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Colloquium Nuclear reprogramming and stem cell creation J. B. Gurdon*, J. A. Byrne, and S. Simonsson Wellcome Trust/Cancer Research UK Institute, Tennis Court Road, Cambridge CB2 1QR, United Kingdom; and Department of Zoology, University of Cambridge, Cambridge CB2 3EJ, United Kingdom The transplantation of a somatic cell nucleus to an enucleated egg results in a major reprogramming of gene expression and switch in cell fate. We review the efficiency of nuclear reprogramming by nuclear transfer. The serial transplantation of nuclei from defective first-transfer embryos and the grafting of cells from such embryos to normal host embryos greatly increases the proportion of nuclei that can be seen to have been reprogrammed. We discuss possible reasons for the early failure of most nuclear transfers from differ- entiated cells and describe the potential value of growing oocytes, rather than unfertilized eggs, as a source of nuclear reprogram- ming molecules and for the eventual identification of these mol- ecules. Nuclear transfer provides a possible route for the creation of stem cells from adult somatic cells. Nuclear reprogramming is a term used to describe changes in gene activity that are induced experimentally by introducing nuclei into a new cytoplasmic environment. When nuclei from partially or fully differentiated cells are transplanted to enucle- ated eggs of Amphibia or mammals in second meiotic meta- phase, blastula or blastocyst embryos can be obtained, and these can form a wide range of tissues and cell types. The multipotency of these nuclear transplant embryos means that they share some characteristics of early stem cells. Indeed those nuclear trans- plant embryos that undergo growth (after feeding in Amphibia, and after implantation in mammals) to become adults must contain stem cells for renewing tissues. To this extent nuclear transplantation can achieve the creation of stem cells or stem- like cells from somatic cells of very restricted developmental potential. In contrast, the identification and isolation of natural stem cells from normal tissues is a difficult process and has not yet been successful for most vertebrate tissues (1~. Furthermore the differentiated state of cells is very stable, and it is hard to induce cells that have embarked on one pathway of differentiation to switch to another. Therefore nuclear transplantation is at present the most reliable way of deriving multipotential cells from a tissue of any kind. For this reason, nuclear reprogram- ming is of interest as a means of creating a range of replacement cells of the same genetic type as the donor source, thereby avoiding the need for immunosuppression as is required with most genetically nonhomologous grafts or implants. The aim of this article is to summarize the efficiency of nuclear reprogram- ming by nuclear transfer and hence to comment on its potential as a source of stem cells. Nuclear Transplant Embryo Development How efficiently and effectively can somatic cell nuclei be repro- grammed to an embryonic state? In the case of Amphibia, these questions have been addressed by the early nuclear transfer experiments carried out with Rana and Xenopus. For reasons that are still not clear, nuclear transfer success declines rapidly with increasing donor age in Rana (2), and nuclear transplant development is much more successful in Xenopus. In the endoderm lineage, which has been analyzed in greatest detail (3, 4), it is now known that cells express the endoderm-lineage www.pnas.org/cgi/doi/10.1 073/pnas.1 834207100 marker endodermin (5) from the late neurula stage onward. Well before this stage, endoderm cells are specified [i.e., they form only endoderm derivatives as explants (6, 7~] and are determined ti.e., they form only endoderm derivatives if trans- planted to ectopic sites (844. Yet nuclei transplanted from more advanced-stage endoderm cells into unfertilized eggs form func- tional muscle and nerve cells in ~20% of all cases (Table 1~. Even at the heartbeat stage, when the endoderm has begun regional differentiation (9, 10), 13% of nuclear transfers from the endoderm can form functional muscle and nerve cells (Table 1~. Likewise, the region of the mesoderm destined to form muscle expresses the myogenic genes MyfS and MyoD by the late gastrula stage (11, 12), and cells from this region continue to express muscle markers even when these cells are transplanted singly to the endoderm (134. Yet