of embryonic cells can be carried out by injection of linearized DNA constructs in which the protein-coding sequence is cloned downstream of tissue-specific promoters
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(15). Because DNA incorporation in transgenic sea urchin embryos occurs in a mosaic fashion (38,39), the transgene, properly guided by the promoter, is expressed only in a clone of embryonic cells that represent a sub-fraction of the wider gene expression domain. An example of such clonal expression is shown in Fig. 13.2F-F0 . In this case, we injected a plasmid construct in which the sequence of the Otp-GFP-encoding fusion-protein was cloned downstream of the P. lividus Hatching enzyme (HE) promoter. This promoter region targets the expression of the transgene solely to the ectoderm territory (40). Because of the fluorescence of the GFP reporter, we easily visualized the cellular clones expressing the transgene and exactly correlated them with the mineralization of supernumerary embryonic skeletal elements. The advantages/disadvantages of the GFP reporter system are presented in further details in the section below. 3.4. Identification of Regulatory Elements by In Vivo Promoter/ Reporter Transgene Assays
Although the extent of mosaicism of injected transgenes can be counterbalanced by injecting large numbers of embryos, there is the possibility of unexpected artificial effects due to the presence of the plasmid DNA sequences that might lead to the suppression of the transgene expression levels or the ectopic enlargement of the spatial expression domains. These artefacts might be misleading especially if one wishes to study the cis-regulatory apparatus of a particular gene. In order to overcome the above-mentioned problems, a straightforward strategy involves the microinjection of purified PCR-amplified DNA fragments that consist only of either the entire or partially deleted cis-regulatory regions placed upstream of a reporter gene (10). It should be emphasized that in our hands, the use of promoter/reporter DNA fragments generally results in higher number of reporter-expressing embryos and the size of the spatial expression domains is usually larger than that observed in embryos injected with the corresponding plasmid constructs (10). A convenient reporter system is given by the GFP gene from the hydromedusa Aequorea Victoria, which encodes a naturally green fluorescent protein that allows nondestructive detection of spatial gene expression within living embryos (41). By this method we easily and quickly obtained a deleted series of constructs containing genomic Otp 50 -flanking regions placed upstream of the GFP-derived Green Lantern-1 reporter gene and successfully dissected the gene regulatory regions (10). These and other examples are shown in Fig. 13.3. A general approach is given as follows: 1. Fuse a large 50 -flanking genomic fragment of the gene of interest, including the transcription and translation start sites, upstream and in frame with the GFP reporter gene. 2. Use the DNA plasmid construct as template to generate, by use of high-fidelity Pfu DNA polymerase, a functional
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Fig. 13.3. cis-Regulatory analysis in sea urchin embryos. Brightfield (A, B), fluorescent (A0 , B0 ), and merged (C–F) images of embryos after 70 h (A-D), 10 h (E), 48 h (F) of development. Zygotes were injected with three different constructs. (A-B) A 2.4 kb Otp promoter region containing the ATG start codon of the Otp gene at the 30 end, fused in frame with the green fluorescent protein reporter gene (pOtp-GFP 2.4) drives GFP localization at the sites of expression of the endogenous Otp gene, viz, the tips of the post-oral arms (A-A0 ) and flanking the mouth (B, B0 ). On the other hand, a deleted pOtp-GFP 0.85 transgene allows ectopic GFP expression in mouth and foregut (C, D). Moreover, upon injection of the deleted regulatory region, an increase of GFP expression was observed, probably due to the lack of some negative regulatory elements that maintain the highly restricted Otp expression pattern. (E) A 10 h old embryo injected with a 2.9 kb HE promoter construct (pHE-GFP). Labeled cells are localized in the ectoderm of the blastula stage embryo. (F) A 48 h old embryo injected with a 1.45 kb Hbox12 promoter construct (pHbox12-GFP). PlHbox12 is an early-expressed homeobox gene (7). The use of the GFP reporter allows the observation of the expression pattern even at stages in which the gene is firmly silenced. Fluorescence is clearly restricted to cell clones of the aboral ectoderm territory (unpublished results).
transgene in which the plasmid sequences are excluded (see Note 8). For each amplification reaction the following components are mixed together in a total volume of 50 ml (final amounts/concentrations are indicated in brackets): Template DNA (1–5 ng) Forward and Reverse primers (0.2–0.4 mM) dNTP mix (0.2 mM) PCR buffer (1X) Pfu DNA polymerase (1–2 U) A common PCR cycling program usually gives sufficient product after 30 cycles of: strand denaturation at 94°C for 10–60 s, primer annealing at 55–65°C (depending on the particular primer pair) for 30–60 s, primer extension at 72° C for 90 s per kilobase of expected PCR product. An additional final extension step of 5–10 min at 72°C ensures that all amplicons are fully extended. 3. Purify the PCR product DNA fragment to a microinjection grade (we use the spin-column-based QIAquick PCR Purification Kit, Qiagen). 4. Quantify the purified DNA fragments by spectrophotometer readings at 260/280 nm.
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5. Assemble the microinjection solution and inject the promoter/reporter transgene construct to assess if its spatial expression pattern is congruent with that of the endogenous gene. If this is not the case, try to clone a larger genomic fragment. 6. Repeat steps 2–4 in order to prepare a series of progressively shorter DNA fragments truncated starting from the upstream regulatory sequences. 7. Assemble the microinjection solutions and check the spatial expression pattern of each deleted construct during embryonic development (see Note 9). Due to the extreme stability of the GFP protein in the sea urchin embryo (41), the fluorescence detected at a given time must to be viewed as the sum of all prior expression, hampering the identification of both the shutoff of gene expression and the cis-regulatory elements involved in the downregulation. In order to overcome this limit in the promoter analysis of a given transgene, the GFP mRNA level can be measured by semi-quantitative RT-PCR or real-time quantitative PCR assays in total RNA extracted from transgenic embryos (10,11,14).
4. Notes 1. Unless stated otherwise, all injection solutions and samples should be prepared in UltraPure distilled water (Invitrogen) tested for the absence of nuclease activity. This standard is referred to as ‘‘water’’ in this text. 2. After prolonged storage, some protamine sulphate might precipitate. Each time plates are prepared, the solution should be incubated on a orbital shaker at room temperature until fully dissolved. 3. pCS2 vectors are not commercially available but can be requested to Dave Turner through the website http://sitemaker.umich.edu/dlturner.vectors. 4. The addition of the freely diffusing internal fluorescent standard TRCD easily allows to distinguish injected from uninjected eggs. 5. In order to avoid any RNAse contamination, all solutions, whenever possible, should be treated with 0.1% diethyl pyrocarbonate (DEPC) and autoclaved. The use of sterile disposable plasticware is recommended as well as the baking of glassware at 180°C for 3 h before use. Electrophoresis tanks should be cleaned with 1 N NaOH and abundantly rinsed with nuclease-free water. Gloves should always be worn and changed frequently.
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6. High yields of in vitro transcription are obtained if plasmid templates are cut with restriction enzymes leaving 50 -overhanging ends (e.g., EcoRI, BglII, HindIII, etc.). 7. Before or after DNAse treatment, an aliquot of the reaction should be run on a denaturing agarose gel to assess the yield of transcription. 8. Depending on the particular length of the putative genomic cis-regulatory region, the Elongase Amplification System (Invitrogen) could be suitable for the amplification of targets longer than 5 kb with high yields. 9. Approximately 5000 molecules per embryo either of plasmid or PCR-amplified DNA transgene constructs are injected in a volume of 1–2 pl.
References 1. Angerer L. M., Oleksyn D. W., Levine A. M., Li X., Klein W. H., Angerer R. C. (2001) Sea urchin goosecoid function links fate specification along the animal–vegetal and oral–aboral embryonic axes. Development, 128, 4393–4404. 2. Mao C. A., Wikramanayake A. H., Gan L., Chuang C. K., Summers R. G., Klein W. H. (1996) Altering cell fates in sea urchin embryos by overexpressing SpOtx, an orthodenticle-related protein. Development, 122, 1489–1498. 3. Levine M., Davidson E. H. (2005) Gene regulatory networks for development. Proc. Natl. Acad. Sci. USA, 102 (14), 4936–4942. 4. Oliveri P., Davidson E. H. (2004) Gene regulatory network controlling embryonic specification in the sea urchin. Curr. Opin. in Gen. & Dev., 14, 351–360. 5. Davidson E. H., Rast J. P., Oliveri P., Ransick A., Calestani C., Yuh C. H., Minokawa T., Amore G., Hinman V., Arenas-Mena C., Otim O., Brown C. T., Livi C. B., Lee P. Y., Revilla R., Rust A. G., Pan Z., Schilstra M. J., Clarke P. J., Arnone M. I., Rowen L., Cameron R. A., McClay D. R., Hood L., Bolouri H. (2002) A genomic regulatory network for development. Science, 295 (5560), 1669–1678. 6. Illies M. R., Peeler M. T., Dechtiaruk A. M., Ettensohn C. A. (2002) Identification and developmental expression of new biomineralization proteins in the sea urchin
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wnt8 gene in the sea urchin endomesoderm network. Dev. Biol., 288, 545–558. Revilla-i-Domingo R., Minokawa T., Davidson E. H. (2004) R11: a cis-regulatory node of the sea urchin embryo gene network that controls early expression of SpDelta in micromeres. Dev. Biol., 274, 438–451. Cavalieri V., Spinelli G., Di Bernardo M. (2003) Impairing Otp homeodomain function in oral ectoderm cells affects skeletogenesis in sea urchin embryos. Dev. Biol., 262 (1), 107–118. Di Bernardo M., Castagnetti S., Bellomonte D., Oliveri P., Melfi R., Palla F., Spinelli G. (1999) Spatially restricted expression of PlOtp, a Paracentrotus lividus orthopediarelated homeobox gene, is correlated with oral ectodermal patterning and skeletal morphogenesis in late cleavage sea urchin embryos. Development, 126, 2171–2179. Di Caro D., Melfi R., Alessandro C., Serio G., Di Caro V., Cavalieri V., Palla F., Spinelli G. (2004) Down-regulation of early sea urchin histone H2A gene relies on cis-regulative sequences located in the 50 and 30 regions and including the enhancer blocker sns. J. Mol. Biol., 342, 1367–1377. Croce J., Lhomond G., Gache C. (2003) Coquillette, a sea urchin T-box gene of the Tbx2 subfamily, is expressed asymmetrically along the oral–aboral axis of the embryo and is involved in skeletogenesis. Mech. Dev., 120, 561–572. Ettensohn C. A., Illies M. R., Oliveri P., De Jong D. L. (2003) Alx1, a member of the Cart1/Alx3/Alx4 subfamily of paired-class homeodomain proteins, is an essential component of the gene network controlling skeletogenic fate specification in the sea urchin embryo. Development, 130, 2917–2928. Puchi M., Quin ˜ ones K., Concha C., Iribarren C., Bustos P., Morin V., Genevie`re A. M., Imschenetzky M. (2006) Microinjection of an antibody against the cysteine-protease involved in male chromatin remodeling blocks the development of sea urchin embryos at the initial cell cycle. J. Cell. Bioch., 98, 335–342. Salaun P., Boulben S., Mulner-Lorillon O., Belle R., Sonenberg N., Morales J., Cormier P. (2005) Embryonic-stage-dependent changes in the level of eIF4E-binding proteins during early development of sea urchin embryos. J. Cell. Sci., 118, 1385–1394.
