The ovine uterus as a host for in vitro-produced bovine embryos

ELSEVIER

THE OVINE UTERUS AS A HOST FOR IN VITRG-PRODUCED

BOVINE

EMBRYOS

C. E. Rexroad, Jr. and A. M. Powell ’ USDA, Agricultural Research Service Gene Evaluation and Mapping Laboratory Beltsville, MD 20705 USA Received for publication: Accepted:

21 January 17 February

1998 1999

ABSTRACT A series of experiments were conducted to determine whether bovine blastocysts would develop beyond the blastocyst stage in the ovine uterine environment. In Experiment 1, in vitro matured, fertilized and cultured (IVM/IVF/IVC) expanded bovine blastocysts were transferred into uteri of ewes on Day 7 or 9 of the estrous cycle and collected on Day 14 or 15 to determine if the bovine blastocysts would elongate and form an embryonic disk. Springtime trials with ewes that were synchronized with a medroxyprogesterone acetate (MAP) sponge resulted in a 78% blastocyst recovery rate, and 68% of the recovered spherical or elongated embryos had embryonic disks. In Experiment 2, transfer of 4-cell bovine embryos to the oviducts of ewes at Day 3 resulted in a lower recovery (47 vs 80%) than the transfer of blastocysts at Day 7 when embryos were recovered at Day 14. However, the percentage of embryos containing embryonic disks was higher for embryos transferred at the 4-cell stage (71%) than for embryos transferred as blastocysts (50%). In Experiment 3, IVF embryos from super-ovulated cows or Day 8 in vitro produced embryos transferred to cows were collected at Day 14 and were found to be similar in size to those produced by transfer to ewes in Experiment 2. In Experiment 4, the transfer of bovine blastocysts to ewes did not prolong the ovine estrous cycle. In Experiment 5, extension of the ovine estrous cycle by administration of a MAP releasing intravaginal device allowed bovine embryos to elongate extensively and to become filamentous. In Experiment 6, uterine flushings on Day 14 or Day 16 contained elevated levels of interferon-tau when bovine blastocyst were transferred on Day 7. Transfer of bovine embryos to the reproductive tract of a ewe allows some embryos to develop normally to advanced perimplantation stages and may be a useful tool for studying critical stages of embryo development and the developmental capacity of experimental embryos. Published by Elsevier Science Inc.

Key words:

bovine, in vitro production, ovine, transfer, embryo

Acknowledgments Special thanks to Harold Hawk for his assistance in the collection and transfer of in vivo bovine embryos. We thank Alan Ealy for assaying the uterine flushings for boIFN-t, Paul Graninger for care of experimental animals and surgical assistance, Ken Bondioli for supplying us with in vitro produced bovine embryos and Ms. Shirley Weese for putting the manuscript in final form. ’ Correspondence and reprint requests: Bldg. 200, Rm 8, BARC-East, Beltsville, MD 20705. Theriogenology 52351964, 1999 Published by Elsevier Science Inc.

0095691X/99/$-see front matter PII SOO93-691X(99)00134-X

352

Theriogenology INTRODUCTION

Early ruminant embryos undergo a rapid and great expansion of their outer trophectoderm cells and inner endoderm cells after hatching horn the zona pellucida (4,9). This early developmental event anatomically prepares the embryo for implantation in the uterine epithelium and results in the production of embryonic factors necessary for maternal recognition of pregnancy (4, 17, 11). It is not well understood what intrinsic (embryonic) or extrinsic (oviduct/uterine) factors are involved in initiating and controlling this rapid change in the bovine embryo’s extra-embryonic tissues. On the maternal side, studies in other species as well as in ruminants have shown that specific maternal factors such as hormones, cytokines, and growth factors affect blastocyst (or trophectoderm) elongation and differentiation (reviewed in 11,22, 35). For the bovine embryo, most of what is known about preimplantation gene expression and the effects of various hormones and growth factors on blastocyst development have been assayed in the prehatching or perihatching blastocyst, and most of this from in vitro studies (12, 15,23, 32, 37, 36). However, it was discovered, and is now well described, that elongation stage ruminant embryos produce (from the trophectoderm) a family of interferon molecules that mediate recognition of pregnancy (3, 21, 24, 3 1,26). Moreover, a family of pregnancy-associated glycoproteins (PAGs) that are trophectoderm-derived have been identified and cloned, but their function is yet unknown (7,40). In the near future, gene expression assays such as PCR based differential display may identify new embryo factors that play critical roles in peri-implantation development and implantation (16). The experimental investigation of the known and yet to be discovered gene products that control bovine blastocyst elongation and implantation is difficult because of the high cost of acquiring and maintaining cattle. In contrast, the production of bovine embryos from bovine oocytes through in vitro production is relatively inexpensive and allows for direct experimental manipulation in order to study various facets of early embryonic development. However, in vitro production does not suffice for per&implantation studies, since current culture systems do not support the development of a well-organized, elongated bovine embryo with an embryonic disk. Furthermore, an in vivo host is needed to study interactions of the conceptus and the uterus during the period of elongation and implantation (4, 17,22). Thus, an experimental model system is needed to advance experimentation on peri-implantation bovine embryo development and on placentation failure in the cow. Study of this developmental stage is particularly important since a large proportion of the of the reproductive failures in domestic farm animals, estimated to be as high as 80%, results from abnormal per&implantation development and unsuccessful implantation (8, 11, 19). One possibility is that in vivo development of bovine embryos in other domesticated and less costly species may provide a system that is experimentally viable and relevant. Before culture media were available that supported development of bovine embryos beyond the 8- to 16-cell block, ligated sheep oviducts were used for in vivo culture to permit g-cell bovine embryos to develop to late morulae or blastocysts (38,39). Subsequently, Eyestone et al.