the nuclei of myogenic cells generate a functional nervous system in >5% of nuclear trans- plants. These results cannot be attributed to escaped germ cells or other rare cell types residing in the endoderm or muscle, because the success rate is too high. Using nuclei from differentiated or adult cells, the success rate of nuclear transfers is much lower than from larval or embryo cells (Table 14. In the case of adult Xenopus tissues, the cells that grow out from explants are of fibroblastic morphology and often do not express differentiation markers. However, for the even- tual purposes of cell replacement, the accessibility of adult tissue, as in the case of skin or blood, is much more important than the definition of cell type. In Xenopus experiments, cells from adult skin have been obtained by outgrowth in culture and retain expression of an epidermal keratin marker. Nuclei from these cells give nuclear transfer results with the same efficiency as cells from other adult organs (14~. About 1% of eggs receiving transplanted nuclei from cells of adult skin reach the muscular response stage and therefore have functional muscle and nerve cells (Table 1~. Work with mammals has given comparable results (15), although relatively few experiments have been done in which nuclei of defined cell types have been transplanted to enucleated eggs. The overall conclusion from these direct nuclear transfer experiments is that a substantial proportion of nuclei from specified or determined embryonic cells expressing differentia- tion markers undergo major reprogramming when transplanted to enucleated eggs. Serial Nuclear Transfers and Grafts The question arises as to whether the low percentages of nuclear transfer success shown in Table 1 for nonendoderm nuclei mean that only a minority of cells in a tissue have the capacity to be reprogrammed or that this capacity exists but has not been demonstrated for technical or other reasons. In Amphibia, a This paper results from the Arthur M. Sackier Coiloquium of the Nationai Acaclemy of Sciences, "Regenerative Medicine," held October 18-22, 2002, at the Arnoicl ancl Mabei Beckman Center of the National Acaclemies of Science and Engineering in irvine, CA. *To whom correspondence shouicl be aciciressecl. E-maii: email@example.com. @) 2003 by The Nationai Acaclemy of Sciences of the USA PNAS | September 30, 2003 I vol. 100 I suppl. ~ | 11819-~822 , . i,
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Table 1. First nuclear transfers using determined and differentiated cells % of total nuclear transfers reaching muscular Species Donor tissue Donor stage response stage Ref. Xenopus la evis Endoderm Muscular response stages 23-26 17-24 3 X. Iaevis Endoderm Heart beatstage 36 9-13 3 X. Iaevis Muscle Muscular response stage 26 2 42 X. Iaevis Intestinal epithelium Earlyfeeding tadpole stage47 1-3 4 X. Iaevis Skin outgrowth Adult 1.3 14 Cow Innercell mass cells Embryonic 0.6 43 Primate 4-32 cell embryos Embryonic 1.4 44 Sheep Mammary Adult 0.4 45 Mouse Cumulus Adult 2.8 46 substantial proportion of nuclear transplant embryos cleave abnormally and die as partial blastulae within 24 h of nuclear transfer. In the case of nuclei from differentiated or adult cells, partial cleavage results from one-quarter to one-third of all nuclear transfers and is far more frequent than complete cleav- age (4~. However, it has been found that the normal-appearing cells of partial blastulae can be used as donors for a second, serial set of nuclear transfers to more enucleated eggs, an experimental design first used by King and Briggs in 1956 (164. When this is done with partial blastulae derived from intestinal epithelium cells of Xenopus, it is found that many of the serial nuclear transplant embryos develop remarkably well, sometimes reach- ing the normal tadpole stage (49. These serially derived tadpoles reflect the developmental potential of the originally trans- planted intestinal epithelium cell nucleus, even though this potential was not revealed by the first nuclear transfers. The best explanation for this apparent improvement in nuclear transfer success is the following. It is believed that nuclei from slow-dividing somatic cells cannot complete their chromosome replication in time for the first cleavage of a recipient egg, which always takes place according to the time schedule of the egg, for example at 1.5 h for the first cleavage in Xenopus. Somatic cells take some 6 h to complete chromosome replication. When