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22. FitzHarris G., Larman M., Richards C., Carroll J. (2005) An increase in [Ca2+]i is sufficient but not necessary for driving mitosis in early mouse embryos. J. Cell. Sci., 118, 4563–4575. 23. Peterson R. E., McClay D. R. (2005) A fringe-modified notch signal affects specification of mesoderm and endoderm in the sea urchin embryo. Dev. Biol., 282, 126–137. 24. Duboc V., R¨ottinger E., Besnardeau L., Lepage T. (2004) Nodal and BMP2/4 signaling organizes the oral–aboral axis of the sea urchin embryo. Dev. Cell., 6, 397–410. 25. Kenny A. P., Oleksyn D. W., Newman L. A., Angerer R. C., Angerer L. M. (2003) Tight regulation of SpSoxB factors is required for patterning and morphogenesis in sea urchin embryos. Dev. Biol., 261, 412–425. 26. Amore G., Yavrouian R. G., Peterson K. J., Ransick A., McClay D. R., Davidson E. H. (2003) Spdeadringer, a sea urchin embryo gene required separately in skeletogenic and oral ectoderm gene regulatory networks. Dev. Biol., 261, 55–81. 27. Coffman J. A., Davidson E. H. (2001) Oralaboral axis specification in the sea urchin embryo. Dev. Biol., 230, 18–28. 28. R¨ottinger E., Besnardeau L., Lepage T. (2004) A Raf/MEK/ERK signaling pathway is required for development of the sea urchin embryo micromere lineage through phosphorylation of the transcription factor Ets. Development, 131, 1075–1087. 29. Gross J. M., McClay D. R. (2001) The role of Brachyury (T) during gastrulation movements in the sea urchin Lytechinus variegatus. Dev. Biol., 239, 132–147. 30. Rupp R. A. W., Snider L., Weintraub H. (1994) Xenopus embryos regulate the nuclear localization of XMyoD. Gene Dev., 8, 1311–1323. 31. Turner D. L., Weintraub H. (1994) Expression of achaete-scute homolog 3 in Xenopus embryos converts ectodermal cells to a neural fate. Gene Dev., 8, 1434–1447. 32. Range R. C., Venuti J. M., McClay D. R. (2005) LvGroucho and nuclear b-catenin functionally compete for Tcf binding to influence activation of the endomesoderm gene regulatory network in the sea urchin embryo. Dev. Biol., 279, 252–267. 33. Tan H., Ransick A., Wu H., Dobias S., Liu Y. H., Maxson R. (1998) Disruption of primary mesenchyme cell patterning by misregulated
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Differential stability of expression of similarly specified endogenous and exogenous genes in the sea urchin embryo. Development, 113, 385–398. 39. McMahon A. P., Flytzanis C. N., HoughEvans B. R., Katula K. S., Britten R. J., Davidson E. H. (1985) Introduction of cloned DNA into sea urchin egg cytoplasm: replication and persistence during embryogenesis. Dev. Biol., 108, 420–430. 40. Ghiglione C., Emily-Fenouil F., Lhomond G., Gache C. (1997) Organization of the proximal promoter of the hatching-enzyme gene, the earliest zygotic gene expressed in the sea urchin embryo. Eur. J. Biochem., 250 (2), 502–513. 41. Arnone M. I., Bogarad L. D., Collazo A., Kirchhamer C. V., Cameron R. A., Rast J. P., Gregorians A., Davidson E. H. (1997) Green fluorescent protein in the sea urchin: new experimental approaches to transcriptional regulatory analysis in embryos and larvae. Development, 124, 4649–4659.
Chapter 14 Exploring the Cytoskeleton During Intracytoplasmic Sperm Injection in Humans Vanesa Y. Rawe and He´ctor Chemes Abstract Understanding the cellular events during fertilization in mammals is a major challenge that can contribute to the improvement of future infertility treatments in humans and reproductive performance in farm animals. Of special interest is the role of the oocyte and sperm cytoskeleton during the initial interaction between gametes. The aim of this chapter is to describe methods for studying cytoskeletal features during in vitro fertilization after intracytoplasmic sperm injection (ICSI) in humans. The following protocols will provide a detailed description of how to perform immunodetection and imaging of human eggs, zygotes, and sperm by fluorescence (confocal and epifluorescence) and electron microscopy. Key words: ICSI, infertility, oocytes, zygotes, sperm, immunofluorescence, immunocytochemistryl, cytoskeleton, electron microscopy, sperm pathology, fertilization, fertilization failure.
1. Introduction The interaction of mammalian spermatozoa with the oocyte after gamete fusion is a complex and meticulously orchestrated cascade of events. While intracytoplasmic sperm injection (ICSI) is an efficient treatment for male infertility (1), there are a significant number of clinical cases of fertilization failure that remain unclear (2,3). The cytoskeleton has diverse functions during fertilization (4). The mouse oocyte and zygote is the most useful and popular model for studying in vitro gamete development and fertilization events but there are huge differences in terms of cytoskeletal organization compared with humans (5). During human fertilization, the microtubules, but not the actin filaments (rodents), are David J. Carroll (ed.), Microinjection: Methods and Applications, Vol. 518 Ó 2009 Humana Press, a part of Springer ScienceþBusiness Media, LLC DOI 10.1007/978-1-59745-202-1_14
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required for pronuclear migration and apposition as well as the formation of the first mitotic spindle. Additionally, the centrosome degenerates in the oocyte and is retained in the sperm during the maturation process. Contrarily to rodents, human spermatozoa contribute the centrosome during fertilization (6), which serves as the dominant microtubule-organizing center in the zygote, and nucleates microtubules from a central structure known as the g-tubulin ring complex. In summary, attention must be taken to avoid extrapolation between animal (rodents) models and humans. The direct study of the internal organization of the human egg and sperm cytoskeleton and the topographical view of the whole flagellum provides new knowledge in the improvement of infertility treatments in humans. Despite extensive research in the area of human reproductive biology, much is still to be understood at the cellular level of how eggs control the assembly and disassembly of sperm’s components. Understanding the role of the cytoskeleton during ICSI is an important step to improve fertilization. The aim of this chapter is to show methods for studying cytoskeletal features during in vitro fertilization after ICSI among humans. The following protocols are going to provide a detailed description of how to perform immunodetection and imaging of human egg, zygote, and sperm components by fluorescence (confocal and epifluorescence) and electron microscopy.
2. Materials 2.1. Media/Chemicals
1. Hepes-Human Tubal Fluid (H-HTF, Irvine Scientific 90126) 2. Synthetic serum substitute (SSS, Irvine Scientific 99193) 3. Bovine serum albumin (BSA, Sigma A4503) 4. Phosphate-buffered saline (PBS, Sigma P3813): in 1l of ultrapure water, pH 7.4. 5. Formaldehyde (formaline 37% solution, Sigma F1635) 6. Taxol (plaquitaxel, Sigma T1912) 7. Triton X-100 (Sigma X100) 8. Poly-L-lysine (MW 300,000; Sigma P8920) 9. Hyaluronidase (Sigma H3884) 10. Tyrodes acid (Sigma T1788) 11. Blocking solution for oocytes and zygotes: 17.5 ml PBS – Sigma P-3813; 0.5 g BSA (fraction V) – Sigma A-3311; 0.5
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ml FBS; 0.1875 g Glycine – Sigma G-7403; 2.5 ml Triton X100 – Sigma X-100; 7.5 ml PBS – Sigma P-3813. Filter with Millipore 0.2 mm (Corning 431219). Store at 4°C. 12. Solution I (Formaldehyde fixation): 400 ml Milli-Q Water; 15.12 g Pipes – Sigma P-8203. Solution will be clear once the pH reaches 7.0. Then add 0.5 g MgCl6H2O – Sigma M2670; 0.48 g EGTA – Sigma E-3889. Adjust volume to 500 ml with Milli-Q Water and filter with Millipore 0.2 mm. Store at 4°C and keep for 3 months. 13. Solution II (prepare the same day of FA fixation): 11.64 ml of Solution I, 0.67 ml of 37% Formaldehyde solution – Sigma F1635; 0.0625 ml Triton X-100 – Sigma X-100; 0.125 ml of stock solution of Paclitaxel (1 mM in DMSO) – Sigma T-1912. 14. Buffer M (all solutions for methanol fixation must be prepared in glassware): 250 ml Milli-Q Water; 125 ml Glycerol – Amresco P-0085440; 1.864 g KCl – Sigma P-5405; 0.051 g MgCl26H2O – Sigma M-2670; 1.702 g Imidazole – Sigma I-3386; 250 ml EDTA Solution (200 mM) – Sigma E-4884; 2.5 ml EGTA Solution (200 mM) – Sigma E-3889. Adjust volume to 500 ml with Milli-Q Water, pH 6.8. Store at 4°C and keep for 6 months. 15. Solution A (Prepare the same day of methanol fixation in glassware): 89 ml solution B, 10 ml methanol, 1 ml Triton X-100 – Sigma X-100. 16. Solution B (Prepare the same day of methanol fixation in glassware): 250 ml Buffer M, 2.5 ml Triton X-100 – Sigma X-100; 0.1755 ml b-Mercaptoethanol – Sigma M-7522; 0.5 ml PMSF (100 mM) – Sigma P-7626. 17. Talp-Hepes Ca2+ free, BSA free (To use for methanol fixation): 2.58 ml KCl (310 mM) – Sigma P-5405; 28.5 ml NaCl (1 M) – Sigma M-2670; 3.44 ml NaH2PO4H2O (29 mM); 0.465 ml Lactic Acid – Sigma L-4263; 2 ml NaHCO3 (250 mM) – Sigma S-1554; 2.5 ml Hepes (1 M) – Sigma H0763; 0.25 ml Penicillin (100.000 IU/ml) – (Pen/Strep) Sigma P-3539; 50 ml Embryo tested water – Sigma W-1503; 3.125 ml MgCl26H2O – Sigma M-2670; 4 mg Phenol red – Sigma P-5530; 250 ml Gentamycin – Sigma G-1397. Adjust volume to 250 ml with embryo-tested water. Filter with Millipore 0.2 mm (Corning 431219), pH 7.4, Osmolarity 255–270 mOsm. Store at 4°C and keep for 3 months. 18. 0.1 M Phosphate buffer: 90 ml of 0.2 M dibasic Na or K phosphate + 10 ml of 0.2 M monobasic phosphate. Complete to 200 ml with distilled water. Correct pH to 7.4 with 0.2 M monobasic phosphate. Use to prepare glutaraldehyde and osmium solutions and for rinses between fixatives.