their transplanted nuclei are forced into early mitosis with incom- pletely replicated chromosomes, they are likely to suffer chro- mosomal damage (17, 18) and generate chromosomally defective embryos, which cannot survive. However, it sometimes happens that a transplanted nucleus fails to undergo chromosome seg- regation when the recipient egg divides into two cells and the whole replicating transplanted nucleus moves into one of the first two blastomeres. It then has a second chance to complete the replication of its chromosomes before undergoing mitosis when the egg goes from a two- to four-cell stage. As a result, partial blastulae are obtained because one of the first two blastomeres undergoes cleavage with the transplanted nucleus, while the other blastomere, having no nucleus, dies. These partial blastu- lae are more likely to contain nuclei with completely replicated chromosome sets than are those nuclear transplant embryos that undergo chromosome segregation at the first mitosis. The result of carrying out serial nuclear transplantation shows that a substantial proportion of the partially cleaved blastulae contain nuclei with wide developmental potential. When this is taken into account, the proportion of original intestinal epithe- lium cells whose nuclei can promote muscle and nerve differ- entiation rises to 20% (Table 2 and ref. 4~. Using a similar line of thinking, it was found that the normal- appearing cells of partial blastulae can be grafted to host embryos and then reveal a wide range of developmental poten- tial. Using GFP-marked donor nuclei, it has been calculated that at least 16% of differentiated larval intestinal epithelium cells contain nuclei capable of completely different pathways of differentiation (193. In conclusion, the use of serial nuclear transfer and grafts shows that a much higher proportion of differentiated cells contain nuclei that undergo a major repro- gramming by egg cytoplasm than the 1-3% apparent when considering only first nuclear transfers. Possible Reasons for Failures After the transfer of nuclei from differentiated cells to enucle- ated eggs, whether in Amphibia or mammals, only a few of the nuclear transplants develop into adult animals. There are several possibilities that could account for this low success rate, and the primary reason for developmental failure may differ between Amphibia and mammals. In Amphibia, the majority of eggs receiving transplanted nuclei from differentiated or adult cells undergo only a few irregular cleavages or fail to divide at all. Thus, the primary loss in amphibian somatic cell nuclear transfer is during the very earliest cleavages. In mammals, the develop- mental loss tends to be greater after the initial cleavages. In primates, development to the early eight-cell stage has been shown to be similar in somatic and embryonic cell nuclear transfers and after intra-cytoplasmic sperm injection (ICSI) (between 80~o alla 90% in every case). However, the develop- ment of cloned embryos to the later blastocyst stage was markedly different; only 1% of somatic cell nuclear transfers reach the blastocyst stage, whereas 34% of embryonic nuclear transfers and 46% of ICSI controls reach this stage (204. Table 2. The combination of first and serial nuclear transfer results % of total nuclear transfers reaching muscular Species Donor tissue Donor stage response stage Ref. X. Iaevis Intestinal epithelium Early feeding tadpole stage47 20 4 X. Iaevis Skin outgrowth Adult 11-12 14 X. Iaevis Kidney, lung, heart Adult 13 47 11820 1 www.pnas.org/cgi/doi/10.1073/pnas.1834207100 ., it, . . Gurdon et al.
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These variations in the stage of developmental loss probably reflect different problems encountered after somatic cell nuclear transfer in Amphibia and mammals. In Amphibia the primary problem may be the previously mentioned difficulty that somatic cell nuclei have in completing chromosome replication within the very limited time (only 90 min in Xenopus) before eggs undergo cleavage (21~. In mammals there are 20 h for the mouse, or longer for humans, between the time of fertilization and the first cleavage division. This makes it unlikely that incomplete chromosome replication is a problem in mammals. However, mammals have imprinted genes (22) that Amphibia do not (23), and it has been suggested that the developmental loss observed after mammalian somatic cell nuclear transfer could be caused by the incomplete reprogramming