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19. 8% Glutaraldehyde (EM grade, Pelco 18421, glass ampoules sealed with inert gas). 20. Osmium tetroxide in distilled water (Pelco 18451). Osmium comes as crystals in sealed ampoules. Immerse the ampoule in 50–60°C tap water until crystals melt completely. Take the ampoule out of the hot water and rotate it until the liquid osmium solidifies again as a thin layer on the inner face of the glass ampoule. Place the ampoule within a thick-walled glass container, close and shake vigorously until the ampoule breaks. Add distilled water to make a 2% STOCK solution. Close the bottle and allow to stay overnight at room temperature. Keep well sealed at 4°C. To prepare fixative mix 2 parts phosphate buffer with 1 part osmium 2% solution (see Note 1). 21. Ethanol (50%, 70%, 96%, and 100%) 22. Propylene Oxide, EM grade (Pelco 18601) 23. Eponate 12 – Araldite 502 resin embedding kit (Pelco 18028) 24. Lead Citrate solution (to stain EM sections) For each 25 ml of distilled water use 100 mg Na (OH) and 50 mg lead citrate. Weigh 1 pearl of Na (OH) and adapt all other quantities accordingly. Distilled water should be previously boiled (to eliminate CO2) and allowed to cool to room temperature. Add the NaOH to the appropriate amount of water and then dissolve the lead citrate. Avoid breathing over the solution. Keep the solutions in syringes eliminating all the air. 25. Uranyl acetate solution (to stain EM sections) Prepare a saturated solution of uranyl acetate in distilled water. Keep well sealed at 4°C. To use, mix equal amounts of uranyl solution and absolute acetone. 26. LR white resin, medium grade (Pelco 18181) 27. Blocking buffer (TBS blocking for electron microscopy): 50 ml of 40 mM Trizma hydrochloride (pH 7.4) + 50 ml of 450 mM NaCl. Add normal goat serum to 1% concentration. 28. TBS (Tris buffer saline, 50 mM Tris, 150 mM NaCl): 50 ml of 100 mM Trizma hydrochloride + 50 ml of 300 mM NaCl. Adjust pH to 7.4. 29. Na citrate buffer (for microwave antigen retrieval): 10 mM Citric acid. Correct pH to 6 with NaOH 2.2. Antibodies and Counterstainings
1. Primary antibodies : anti-alpha and beta Tubulin (Cytoskeleton, cat. Number: ATN02), anti p150Glued dynactin (BD Transduction Laboratories, cat. number: 610474), anti intermediate filaments (Vimentin, Santa Cruz Biotechnology, cat. number: sc-7557), anti F-actin (phalloidin-568,
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Molecular probes), anti Arp 2–3 complex (BD Transduction Laboratories, cat. number: 612134), anti Profilin (Immunoglobe, cat. number 0022-01), anti acetylated alpha Tubulin (Sigma T-6793), anti AKAP4 (kindly provided by Dr. Mitch Eddy). 2. Secondary antibodies: anti-mouse, anti-rabbit IgGs conjugated with FITC or TRITC. Goat anti-rabbit IgG, Colloidal gold labeled (15 nm), Pelco 15727. 3. VectaShield with DAPI (H1200, Vector Laboratories), Hoechst 33342 (Sigma-Aldrich) at 5 mg/ml or TOTO-3 (Molecular Probes T3604) at 10 mg/ml, depending on the microscope to be used (epifluorescence with DAPI or Hoechst or confocal with TOTO-3). 2.3. Consumables and Disposables
1. Falcon dishes (Falcon 3037, 60 15 mm) 2. Six-wells cell culture cluster, flat bottom (Corning 3516) 3. Falcon X plates (Falcon 351009) 4. Kodak Technical Pan EM film 5. 300-mesh nickel grids 6. Humid chamber 7. Microscopy slides 8. Microscopy coverslips (18 18 mm) 9. 15 ml conical centrifuge tubes (Corning 430055) 10. 1.5 ml eppendorf tubes 11. Nail polish 12. Lint-free lens paper
2.4. Equipment
1. Slide warmer 2. Dissecting microscope 3. Confocal and epifluorescence microscope 4. Zeis EM 109 Electron Microscope 5. 10 ml, 20 ml, 200 ml, and 1000 ml pipetters (Gilson or similar) 6. Pipetter’s and strippers tips 7. Timers 8. Incubator (37°C, 5% CO2)
2.5. Tools
1. Scalpels and blades 2. Fine tweezers 3. Strippers 4. Sterile glassware and funnels
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3. Methods Injected oocytes and zygotes shown in this chapter are discarded human material obtained from couples undergoing ICSI at CEGyR who gave informed consent in writing. Internal Review Board approved the use of the mentioned material. The methods described below outline the study of human non-fertilized oocytes and zygotes by immunocytochemistry (ICC) and fluorescence microscopy during ICSI. Formaldehyde and methanol fixation are included, and the visualization of human sperm pathology through epifluorescence and electron microscopy. 3.1. Study of Human Non-fertilized Oocytes and Zygotes by ICC and Fluorescence Microscopy During ICSI
Human oocytes and zygotes are frequently very few and are handled in drops using the mouth pipette (when allowed and depending on the regulations of every Research Institute) or strippers. For ICC, all the material needs to be completely denuded from cumulus cells and zona pellucida and then fixed with formaldehyde or methanol.
3.1.1. Removal of Cumulus Cells
1. Take discarded human oocytes and zygotes and remove cumulus cells with short incubations (approximately 40 s) in a volume of human tubal fluid supplemented with Hepes (HHTF) containing 80 IU/ml of hyaluronidase. 2. Use the stripper with the appropriate capillary diameter (135 mm or 150 mm). 3. Once the human material is stripped, place the oocytes or zygotes in five-seven 50 ml drops of 0.3%BSA +H-HTF for washing.
3.1.2. Removal of Zona Pellucida
1. The zona pellucida (ZP) is removed after incubation in a drop of Tyrodes acid (pH 2.3) (Fig. 14.1a). 2. For this purpose, place human cumulus cells-free oocytes and zygotes in a 30 ml drop of Tyrodes acid for 15 s. After removal of the ZP, wash denuded human material 2–3 times in 35 ml drops of 0.3%BSA +H-HTF or blocking solution. 3. At this point, the material is ready to be handled for fixation and permeabilization with 2% formaldehyde (FA) and 0.5% Triton X-100, respectively (Method modified and based on Messinger and Albertini, 1991 (7) (see Note 2).
3.1.3. Formaldehyde Fixation, Permeabilization, and Antibody Labeling
1. Place the samples in a one-well dish containing 500 ml of Solution II during 30 min at 37°C. 2. After the fixation and permeabilization step, take the oocytes and wash them by transferring into different drops of blocking
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Fig. 14.1. Preparation of slides for immunocytochemistry. (a) The zona pellucida (ZP) is removed after incubation in a drop of Tyrodes Acid (pH 2.3). (b) Oocytes are maintained in a drop of blocking solution for 1 h at RT under oil or at 4°C until ICC is performed. (c) An 18 18 mm coverslip is used to mount the sample. (d) Seal all four sides of the coverslip with clear nail polish (available at local drug stores) to prevent drying.
solution. Keep them in a drop of blocking solution for 1 h at RT under oil or at 4°C until ICC is performed (Fig. 14.1b). 3. An alternative to formaldehyde fixation is 100% methanol fixation for 10 min at –20°C. After that, coverslips are stored in PBS and 0.1% Triton X-100. Permeabilization is not necessary. 4. Antibodies are applied by placing the oocytes in 30–50 ml drops of the appropriate dilution of the antibody (experimentally determined) for 40 min at RT or 1 h at 37°C under oil (raising temperature to 37°C may improve antibody binding to epitopes), or overnight at 4°C. 5. After primary antibody incubation, samples are then rinsed by transferring the oocytes into drops of blocking solution (see Note 3). 6. To detect the primary antibodies, fluorochrome-conjugated secondary antibodies are applied in drops for 1 h at RT in the dark. 7. Following incubation with secondary antibodies, samples are once again rinsed with drops of blocking solution. 8. For DNA labeling and visualization at the confocal microscope, incubate the material for 25 min at RT in TOTO-3 at 10 mg/ml. For epifluorescence, DNA is counterstained using
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Vectashield with DAPI or Hoechst 33342 at 5 mg/ml (25 min at RT). The stock of Hoechst can be kept for several months at 4°C, shielded from light by wrapping in aluminum foil, and used as needed. 3.1.4. Mounting and Visualization of Formaldehyde-Fixed Samples
1. A 5 ml drop of the anti-fade medium with the oocytes is placed in a slide. An 18 18 mm coverslip is used to mount the sample (Fig. 14.1c). 2. Seal all four sides of the coverslip with clear nail polish (available at local drug stores) to prevent drying (Fig. 14.1d). 3. Fixed oocytes and zygotes shown in this chapter were imaged with Olympus spectral confocal microscope, using laser lines at 488 nm, 568 nm, and 633 nm wavelengths (University of Buenos Aires, Faculty of Exact and Natural Science). 4. Epifluorescence images were taken with an Olympus BX40 microscope associated to a UV light source.
3.1.5. Methanol Fixation and Antibody Labeling (Based on Simerly and Schatten, 1993 (8) )
For methanol fixation it is mandatory to work in a Warm Room in order to preserve the oocyte/zygote’s cytoskeleton structure (Fig. 14.2a). Microtubules are very sensitive to temperature changes, so working on a slide warmer and a warm room is recommended (Fig. 14.2b). 1. Cumulus cell and ZP removal is done as stated before. 2. Prepare a six-wells culture dish by placing an 18 18 mm poly-L-lysine coverslip in each well. Add 4 ml of Talp Hepes Ca2+-free, BSA-free (Fig. 14.2b). Take the oocytes and zygotes (ZP free) and gently place them in each well. Oocytes must decant from the media surface to the bottom of the well (where the coverslip is placed) in order to ‘‘wash’’ them from any protein left in its surface (Fig. 14.2c). 3. Remove 2 ml of Talp Hepes Ca2+-free, BSA-free and add (very gently) 9 ml of Solution A (Fig. 14.2d). 4. After approximately 10 min, oocytes and zygotes become clear (signal that they have been extracted). Using a mouth pipette, gently remove oocytes and zygotes and place them in an 18 18 mm of a poly-L-lysine coverslip seated in a quarter of an X plate (Fig. 14.2e). Suck out the excess liquid (without drying them!) and let them attach to the coverslip. Add 9 ml of Solution B until the quarter is filled (Fig. 14.2e). 5. Progressively add 100% methanol (kept at –20°C) to start dehydratating the oocytes until the solution is fully covered with 100% methanol (Fig. 14.2f). Gradually pour off the solution B towards to other three-quarters of the X plate.
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Fig. 14.2. Methanol fixation of oocytes and zygotes. (a) Warm room. (b) A slide warmer is recommended when studying microtubules structures. The material is placed in a six-well culture dish on 18 18 mm poly-L-lysine coverslips (each well) covered with Talp Hepes Ca2+-free, BSA-free. (c) Oocytes decant from the media surface to the bottom of the well. (d) Talp Hepes Ca2+-free, BSA-free is removed and Solution A is added. (e) Removal of oocytes and zygotes to place them in an 18 18 mm of a poly-L-lysine coverslip seated in a quarter of an X plate. After that, solution B is added until the quarter is filled. (f) Addition of 100% methanol (–20°C) until it fully covers the material. Gradually pour off the solution B towards other three-quarters of the X plate (hydratation step). (g) Primary and secondary antibodies are applied on the coverslip containing the oocytes and zygotes and using the edges of an X plate. (h) Coverslips are mounted in an anti-fade medium to retard photobleaching. (i) Sealing all four sides of the coverslip with clear nail polish to prevent drying.
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6. After oocytes and zygotes are completely immersed in 100% methanol (fixation step), the hydratation step begins. For this purpose, gradually add 10 mM PBS + 0.1% Triton X100 and pour off the methanol towards other three-quarters of the X plate (as before) until it is fully covered. Methanolfixed oocytes can be kept at 4°C for a week until ICC is performed. 7. Primary antibodies are applied on the coverslip containing the oocytes and zygotes and using the edges of an X plate (Fig. 14.2 g). Place approximately 200 ml of the appropriate dilution of the primary antibody (experimentally determined) for 40 min at or overnight at 4°C. Alternatively, the quarters of the X plate can be filled with water (humified chamber), covered with the lid, and the incubation can take place for 1 h at 37°C. 8. After primary antibody incubation, samples are rinsed by removing the primary antibody dilution and adding 200 ml of PBS+ 0.1% Triton X-100. 9. To detect the primary antibodies, fluorochrome-conjugated secondary antibodies are applied on the coverslip for 1 h at RT in the dark. 10. Following incubation with secondary antibodies, samples are once again rinsed by adding 200 ml of PBS+ 0.1% Triton X-100. 11. DNA labeling is done as stated before (formaldehyde fixation). 3.1.6. Mounting and Visualization
1. Coverslips are mounted in an anti-fade medium to retard photobleaching (as above). A 5 ml drop of the anti-fade medium is placed in a slide and the coverslip (with the processed material attached) is mounted on it (Fig. 14.2 h). 2. Seal all four sides of the coverslip with clear nail polish to prevent drying (Fig. 14.2i). 3. Examples of the visualization of cytoskeleton dynamics during fertilization and fertilization failure after ICSI are seen in Figs. 14.3 and 14.4 respectively.
3.2. Visualization of Human Sperm Pathology
Sperm Pathology, the discipline of characterizing structural and functional deficiencies in abnormal spermatozoa, allows a deep understanding of the factors responsible for deterioration of sperm quality. In particular, abnormally shaped flagella in severely asthenozoospermic men express different alterations in the structural and molecular organization of the sperm tail. One example of this is the dysplasia of the fibrous sheath (DFS, see Fig. 14.5) a developmental disruption of the tail cytoskeleton, involving most prominently, but not exclusively, the fibrous sheath (FS) of the sperm principal piece (9,10). The FS, a cytoskeletal structure of
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Fig. 14.3. Cytoskeleton dynamics during normal fertilization. (a) Microtubules are widely spread throughout the oocyte cytoplasm when fully apposed and developed of male and female pronuclei are seen (approximately 18 h after ICSI). Tubulin is nucleated to the microtubule organizing center (MTOC), and through the action of microtubule-related proteins like Dynactin (see b) pronuclear apposition occurs. (b) During pronuclear apposition in humans, Dynactin is visualized associated with male and female pronuclear envelopes and concentrated in the center of the zygote (ref. (14)). (c) Intermediate filaments (Vimentin) are strongly seen as a punctuate pattern within all the zygote’s cytoplasm and also associated with both pronuclei. (d) Filamentous actin (F-actin) is observed in the zygote’s cortex as well as a central area within the cytoplasm where pronuclear apposition is taking place. (e) The actin-related protein Arp 2/3 is visualized throughout the cytoplasm and clearly seen associated with both pronuclei participating in what is called the ‘nucleoskeleton’. Arp 2/3 complex stimulates the formation of actin filaments but its specific role during fertilization in humans is unknown. (f) Profilin is a protein that regulates actin polymerization by sequestering actin monomers in association with other actin-related proteins. In pronucleate-stage zygotes, profilin localizes to specific foci inside both pronuclei and the cytoplasm. Actin remodeling is essential for fertilization and embryo development and inhibition of profilin leads with a failure during embryo cytokinesis (15). Normal human zygotes (Fig. 14.3) and supernumerary zygotes that failed to cleave within 40 h after ICSI (Fig. 14.4) were not used for infertility treatment or cryopreservation and were donated for research by consenting adult female donors. All animal and human procedures were approved by CEGyR’s Internal Review Board and Ethic Committee accordingly.
the sperm flagellum, is an assemblage of proteins like AKAP4, sperm thyoredoxins (Sptrx-1 and 2), and glyceraldehyde 3-phosphate dehydrogenase-S (GAPDS) all involved in various essential functions such as cAMP-dependent signaling pathways and protein phosphorylation, disulphide bond reduction, and glycolytic ATP generation (11–13). 3.2.1. ICC and Epifluorescence 3.2.1.1. Fixation and Permeabilization of the Semen Sample
1. After 30 min of ejaculation, take the fresh semen sample and wash it by centrifugation in PBS for 5 min. 2. Remove supernatant and dilute the pellet in a volume of PBS. 3. Place dry 18 18 mm poly-L-lysine coverslips on the edge of a slide warmer (at 37°C). Deposit 100 ml of cell suspension onto each slide and incubate for 15 min to allow the spermatozoa to settle onto the poly-l-lysine layer.
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Fig. 14.4. Cytoskeletal organization and DNA configuration after fertilization failure during ICSI. In all pictures, inserts represent a schematic situation for each selected example. (a and b) Oocyte’s meiotic spindle with the chromosomes in the metaphase plate (human oocytes are arrested at metaphase of the second meiosis before fertilization). In some cases, the sperm can be expulsed and fertilization does not take place. (b) An abnormal distribution of chromosomes can be seen in the meiotic spindle of an old oocyte. (c) A premature chromosome condensation (PCC). In this case, the oocyte meiotic spindle is seen next to the paternal chromosomes associated with the sperm tail. (d) The sperm head failed to form the male pronucleus (arrow) and the female chromosomes are still condensed. (c and d) Lack of activation of the oocyte and presence of an abnormal distribution of actin in d (arrow in c: sperm tail, arrow in d: male pronucleus). (e and f) Two examples of oocyte activation with a failure or incomplete male and/or female pronuclear formation. (e) Microtubules were not homogeneously distributed around pronuclei. (f) Male DNA was not completely decondensed (arrow) while female pronucleus was (asynchronic pronuclear development). Actin was normally distributed in the cytoplasm and cortical region (g and h) Arrest at the first mitotic plate (first embryo mitosis). (g) Disorganization of chromosomes and microtubules can be visualized associated with the sperm tail. (h) Dissociation of the sperm tail-head can be observed within the oocyte cytoplasm.