of various imprinted genes (as well as Oct4) (24~. Another recent suggestion for mammalian developmental failure is that enucleation removes maternal spindle proteins required to maintain ploidy through the initial cleavages (25~. Although amphibian cloned embryos tend to suffer from chromosomal damage caused by incomplete chromosome rep- lication and mammalian cloned embryos tend not to express imprinted genes correctly, there is also a range of other factors that may affect both amphibian and mammalian nuclear trans- plants. Quantitatively incomplete/incorrect reprogramming of gene expression has been found in both amphibian (19) and mammalian (26) cloned embryos, and it has been suggested that this may affect development (27~. Also, it has been suggested that the stage of the donor cell cycle may be critical to avoid aneuploidy through re-replication of the donor genome (28~. Another idea concerns the centriole; this is normally introduced with the sperm at fertilization and the eggs of most animals do not contain their own centriole. Perhaps most differentiated cells that are no longer required to divide do not contain a fully functional centriole. It is possible that technical factors may also be important. Failure to rupture a donor cell would certainly account for the total lack of cleavage. On the other hand, the position in an egg at which a transplanted nucleus is deposited is presumed not to matter on the grounds that sperm, entering from the surface of the egg, always find their way to the correct central position of the egg. In conclusion, there is no definitive explanation for the high frequency with which nuclei transplanted from differentiated or adult cells fail to elicit any cleavage or development of recipient eggs. It is probably a combination of aneuploidy, genetic damage, and incomplete epigenetic reprogramming. For the purposes of cell replacement, this is a serious problem only if the supply of recipient eggs is strictly limited, as might be the case for humans. The Cell Differentiation Potential of an Imperfect Genome A fundamental idea behind the original vertebrate nuclear transfer experiments was that a complete genome is required for an egg to develop to a normal adult. This is very likely to be true; in fact, the completeness of a genome might be defined in this way. But this does not at all mean that each individual pathway of cell differentiation also depends on ~ complete genome. Nuclei that lack essential genes for one developmental pathway may nevertheless be able to proceed along other differentiation routes. Nuclear transfer experiments in both Amphibia and mammals have given support to this idea. In Xenopus it has been found that many of the partially cleaved nuclear transplant embryos (that are developmentally defective) have quantita- tively aberrant expression of early zygotic genes (19~. Despite this, healthy cells from such embryos, genetically marked by GFP, were able, after grafting to host embryos, to participate in the differentiation and growth of normal muscle, notochord, epidermal, and other cells (19~. In mice, it was discovered that whereas only 2% of nuclear transplant embryos could develop into adult animals, 9% of nuclear transplant embryos could Gurdon et a/. Oocyte growth: 9 months Blastula Development: 7 hours 7 hours at 1 8°C DNA replication Cell division Unfertilized egg No transcription Blastula 1 cell 10,000 cells 4daysatl8°C No DNA replication No cell division Oocyte 1 cell Intense transcription Oocyte 1 cell 4 days at 37°C ~ - Unfertilized egg 1 cell DNA replication. Slow cell division. Transcription starts Blastocyst ~100 cells Fig. 1. Diagrams to show the time scale of nuclear reprogramming in Xenopus and the mouse. (A) Oogenesis and early development of Xenopus. (B) In Xenopus egg nuclear transfers, reprogrammed gene expression is seen at the late blastula stage after at least 12 cell division cycles. (C) In Xenopus oocyte nuclear transfers, reprogrammed gene expression is seen in the com- plete absence of DNA replication and cell division. (D) In mouse nuclear transfer to eggs, reprogrammed gene expression has been seen at the blas- tocyst stage. produce embryonic stem cell lines (29~. This finding suggests that defective cloned mammalian embryos, which are incapable of developing into an entire mouse, can still produce useful stem cell lines. An interesting future direction of research will be to investigate the differentiation capacity of transplanted nuclei