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Fig. 14.5. Longitudinal and transverse sections of sperm flagella depicting redundant fibrous sheath in patients with dysplasia of the fibrous sheath. (A and C) Regular transmission electron microscopy. Excessive fibrous sheath elements surround the central axoneme. (B, D, and E) Corresponding pictures showing positive immunogold localization for AKAP4. Gold particles overlay exclusively the fibrous sheath but not the central axoneme (asterisks). F. Specificity of AKAP4 localization is ascertained by the sparsity of gold particles in a negative control section. Scale bars represent 0.2 mm. Panels D, E, and F were originally published in Chemes et al. (2001) J. Androl., 22, 302–315 by permission.
4. Shake off the medium with excess cells and submerge the coverslip in 2% FA in PBS (pH 7.2–7.3) for 40 min. Best preservation of cell structure is achieved when the coverslips with cells are carefully transferred, coated face up, into six-wells dish with PBS+2%FA+ 1%Triton X-100. (To prepare 12 wells mix 37.44 ml PBS+ 2.16 ml FA (2%)+ 0.4 ml Triton X-100 (1%)). 5. Remove the coverslips with cells from formaldehyde solution using fine-pointed-tweezers, wash briefly in PBS, and store in pure PBS or in PBS containing 1% Triton X-100 (permeabilization agent).
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3.2.1.2. ICC and Visualization of Spermatozoa
1. Place coverslips on the cross of a four-well petri dish (see Fig. 14.2 g). Block non-specific antibody binding to residual free aldehyde groups by 40 min incubation in blocking solution. 2. Incubations with primary and secondary antibodies are carried out following the methods previously described for methanol fixation of oocytes and zygotes (see Section 3.1.4).
3.2.2. Electron Microscopy 3.2.2.1. Fixation and Processing of the Semen Sample for Regular Electron Microscopy
1. Semen samples were studied within 30–45 min of ejaculation after complete liquefaction. Samples were diluted 1:4 with phosphate buffer (0.1 M, pH 7.4) at room temperature and thoroughly mixed until a homogeneous suspension is obtained (see Note 4). 2. When a uniform and thin sperm suspension is obtained it is transferred to a conical tip tube cell and centrifuged at 1500–2000 rpm for 10 min. Ideally, a well-defined pellet should be obtained. If this is not the case, discard supernatant, resuspend the pellet in 10 ml phosphate buffer, and re-centrifuge (see Note 5). 3. Fixation is accomplished by slowly adding 3% glutaraldehyde in phosphate buffer at 4°C through the tube wall. If the pellet is more than 2 mm thick it should be carefully dislodged from the bottom of the tube so that the fixative reaches both faces of the pellet. Fixation time varies between 3 and 5 h, followed by two rinses of 30 min each in phosphate buffer. Secondary fixation is accomplished with 1.3% osmium tetroxide at 4°C for 2 h and subsequent two 30-min rinses in phosphate buffer. 4. Dehydrate in an ascending series of ethanol (50%, 70%, 96%) in the cold. Perform three rinses of 10 min for each ethanol concentration. The last 96% rinse should be performed at room temperature. 5. Place in 100% ethanol (3 times of 20 min each), followed by three rinses in propylene oxide (20 min each). These two steps should be performed at room temperature. 6. Place in a 1:1 solution of propylene oxide: Eponate–Araldite mix and let infiltrate for 2 h at room temperature. 7. Change for fresh Eponate-Araldite mix and cure in 60°C oven for 24–48 h in appropriate molds. 8. Eponate–Araldite mix: Eponate 12 Resin (10 parts) + Araldite 502 Resin (10 parts) + DDSA (30 parts). 9. Mix well. The mixture can be moderately heated to facilitate mixing. Keep at 4°C. 10. Before using add BDMA at 3.2% final concentration.
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11. Thin sections (silver to pale golden) were obtained in a RMC MT-7000 Ultramicrotome using a diamond knife and examined and photographed in a Zeiss 109 Electron microscope (Zeiss Oberkochen, Germany) after double-staining with uranyl acetate and lead citrate (see Note 6).
3.2.2.2. Ultrastructural Immunocytochemistry of Cytoskeletal Components
Initial processing and centrifugation of the semen sample is similar to that for regular electron microscopy. 1. For ultrastructural immunocytochemistry sperm pellets were fixed for 1 h at 4°C in 5% phosphate-buffered formaldehyde (0.1 M, pH 7.4), rinsed in buffer, dehydrated in an increasing series of ethanol, infiltrated in LR-White Resin, medium grade (London Resin Co., Ltd, Reading, England) and polymerized at 60°C for 24 h (see Note 7). 2. Thin sections displaying pale gold to silver interference colors were obtained in a RMC MT-7000 Ultramicrotome with a diamond knife, mounted on 300-mesh nickel grids and dried at room temperature. 3. Ultrastructural immunocytochemical localization of selected cytoskeletal components (AKAP4 in our figures) was performed on section with or without pretreatment with microwaves. 4. The grids were hydrated by flotation on distilled water drops. 5. After antigen retrieval (see Note 8), grids were subsequently washed and incubated for 30 min at room temperature in blocking buffer (Tris-buffered saline: TBS 225 mM, pH 7.5 + 10% normal goat serum) and then floated on drops of primary antibody and incubated on a humid chamber overnight at 4°C. Primary antibodies were used at appropriate dilutions. 6. After three washes in TBS the grids were incubated for 1 h at 4°C with blocking buffer containing 15 nm colloidal gold-labeled goat anti-rabbit IgG (Pelco International, Redding, Ca) at 1:25 or 1:50 dilutions, and rinsed three times in TBS. 7. Grids were subsequently counterstained with 1% osmium tetroxide followed by 1:1 aqueous uranyl acetate: absolute acetone or were left without any further staining. 8. Specimens were studied and photographed in a Zeiss EM109 transmission Electron Microscope (EM, Zeiss, Oberkochen, Germany). Negative controls were processed identically replacing the primary antibody by similar dilutions of primary antibodies preadsorbed with excess antigen or omitting the first antibody step.
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4. Notes 1. Osmium is very volatile. Avoid contact with skin, eyes, and do not inhale vapors. Always use under hood. Dispose of used solutions with toxic materials. 2. Aldehydes are carcinogenic, thus during ICC, fixations must be performed under a fume hood or with a facemask with special filters. It is crucial to always keep the coverslip with the cell-coated side up. 3. In order to better visualize internal cellular structures, for immunocytochemistry of oocytes and zygotes, no more than a week of incubation in blocking solution is recommended before incubation with primary and secondary antibodies. 4. During processing of the human semen samples, when semen is very thick this step requires more thorough mixing or the use of repeated aspirations through a glass Pasteur pipette or even a disposable 10 ml syringe fitted with a 21G needle. In these cases a higher than 1:4 dilution may be needed. 5. Care should be exercised not to resuspend the pellet. This is accomplished by adding all solutions drop by drop through the tube wall and discarding supernatants by slowly pouring them. Avoid use of pipettes at these steps or any other device that may create liquid turbulences and resuspend the pellet. 6. To stain EM sections place a square piece of parafilm under a petri dish. Mix equal amounts (1 or 2 drops) of uranyl acetate and absolute acetone on the parafilm surface. Float grids face down. Allow to stain for 2 min. Pick up grids with fine forceps and rinse well in distilled water. Without drying, transfer grids to a few drops of lead citrate and float them face down. Stain for 2 min. Rinse well with distilled water and allow grids to dry face up on filter paper. 7. LR white mix (embedment for use on sectioned ultrastructural immunocytochemistry). Do not use osmium fixation. Tissue fragments or pellets are dehydrated in an ascending series of ethanol (50%, 70%, 96%). It is not necessary to reach 100% ethanol solutions. Infiltrate in LR White by 4–6 changes, 30 min each, at room temperature. Change to fresh LR White and polymerize for 24 h in 60°C oven in molds closed without air bubbles. 8. To perform antigen retrieval the grids were immersed section side up in a glass petri dish containing 10 mM Na citrate buffer pH 6.0 and subjected to 1 min of microwave irradiation at 800 W.
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Acknowledgments Methods described in this chapter are based on protocols developed by V. Rawe at CEGyR and during her postdoctoral training in Dr. Gerald Schatten’s lab. (Pittsburgh Development Center, Pittsburgh, PA, USA). H. Chemes contributions were developed at his laboratory in the CEDIE (National Research Council of Argentina). Special thanks to Gerald Schatten, Calvin Simerly, and Peter Sutovsky for continuous scientific support and fruitful collaborations. Part of the information presented in this chapter is the subject of the ‘‘Summer Course in Biology of Reproduction’’ annually organized at CEGyR (for general information:
[email protected]/
[email protected]). Supported by CEGyR Foundation and Grants from CONICET (PIP 2565), ANPCyT (PICT 9591). References 1. Palermo, G., Joris, H., Devroey, P., and Van Steirteghem, A.C. (1992) Pregnancies after intracytoplasmic injection of single spermatozoon into an oocyte. Lancet 340, 17–8. 2. Asch, R., Simerly, C., Ord, T., Ord, V.A., and Schatten, G. (1995) The stages at which human fertilization arrests: microtubule and chromosome configurations in inseminated oocytes which failed to complete fertilization and development in humans. Hum Reprod. 10, 1897–906. 3. Rawe, V.Y., Brugo Olmedo, S., Nodar, F.N., Doncel, G.D., Acosta, A.A., and Vitullo, A.D. (2000) Cytoskeletal organization defects and abortive activation in human oocytes after IVF and ICSI failure. Mol. Hum. Reprod. 6, 510–16. 4. Longo, F.J. (1989) Egg cortical architecture. In ‘The Biology of Fertilization’ (H. Schatten and G. Schatten, eds.), pp. 105–58. Academic Press, San Diego. 5. Menezo, Y. and Herubel, F. (2002) Mouse and bovine models for human IVF. RBM Online. 2, 170–75. 6. Schatten, G. (1994) The centrosome and its mode of inheritance: the reduction of the centrosome during gametogenesis and its restoration during fertilization. Dev Biol. 165, 299–335. 7. Messinger, S.M. and Albertini, D.F. (1991) Centrosome and microtubule dynamics
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during meiotic progression in the mouse oocyte. J Cell Sci. 100, 289–98. Simerly, C. and Schatten, G. (1993) Techniques for localization of specific molecules in oocytes and embryos. Methods Enzymol 225, 516–53. Chemes, H.E., Brugo, S., Zanchetti, F., Carrere, C. and Lavieri, J.C. (1987) Dysplasia of the fibrous sheath. An ultrastructural defect of human spermatozoa associated with sperm immotility and primary sterility. Fert Steril. 48, 664–69. Chemes, H.E., Brugo, S., Carrere, C., Oses, R., Carizza, C., Leisner, M., Blaquier, J. (1998) Ultrastructural pathology of the sperm flagellum. Association between flagellar pathology and fertility prognosis in severely asthenozoospermic men. Hum. Reprod. 13, 2521–26. Carrera, A., Gerton, G.L., and Moss, S.B. (1994) The major fibrous sheath polypeptide of mouse sperm: structural and functional similarities to the A-kinase anchoring proteins. Dev Biol. 165, 272–84. Miranda-Vizuete, A., Ljung, J., Damdimopoulos, A.E., Gustafsson, J.A., Oko, R., Pelto-Huikko, M. and Spyrou, G. (2001) Characterization of Sptrx, a novel member of the thioredoxin family specifically expressed in human spermatozoa. J Biol Chem. 276, 31567–74.