carrying known chromosomal or gene deficiencies. Perhaps genetically deficient cells may be entirely suitable for somatic cell replacement. Reprogramming Without Replication The great majority of nuclear transfer experiments in both mammals and Amphibia have been carried out with eggs in second meiotic metaphase as recipient cells. In all of these cases, the first response of an enucleated egg to a transplanted nucleus is the induction of DNA synthesis and cell division (Fig. 1~. In Amphibia, new transcription, and hence evidence of nuclear reprogramming, commences after 12 cell cycles at the midblas- tula transition, 5 h after nuclear injection. In mice, new tran- scription starts at the early two-cell stage, ~24 h after nuclear injection, and new transcription starts later in other mammalian species. This situation raises the possibility that nuclear repro- gramming requires DNA replication and/or cell division to reset an epigenetic program, a suggestion made by Tada and Tada (30~. PNAS | September 30, 2003 I vol. 300 | suppl. ~ | 11821
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To test this possibility, we have transplanted somatic cell nuclei into nondividing amphibian oocytes in the prophase of first meiosis. These cells cannot be fertilized, are inactive in DNA synthesis, but are intensely active in transcription (Fig. 1~. Their active genes are maximally packed with RNA polymerase, as seen in the spectacular transcription complexes of Miller (31~. We transplanted between 10 and 100 somatic cell nuclei to a single oocyte to obtain a detectable response, and the results were assessed by 2D protein analysis. The injected nuclei un- derwent a large increase in volume and dispersion of their chromatin over the several days for which injected oocytes can be cultured. Protein analysis showed that new proteins were synthesized by oocytes injected with mammalian nuclei from cultured HeLa cells, although the identity of the proteins was not known (32, 33~. In the case of nuclei from one species of amphibian (Pleurodeles) transplanted to oocytes of Xenopus, some new proteins were synthesized with the size and charge properties of oocyte expressed genes (34~. In other experiments, oocyte-specific 5S genes were activated when Xenopus somatic cell nuclei with inactive oocyte-type SS genes were injected into oocytes (35) and liver-specific enzymes were inhibited in exper- iments with two species of Ambystoma (364. Very recently we have extended our analysis of nuclear transfer in Xenopus oocytes. Quite surprisingly, we find that the nuclei of differentiated adult cells of mice (thymocytes) and humans (white blood cells) can be to some extent reprogrammed by Xenopus oocytes (37~. In particular, the diagnostic pluripo- tency stem cell marker gene oct4 is induced in these mammalian nuclei after injection into the germinal vesicle (enlarged oocyte nucleus'. The oct4 transcripts have the human or mouse se- quence when oocytes are injected with human or mouse somatic 1. McKay, R. (2000) Nature 406, 361-364. 2. Briggs, R. & King, T. J. (1957) J. Morphol. 100, 269-312. 3. Gurdon, J. B. (1960) J. Embryol. Exp. Morphol. 8, 505-526. 4. Gurdon, J. B. (1962) J. Embryol. Exp. Morphol. 10, 622-640. 5. Sasai, Y., Lu, B., Piccolo, S. & De Robertis, E. M. (1996) EMBO J. 15, 4547-4555. 6. Hudson, C., Clements, D., Friday, R. V., Stott, D. & Woodland, H. R. (1997) Cell 91, 397-405. 7. Newman, C. S., Chia, F. & Krieg, P. A. (1997) Mech. Dev. 66, 83-93. 8. Wylie, C. C., Snape, A., Heasman, J. & Smith, J. C. (1987) Dev. Biol. 119, 496-502. 9. Chalmers, A. D. & Slack, J. M. (1998) Dev. Dyn. 212, 509-521. 10. Zorn, A. M. & Mason, J. (2001) Mech. Dev. 103, 153-157. 11. Harvey, R. P. (1990) Development (Cambr~dge, U.K) 108, 669-680. 12. Hopwood, N. D., Pluck, A. & Gurdon, J. B. (1989) EMBO J. 8, 3409-3417. 13. Kato, K. & Gurdon, J. B. (1993) Proc. Natl. Acad. Sci. USA 90, 1310-1314. 14. Gurdon, J. B., Laskey, R. A. & Reeves, O. R. (1975) J. Embryol. Exp. Morphol. 34, 93-112. 15. Campbell, K. H., McWhir, J., Ritchie, W. A. & Wilmut, I. (1996) Nature 380, 64-66. 16. King, T. J. & Briggs, R. (1956) Cold Spring Harbor Symp. Quant. Biol. 21, 271-290. 17. Briggs, R., King, T. J. & Di Berardino, M. A. (1960) Symp. Germ Cells Dev. 441-477. 18. DiBerardino, M. A. & King, T. J. (1967) Dev. Biol. 15, 102-128. 19. Byrne, J. A., Simonsson, S. & Gurdon, J. B. (2002) Proc. Natl. Acad. Sci. USA 99, 6059-6063. 20. Mitalipov, S. M., Yeoman, R. R., Nusser, K. D. & Wolf, D. P. (2002) Biol. Reprod. 