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13. Miki, K., Qu, W., Goulding, E.H., Willis, W.D., Bunch, D.O., Strader, L.F., Perreault, S.D., Eddy, E.M., O’Brien, D.A. (2004) Glyceraldehyde 3-phosphate dehydrogenase-S, a sperm-specific glycolytic enzyme, is required for sperm motility and male fertility. Proc. Natl. Acad. Sci. USA 23, 16501–06. 14. Payne, C., Rawe, V.Y., Ramalho-Santos, J., Simerly, C. and Schatten, G. (2003)
Cytoplasmic dynein/dynactin association with nucleoporins and vimentin mediates genomic union during mammalian fertilization. J Cell Sci 116, 4727–38. 15. Rawe, V.Y., Payne, C. and Schatten, G. (2006) Profilin and actin-related proteins regulate microfilament dynamics during early mammalian embryogenesis. Hum. Reprod. 5, 1143–53.
Chapter 15 Somatic Cell Nuclear Transfer in the Mouse Satoshi Kishigami and Teruhiko Wakayama Abstract Somatic cell nuclear transfer (SCNT) has become a unique and powerful tool for epigenetic reprogramming research and gene manipulation in animals since ‘‘Dolly,’’ the first animal cloned from an adult cell was reported in 1997. Although the success rates of somatic cloning have been inefficient and the mechanism of reprogramming is still largely unknown, this technique has been proven to work in more than 10 mammalian species. Among them, the mouse provides the best model for both basic and applied research of somatic cloning because of its abounding genetic resources, rapid sexual maturity and propagation, minimal requirements for housing, etc. This chapter describes a basic protocol for mouse cloning using cumulus cells, the most popular cell type for NT, in which donor nuclei are directly injected into the oocyte using a piezo-actuated micromanipulator. In particular, we focus on a new, more efficient mouse cloning protocol using trichostatin A (TSA), a histone deacetylase (HDAC) inhibitor, which increases both in vitro and in vivo developmental rates from twofold to fivefold. This new method including TSA will be helpful to establish mouse cloning in many laboratories. Key words: Nuclear transfer, somatic cell clone, mouse, trichostatin A, HDAC inhibitor.
1. Introduction The first adult somatic cell cloned animal, a sheep named ‘‘Dolly’’ was reported in 1997 (1). Subsequently, researchers have succeeded in cloning more than 10 mammals using both embryonic and adult donor cells (2). These successes in somatic cell cloning give promise to applications such as gene manipulation, species preservation, livestock propagation, and cell therapy for medical treatment by nuclear transfer embryonic stem (ES) cells (NT-ES cells) (3–6). Mouse cloning by somatic cell nuclear transfer (SCNT) has been inefficient since the first cloned mouse, ‘‘Cumulina,’’ was born in 1997 (7). Many different attempts to improve the low David J. Carroll (ed.), Microinjection: Methods and Applications, Vol. 518 Ó 2009 Humana Press, a part of Springer ScienceþBusiness Media, LLC DOI 10.1007/978-1-59745-202-1_15
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developmental rate of cloned mouse embryos have been made. These include optimizations of the NT protocol (7) such as oocyte activation (8), and timing of enucleation (9). In addition, the simple treatment of cloned embryos with dimethyl sulfoxide (DMSO) (10) to induce DNA demethylation (11) and more laborious methods including serial nuclear transfer (12) and clone–clone aggregation (13) have been also developed to increase the success rate of mouse cloning. However, despite these numerous creative methods, improvement was minimal. Recently, we and the laboratory of Tsunoda have published a new cloning method where embryos are treated with TSA, a histone deacetylase (HDAC) inhibitor, following nuclear transfer (14–16), which leads to 2–5 times higher success rates for both reproductive and therapeutic cloning (Fig. 15.1), suggesting that TSA enhances reprogramming of transferred somatic nuclei in oocytes. It is noted that TSA treatment of normal fertilized embryos perturbs their subsequent embryonic development (17, 18). Most nuclear transfer experiments are done by means of cell fusion, which is widely used for animal cloning experiments (19). The alternative method requires using a donor cell nucleus injected directly into the oocyte. However, mouse oocytes in particular are so fragile that conventional microinjection through pipettes wider than 1 mm at their tips is impracticable and leads to lysis. This dilemma was eventually overcome with the application of piezo-actuated micromanipulation (20), which permitted the use of larger microinjection pipette for intracytoplasmic sperm injection (ICSI). By 1998, a mouse SCNT method that allows for injecting donor nuclei into enucleated oocytes using a piezo unit was successful in producing the first cloned mice (7). Once the piezo unit is properly set up on the micromanipulator, it will be of considerable assistance not only in doing ICSI or NT, but also in other forms of micromanipulation such as ES cell injection into blastocysts (21, 22). Moreover, using the piezo unit simplified pipette preparation, allowing the use of blunt-tip pipettes
Fig. 15.1. TSA treatment of cloned embryos significantly improves success rates of both reproductive and therapeutic cloning (14). Each white or gray bar shows a success rate of cloning without or with TSA, respectively. The success rates are calculated based on the numbers of transferred or cultured embryos, respectively.
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without any additional treatment, whereas without using a piezo unit, the pipette tip must be sharpened to a fine point, requiring lots of skill and time. Now laboratories could reliably produce cloned mice from a variety of cells using the piezo unit. Currently, cumulus cells (7), tail-tip fibroblast cells (23), Sertoli cells (24), fetal cells (25, 26), and ES cells (27) have been routinely used to produce cloned mice. NKT cells (28), primordial germ cells (29), hematopoietic stem cell (30, 31), fetal neuronal cells (32), and newborn neuronal stem cells (33) have also been used. However, limitations to cloning efficiency are mainly posed by the type of nucleus donor cell. For example, the cloning efficiency from ES cells is greater than that from Sertoli cells and progressively less efficient in cumulus cells, fibroblasts, and finally thymocytes. Mouse strain is also restrictive for successful cloning (25, 34). So far, only hybrid mice and the 129 inbred strain have been used successfully as sources of donor nuclei (either somatic or ES cell) or of recipient oocytes. However, the recent TSA-cloning method also allows us to substantially clone even ICR outbred mice (35). Regardless, to first establish a successful mouse cloning system in each laboratory, we recommend researchers begin with cumulus cells as a somatic cell type along with B6D2F1 as a donor and recipient strain following the procedures in this chapter. Even before successful cloning, we have to keep in mind that abnormalities in mice cloned from somatic cells have been reported, such as abnormal gene expression in embryos (36–39), abnormal placentas(23,40),obesity(41),orearlydeath(42).Inparticular,cloned mice have almost 100% penetrance of placentomegaly (43). Such abnormalities notwithstanding, success in generating cloned offspring has opened new avenues to study complex processes, such as genomic reprogramming, imprinting, and embryonic development.
2. Materials 2.1. Equipment
1. Inverted microscope with Hoffman optics (Olympus, model IX71). 2. Micromanipulator set (Narishige, cat. no. MMO-202ND). 3. Pipette puller and pipette (Sutter Instrument, model P-97 and cat. no. B100-75-10). 4. Microforge (Narishige, Model MF-900). 5. Piezo impact drive system (Prime Tech, model PMM-150FU). Speed (S) and Intensity (I) controller settings should be adjusted to 1 (S) and 1 to 3(I), and 5 to 8 (S) and 1 to 3(I), respectively, for two types of piezo impacts, referred here as ‘‘small hammer’’ and ‘‘big hammer’’, respectively.
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2.2. Animals
1. Mature B6D2F1 hybrid mice (C57BL/6 DBA/2) are sacrificed for oocyte and somatic cell donations in this standard cloning experiment (see Note 1). 2. Surrogate females for embryo transfer are prepared by mating ICR (CD-1) females with vasectomized males. Foster females are similarly prepared except using fertile males.
2.3. Media
1. For embryo culture, KSOM medium (Specialty Media, cat. no. MR-020P-5D) is prepared following the provided directions. Then, 1/1000 volume of phenol red solution (Sigma, cat. no. P0290) is added. Sterilize by filtration through a 0.22-mm filter and store in aliquots at 4°C. Chatot-Zimomek-Bavister (CZB) medium (44) supplemented with 5.56-mM D-glucose can also be used as well (7). 2. HEPES-buffered CZB medium (HEPES-CZB) is prepared by modifying CZB medium with 20-mM HEPES-HCl, a reduced amount of NaHCO3 (5 mM), and 0.1 mg/ml polyvinyl alcohol (PVA) instead of BSA (see Note 2). 3. Ca2+-free CZB medium is prepared as CZB without CaCl2. Detailed chemical compositions of HEPES-CZB and Ca2+free CZB media are also described elsewhere (45).
2.4. Reagents
1. Cytochalasin B (CCB; ICN Biomedicals, cat. no.195119) is dissolved at 0.5 mg/ml in DMSO; Sigma, cat. no. D8418). Dispense solution into small aliquots and store at –20°C. 2. Hyaluronidase (Sigma, cat. no. H4272) is dissolved at 100 mg/ml in HEPES-CZB for a 10% stock solution. Dispense solution into small aliquots and store at –20°C. 3. Strontium chloride hexahydrate (Sr2+; Wako, cat. no. 19304182) is dissolved at 100 mM in distilled H2O. Store at room temperature. 4. Polyvinyl pyrrolidone (PVP; Wako, cat. no. 168-17042) is dissolved at 12% (w/v) in HEPES-CZB (see Note 3). Also a commercial 10% PVP solution works well (Irvine Scientific, cat. no. 99311). Sterilize by filtration through a 0.45-mm filter and store in aliquots at 4°C (see Note 4). 5. Trichostatin A (TSA; Sigma, cat. no. T8552) is dissolved in DMSO for a 1-mM stock solution and stored in aliquots at 20°C.
2.5. Making Micromanipulation Pipettes
Micromanipulation for enucleation and injection requires three types of pipettes: holding, enucleation, and injection. Although the inner diameter of the enucleation pipette can be of a variety of widths, to minimize the accidental suction of ooplasm, smaller is better (a suggested inner diameter is 89 mm). The
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inner diameter of the injection pipette should be adjusted depending on the donor cell size such as 5–6 mm for cumulus cell injection (15). 1. Pull out the pipette using a puller with programmed setting for ICSI which have to be optimized depending on the condition of filament (an example of settings is: Heat 860, Pull 30, Velocity 120, and Time 200). 2. Break the tip of the pipette at 90° with the tip blunt using a microforge. 3. Bend the pipette approximately 1–2 mm from the tip using the microforge. The proper angle may depend on the type of micromanipulator (a suggested angle is around 20 degrees). 4. Before the pipette is mounted on the micromanipulator, fill the wider end of the pipette (to approximately 5–10 mm in length) with mercury (Fisher Scientific, cat. no. M-140) using a 26-gauge needle attached to a 1-ml syringe.
3. Methods The somatic cell cloning technique is a complicated process consisting of enucleation and nuclear transfer using a micromanipulator, embryo culture, and embryo transfer (Fig. 15.2). It must be acknowledged the success rates of mouse cloning are inherently labile owing to unique conditions in each laboratory. To obtain successful development of cloned embryos, therefore, it is recommended to carefully confirm the quality of all the reagents, media, and techniques by first producing parthenogenetic and ICSImediated fertilized embryos (see Note 5). After nuclear transfer, cloned embryos can be used for production of cloned mice by way of embryo transfer to surrogate mothers. They can be also used for the establishment of NT-ES cells (6, 15). 3.1. Preparation of Oocytes and Donor Cells (Fig. 15.3)
1. Inject mature females intraperitoneally with 5 IU of PMSG and 5 IU of hCG, with a 48-h period between injections. Normally 3–5 females are necessary for acquisition of more than 100 recipient oocytes. 2. Collect mature oocytes that are surrounded by cumulus cells from the ampullae of the oviducts 15–17 h after the hCG injection. 3. Incubate the collected oocytes in HEPES-CZB medium containing 0.1% hyaluronidase at 37°C until cumulus cells disperse (2–4 min).
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Fig. 15.2. A progressive chart for creation of either delivery of cloned pups or establishment of NT-ES cells. This is based on the ‘‘Honolulu method’’ (7) with a modification for TSA treatment (14,15).
4. Wash the oocytes and place them in a microdrop culture dish containing KSOM medium (see Note 6). 3.2. Enucleating Oocytes (Fig. 15.4A)
1. Place a group of 10–20 oocytes into a drop of HEPES-CZB medium containing 5 mg/ml CCB in a micromanipulator chamber. 2. Hold the oocytes with the metaphase plate between the 8 o’clock and 10 o’clock positions (or between the 2 o’clock and 4 o’clock positions depending on personal preference). 3. Place the tip enucleation pipette onto the surface of the zona pellucida and use a few piezo pulses with the ‘‘big hammer’’ while applying a very slight negative pressure. 4. Remove the metaphase II chromosome–spindle complex, which can be distinguished as a translucent spot, by suction, along with a small volume of cytoplasm into the pipette, and gently pull away from the oocyte until a stretched cytoplasmic bridge is pinched off. 5. After a group of oocytes has been enucleated, wash them several times in successive drops of fresh KSOM medium. Incubate the oocytes for at least 30 min before nuclear transfer injection (see Note 7).
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Fig. 15.3. A scheme for oocyte and cumulus cell (donor) preparation. In this procedure, oocytes are collected without covering with mineral oil. Therefore, oocyte collection should be finished within 10 min. Following oocyte collection, the lid of the dish is set up as an injection chamber. After drawing the lines with ink, three types of drops are made. Then, mineral oil covers all the drops.
3.3. Donor Cell Nuclear Transfer (Fig. 15.4B)
1. Gently mix the donor cumulus cells into the PVP drops in the micromanipulation chamber with fine forceps (see Note 8). 2. Place a group of 10–20 enucleated oocytes in the HEPESCZB in a micromanipulation chamber.
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Fig. 15.4. Techniques for enucleation (A) and nuclear transfer (B). (A) After 10 min culture in a HEPES-CZB drop containing CCB, each oocyte is enucleated with an enucleation pipette. MII plate in a B6D2F1 oocyte is visible under an inverted microscope with Hoffman optics. First, make a hole in the zona with a ‘‘big hammer’’ of piezo pulses and take the tip onto the MII plate. Second, suction the MII plate. Enucleation pipettes can be reused if stored by filling the tip with mineral oil. (B) After picking up donor cells in a PVP drop, carry 5–10 somatic nuclei inside the injection pipette into an injection drop. Next, break the zona. Then, push deeply into the cytoplasmic membrane moving the donor nucleus close to the tip. After a ‘‘small hammer’’ of piezo pulse to break the cytoplasmic membrane, extrude the nucleus slowly and retract the pipette.
3. Remove the nuclei from the donor cells by gently aspirating them in and out of the injection pipette. Then draw up 5–10 of the donor nuclei in a line within the injection pipette. 4. After advancing the injection pipette through the zona pellucida with the ‘‘big hammer’’ of piezo pulses, insert the pipette deep into the ooplasm while pushing forward a donor nucleus to the tip of the pipette. 5. Puncture the plasma membrane by applying a single ‘‘small hammer’’ of piezo pulse, as evidenced by a rapid relaxation of the membrane. Expel the donor nucleus into the ooplasm with a minimal amount of medium. 6. Gently withdraw the pipette from the ooplasm. 7. Leave the injected oocytes on the stage for 10 min after the last one is injected. Wash, then culture, the injected oocytes in KSOM medium until the next step, strontium activation. 3.4. Oocyte Activation and Culture with TSA
1. At 30–60 min after injection, place the reconstructed embryos in the Ca2+-free CZB medium containing 5 mM Sr2+, 5 mg/ml CCB, and 5 nM TSA (see Note 9) for 5–6 h to activate the oocytes (see Note 10). 2. Following oocyte activation, transfer those embryos in KSOM medium containing the same concentration of TSA and
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culture them for 3–4 h more (total time for TSA treatment is 8–10 h). Then, wash these embryos, and then culture until the embryos are transferred into surrogate mothers.
4. Notes 1. Normally we use 8–12-week-old F1 mice. 2. PVA keeps the injection pipette less sticky over a longer period than BSA. 3. To dissolve PVP powder, use a 60-mm culture dish (BD Falcon, ca.no.351007) for 10-mL PVP solution and place it over night at 4°C. The next day, mix well with a pipette tip until it goes into solution. 4. Finally, take time to filtrate PVP solution with little pressure to prevent separation of the syringe and filter. For storage, we normally keep one aliquot at room temperate protected from light and discard after a month. The remaining aliquots should be stored at 4°C protected from light as well. 5. Quality control of media and reagents (optional): Once everything is ready, it is imperative that all prepared reagents and media be tested for quality for use in mouse cloning. Any reduction in quality will result in failure to produce cloned embryos. Here we provide two easy ways to examine their quality. Both experimental procedures are also well-described elsewhere (41). ICSI experiments can be used to confirm quality of the PVP and media (HEPES-CZB and KSOM) as well as the technique of the ‘‘manipulator’’. After 96 h, more than 80% of fertilized embryos (B6D2F1 B6D2F1) should be expanding or expanded blastocysts. If less than 50% expand, there may be problems with either the media or the techniques. Culture of parthenogenetic-activated embryos can be also used to confirm quality of the activation conditions including Sr2+, CCB, Ca2+-free CZB medium. After 96 h, more than 80% of parthenogenetic-activated embryos should be expanding or expanded blastocysts. 6. Every time you change the media for oocytes, wash the oocytes twice in successive drops of fresh media. 7. TSA treatment during this time period could be effective (14). 8. Cumulus cells are transferred to PVP drops by pipette just after collection since these cells are easily clumped within an hour.
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9. The proper TSA concentration may differ between donor cell types. We recommend trying a 1–100-nM range of TSA. 10. The concentrations of Sr2+ and CCB are crucial for the subsequent survival and embryonic development. We recommend these concentrations be optimized using parthenogenetic activation since they could be different among mouse strains as well as laboratories.
Acknowledgments We thank Dr. T. Castranio and all laboratory members for discussion and critical reading of the manuscript. This work in our laboratory was supported by RIKEN (Strategic Program for Research and Development (FY2005) to S.K.) and the Ministry of Education, Culture, Sports, Science and Technology of Japan (17780213 to S.K., 15080211 to T.W., and the Project for the Realization of Regenerative Medicine to T.W.)