66, 1367-1373. 21. Graham, C. F., Arms, K. & Gurdon, J. B. (1966) Dev. Biol. 14, 349-381. 22. Surani, M. A., Barton, S. C. & Norris, M. L. (1984) Nature 308, 548-550. 23. Meehan, R. R. & Stancheva, I. (2001) Essays Biochem. 37, 59-70. 24. Bortvin, A., Eggan, K., Skaletsky, H., Akutsu, H., Berry, D. L., Yanagimachi, R., Page, D. C. & Jaenisch, R. (2003) Development (Cambridge, U.K) 130, 1673-1680. 25. Simerly, C., Dominko, T., Navara, C., Payne, C., Capuano, S., Gosman, G., Chong, K. Y., Takahashi, D., Chace, C., Compton, D., et al. (2003) Science 300, 297. 11822 1 www.pnas.org/cgi/doi/10.1073/pnas.1834207100 cell nuclei, respectively. The ability to activate oct4 expression in the nuclei of adult somatic cells may increase the probability of deriving embryonic stem cells from nuclear transplant embryos (38~. Evidently amphibian oocytes contain molecules and con- ditions that can at least partially reprogram nuclei of adult mammalian cells. Because one Xenopus oocyte has 4,000 times the protein content of a mammalian egg, and because one female Xenopus contains some 25,000 oocytes, the Xenopus ovary constitutes favorable material for the identification of repro- gramming molecules and mechanisms. Proliferation of Reprogrammed Cells To be clinically useful, it will be necessary to be able to extensively proliferate reprogrammed cells. The remarkable discovery, primarily of Evans (39), that mouse blastocyst cells can be made to proliferate almost indefinitely in culture, as embryonic stem cells, without losing their potential for differ- entiation into most, and sometimes all, cell types of an adult gives great encouragement in this requirement. We heard at this meeting about much current research directed toward under- standing the mechanisms and control of embryonic stem cell proliferation and differentiation. It may eventually be necessary to build into cells destined for replacement a finite proliferative capacity to reduce the likelihood of donated cells becoming cancerous. In conclusion, we suggest that the extraordinary reprogram- ming capacity of eggs and oocytes may lead to the identification of reprogramming molecules and mechanisms. These may facil- itate a route toward cell replacement in humans, by a combina- tion of nuclear transfer, stem cell creation, and embryonic stem cell proliferation, as suggested by the work of Munsie et al. (40) and Rideout et al. (41~. 26. Humpherys, D., Eggan, K., Akutsu, H., Friedman, A., Hochedlinger, K., Yanagimachi, R., Lander, E. S., Golub, T. R. & Jaenisch, R. (2002) Proc. Natl. Acad. Sci. USA 99,12889-12894. 27. Rideout, W. M., III, Eggan, K. & Jaenisch, R. (2001) Science 293, 1093-1098. 28. Campbell, K. H., Loi, P., Otaegui, P. J. & Wilmut, I. (1996) Rev. Reprod. 1, 40-46. 29. Wakayama, T., Tabar, V., Rodriguez, I., Perry, A. C., Studer, L. & Mombaerts, P. (2001) Science 292, 740-743. 30. Tada, T. & Tada, M. (2001) Cell Struct. Funct. 26, 149-160. 31. Miller, O. L., Jr., (1965) Natl. Cancer Inst. Monogr. 18, 79-99. 32. Gurdon, J. B. (1976) J. Embryol. Exp. Morphol. 36, 523-540. 33. De Robertis, E. M., Partington, G. A., Longthorne, R. F. & Gurdon, J. B. (1977) J. Embryol. Exp. Morphol. 40, 199-214. 34. De Robertis, E. M. & Gurdon, J. B. (1977) Proc. Natl. Acad. Sci. USA 74, 2470-2474. 35. Korn, L. J. & Gurdon, J. B. (1981) Nature 289, 461-465. 36. Etkin, L. D. (1976) Dev. Biol. 52, 201-209. 37. Byrne, J. A., Simonsson, S., Western, P. S. & Gurdon, J. B. (2003) Curr. Biol. 13, 1206-1213. 38. Boiani, M., Eckardt, S., Scholer, H. R. & McLaughlin, K. J. (2002) Genes Dev. 16, 1209-1219. 39. Evans, M. (1981) J. Reprod. Fertil. 62, 625-631. 40. Munsie, M. J., Michalska, A. E., O'Brien, C. M., Trounson, A. O., Pera, M. F. & Mountford, P. S. (2000) Curr. Biol. 10, 989-992. 41. Rideout, W. M., III, Hochedlinger, K., Kyba, M., Daley, G. Q. & Jaenisch, R. (2002) Cell 109,17-27. 42. Gurdon, J. B., Brennan, S., Fairman, S. & Mohun, T. J. (1984) Cell 38, 691-700. 43. Sims, M. & First, N. L. (1994) Proc. Natl. Acad. Sci. USA 91, 6143-6147. 44. Meng, L., Ely, J. J., Stouffer, R. L. & Wolf, D. P. (1997) Biol. Reprod. 57, 454-459. 45. Wilmut, I., Schnieke, A. E., McWhir, J., Kind, A. J. & Campbell, K. H. (1997) Nature 385, 810-813. 46. Wakayama, T., Perry, A. C., Zuccotti, M., Johnson, K. R. & Yanagimachi, R. (1998) Nature 394, 369-374. 47. Laskey, R. A. & Gurdon, J. B. (1970) Nature 228, 1332-1334. Gurdon et a/. . . , ,~ . . ,
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