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INDEX A Antibodies Immunoblotting,.................................38–39, 57–65, 85 Immunofluorescence,...................79, 80–81, 85–86, 89, 93–94, 96, 189 Inhibitory, .......................................................78–79, 81 Antral follicle, .........................................159, 163, 167–170 Apoptosis assay,......................................2–3, 10, 11, 12–13, 39, 58 Artificial seawater calcium-free, ...................................................59, 60, 61 Assay cell cycle, .... 1–3, 9–12, 31, 40, 43, 44, 57, 89, 100, 110 PARP, ...................................................................12–13 protein,......................................38–39, 89–90, 183–185 reporter gene, ..............99, 114–115, 125, 176, 183, 184 transgene, ..........................................................183–185 TUNEL, .....................................................................12 Asterina miniata,.........................................................58–59
Confocal microscopy, see Microscopy, confocal Coverslips poly-L-lysine,............................................196, 197, 199 cRNA injection, see microinjection, mRNA synthesis, see mRNA, synthesis Cyclin,.................................... 2, 10–12, 18, 44, 45, 57, 184 Cytoskeleton, ..........................................................189–203
D Dimethyl sulfoxide,.........................................................208 DMSO, see Dimethyl sulfoxide DNA electrophoresis,............................................................12 purification,...............................................102, 117, 184 Southern blot, ...........................................................117 dsRNA, see RNAi
E
B Bacteriophage C31,..............................................113–121 Blastomere, see Injection, blastomere Buffer injection, .........................................19, 23, 68, 116, 153 Laemmli,.........................................................34, 62, 66 lysis,...........................................................20, 21, 34, 38
C Caenorhabditis elegans, ....................................58, 123–132 Calcium signaling, ..........................................67–68, 72, 74 cAMP,.............................................................158–159, 199 Cell culture,................ 79–80, 138, 160, 163, 177, 179, 193 Cell cycle assay,........... 1–3, 9–12, 31, 40, 43, 44, 57, 89, 100, 110 Cell lines stable gene transfer, ..........................106–107, 110–111 Cells BHK–21,...........................................................101–111 CHO DG44, ..............................99, 101, 105–106, 110 PtK2, ................................ 79–80, 83, 85, 88, 93, 95, 96 recombinant, .......................................99–101, 107, 110 Centrosome,....................................................................190 Chelex, ..............................................................................19
ECL, see Enhanced Chemiluminescence (ECL) Eggs dejelly, ...........................................................................7 mouse, ...........................................................17–28, 141 Xenopus, .........................1–13, 31–35, 43–51, 113–121 Embryo development,.......................................................17, 199 transgenic, .........................................................115, 120 Enhanced chemiluminescence (ECL),...............20, 25, 46, 63–64, 70, 120, 142, 162–163, 180 Epifluorescence, see Imaging, fluorescence
F Fertilization in vitro,............................................32, 35, 71, 189–190 Fixation formaldehyde, ...................................191, 194–196, 198 methanol, ..............................92, 96, 191, 194–197, 202 Flow cytometry, ......................................................108, 111 Fluorescence, see Imaging, fluorescence Follicle ovarian,..............................................152, 158, 162–163 Follicle stimulating hormone,.................................141, 158
219
MICROINJECTION
220 Index G
Gene expression, ............. 2, 32, 40, 99, 103–106, 110, 113, 125, 176, 183, 185, 209 Gene regulation, .............................................................176 Germinal vesicle breakdown (GVBD), .........44, 48, 52–54, 61–62, 135, 136, 141 GFP, see Green Fluorescent Protein (GFP) GPR3, .....................................................................158–159 Green fluorescent protein (GFP), .......................80, 93, 99, 101, 105–111, 115, 119–120, 125, 128, 132, 175, 181, 183–185 GST, .....................................................................32, 51, 73 GVBD, see Germinal Vesicle Breakdown (GVBD)
H Hybridization in situ, .........................................12, 32, 37, 39, 40, 176
I ICSI, see Intracytoplasmic Sperm Injection (ICSI) Imaging calcium, .......................................................................67 fluorescence,........................................................28, 196 live cells, ................................................................87, 90 luminescence, ..................................................17, 20, 27 Immunoblotting single cell,..................................................38–39, 57–64 Injection blastomere, ..............................................9, 37, 179–183 chamber,................................................................68–69 pressure, ........................................9, 103–106, 109–110 Integrase,.................................................................113–121 Intracytoplasmic sperm injection (ICSI),...............189–203
K Kinases Ser/Thr, ......................................................................58
L Leuteinizing hormone (LH), .........................................158 Luciferase luminescence, ........................................................17–27 quantification, ...........................................20–21, 26–27
M MAPK, see Mitogen-Activated Protein Kinase (MAPK) Maturation promoting factor (MPF), ........................57, 72
Microinjection antibodies, .....................................40, 78–79, 81–83, 86 cRNA, see Microinjection, mRNA,.......2, 99–108, 120, 123–126, 128, 135, 136, 180 inhibitors,........................... 10, 19, 22, 32, 44, 158, 207 morpholino oligos, ................................................31–40 mRNA, ...........................................1–13, 119, 181, 182 peptides, ..........................................................43–44, 52 pressure-based,....................................................17, 179 quantitative, ................................................57, 135–155 recombinant proteins, ...........................................43, 99 zebrafish, ...............................................................67–75 Microinjection chamber, see Injection, chamber Micropipette, see Pipette, injectioncalibration, see Pipette, calibration Microscopy confocal, ......................................................................70 fluorescence,............ 12, 25, 27, 67, 70, 72, 80, 85, 104, 105, 106, 110, 144, 194 Midblastula transition,..................................................2, 11 Mitogen-activation protein kinase (MAPK),.............57–65 Morpholino antisense,.........................................................31, 32–33 Mouse, ........9, 17-18, 19, 20, 24, 25, 27, 28, 60, 101, 135, 141, 143, 151, 157, 158, 159, 163, 164, 167, 168, 169, 170, 172, 207, 208, 209, 211, 215, 216 MPF, see Maturation Promoting Factor (MPF) mRNA poly-A tail,....................................................3, 4, 22–23 quantification, .......................................19, 22, 176, 185 synthesis, .......................................................21, 78, 176
N Needle injection, see Pipette, injection Nuclear transfer,......................................................207–215
O Oocyte activation,..........................................200, 208, 214–215 collecting, ..........................................140–142, 211, 213 defolliculation, ......................................................48, 49 dissection, see Oocyte, collecting enucleating, .......................................................212–213 human, ..............................................................194, 200 maturation,..........................................................58, 136 mouse, .......................135–155, 157–172, 189–190, 208 Oregon Green BAPTA, .............................................19, 25 Orthopedia,.....................................................175, 176, 181 Ovarian follicle,...............................................................158 Ovary dissection,..................................................58–59, 60–61
MICROINJECTION Index 221 P
S
Paracentrotus lividus, 176 PCR, see Polymerase chain reaction (PCR) Peptide, ........ 31–32, 43–44, 49, 51, 53, 137, 160, 175, 179 Phospholipase C zeta (PLC zeta),....................................................17–18 Pipette beveling, ....................................................144–146, 153 calibration, ........................................149, 150, 154, 155 constriction, ......................................137, 146, 151–152 holding, ............................... 20, 23, 67, 70, 74, 75, 135, 136, 137, 139, 140, 143, 146–147, 149, 150, 154 injection, ....... 23, 45, 48–49, 69, 71, 74, 135, 136, 137, 140, 143, 144, 146–151, 159, 167, 169, 170, 172, 208, 211, 214 mercury brake,...................................................147–148 silanized,....................................................................171 transfer, ..................... 37, 46, 51, 87, 135, 136, 137, 138, 141–143, 168 Plasmid pCMV-EGFP-DI-attB, ..........................................120 pCS2, ................................................................177, 181 pET11-phiC31poly(A),....................115–116, 117–118 pGEX,.........................................................................51 pMYK-EGFP-puro,.........................101, 106, 110, 111 pSP64polyA, .............................................................4–5 purification,.......................................................102, 116 Poly-(ADP ribose) polymerase, see Assay, PARP Polymerase chain reaction (PCR),...............4–5, 78, 90, 91, 93, 95, 175, 176, 178, 183–185 Protamine sulphate, ................................................177, 179 14-3-3 proteins, ....................................................31–32, 38 Protein inhibition, .............................................................77–96 recombinant, ...................................43, 51, 99–101, 107
SCNT, see Somatic cell nuclear transfer (SCNT) Sea urchin, see Paracentrotus lividus SiRNA, .........................................77–78, 83, 85, 90–93, 95 Somatic cell nuclear transfer,..................................207–215 Sperm activation,............................................................74, 135 electron microscopy, .................................................202 factor, see Phospholipase C, zeta fixation, .............................................................202–203 immunoelectron microscopy, 203 pathology,..........................................................198–199 Starfish, see Asterina miniata
R
Xenopus laevis,......... 1–13, 31, 32, 40, 43–51, 52, 113–121
RNA quality,...................................................................19, 23 RNAi,..........................77–79, 81, 83, 85–87, 89–91, 93–96 RNAse, .............. 3, 5–7, 12, 19, 21–23, 118, 176–177, 182 Rose chamber,...........................................82, 86, 87, 90, 94
T Transcription, see mRNA, synthesis Transfection, .........................78, 83, 85, 90, 91–96, 99–100 Transformation, ................................................21, 123–132 Transgene,.. 2, 113–121, 125, 132, 175, 178–181, 183–185 Trichostatin A,........................................................207, 210 TSA, see Trichostatin A TUNEL, see Assay, TUNEL
V Vibration isolation, .........................................139, 143, 152
W Western blotting, see Immunoblotting Worms, see Caenorhabditis elegans
X
Z Zebrafish, ....................................................................67–75 Zona pellucida, ...................23, 24, 150, 194–195, 212, 214