Theriogenology

353

(13) showed that bovine embryos transferred into the oviducts of ewes develop from the one cell to the blastocyst stage. The ligated sheep oviduct was subsequently used as an intermediate host for early development of bovine embryos after microinjection with DNA or after transfer of nuclei into enucleated oocytes (6,27). Such use permitted the selection of developing embryos for nonsurgical transfer back into the cow uterus to complete gestation. We initiated the present experiments because we needed an economical source of elongation stage bovine embryonic disks to transfer into severe combined immuno-deficient (SCID) mice to determine their potential as a source of bovine embryonal carcinoma cells. In vitro produced bovine 4-cell and blastocyst stage embryos were transferred into ovine uteri, and their anatomical (elongation and presence of an embryonic disc) and physiological development (interferon-tau production) was assessed. These experiments demonstrated the ability of the ovine uterus to maintain early development of bovine embryos. MATERIALS

AND METHODS

Production of Bovine In Vitro Produced Embryos Oocytes were aspirated from ovaries collected at an abattoir and washed. The cumulus was partially removed by pipetting, and the cumulus-oocyte complexes (COCs) were cultured in maturation medium (Ham’s FlO with 10% fetal bovine serum [FBS] +0.3 pg/mL USDA-PLHB6). After 24 h of culture, the maturation medium was aspirated and replaced with IVF-TL solution (BSS-010-D, Specialty Media, Lavalette, NJ) supplemented with 6 mg/mL fatty-acid free bovine serum albumin (BSA) and 0.275 mg/mL pyruvate. Spermatozoa from 1 straw of commercially prepared semen was washed in 10 mL Dulbecco’s phosphate buffered saline (DPBS) by centrifugation. The spermatozoa were resuspended in 400 pL of IVF-TL solution and were capacitated in the presence of 20 ug heparin for 15 min and subsequently diluted in 11 mL of IVF-TL solution, for a final concentration of approximately 450,000 sperm/ml One-half milliliter of diluted spermatozoa was added to each well of 50 oocytes. Twenty-hours after the addition of spermatozoa, zygotes were vortexed for 3 min in normal DPBS supplemented with 1% BSA to remove adherent cumulus cells. Embryos were cultured in 500 uL CRIaa (28) with 8.9 mg/mL alanine and 75 mg/mL glycine +10% FBS on a monolayer of Buffalo rat liver (BRL) cells (CRL 1442; ATCC, Rockville, MD) that had been plated at 10,000 cells/well 2 to 3 d previously in 4-well Nunc (Roskilde, Denmark) tissue culture plates. Embryos were transferred to new plates every third day of culture. Blastocysts were collected for transfer as indicated for each experiment. Prior to transfer, the embryos were washed and held in DPBS with 1% BSA (Fraction V, Sigma, St. Louis, MO). The experimental protocols used in this study were approved by the Beltsville Area Institutional Animal Care and Use committee.

354

Experiment

1

Day 8 expanded and hatched bovine blastocysts were transferred into one or both uterine horns of luteal phase ewes during mid-ventral laparotomy on Day 7 or 9 of the estrous cycle. During transfer the upper half of the uterine horn was punctured with a blunt probe. Blastocysts were aspirated into a positive displacement pipette in about 10 pL of medium, the pipette was then inserted through the puncture site, and the blastocysts were deposited into the lumen of the uterus. In Trial 1, ewes were synchronized during the spring by insertion of a MAP sponge into the vagina for 10 d. The ewes were mated at estrus 48 h after MAP sponge removal. Ten to 19 blastocysts were transferred into each uterine horn. Trial 2 was subsequently conducted during the fall with ewes that were synchronized with a MAP sponge as described above. Ewes were not bred before 9 to 16 blastocysts (4 to 8/horn) were transferred into each ewe on Day 7. Ewes were sacrificed on Day 14 or 15, and embryos were flushed horn uterine horns using DPBS with 10% ovine serum. Recovered embryos were counted and inspected for the presence of an embryonic disk and were classified as spherical or elongated. In Trial 1, the more advanced filament development in ovine embryos made them readily distinguishable from cow embryos. Experiment 2 Bovine embryos were transferred either into the oviducts of 6 ewes on Day 3 as 4-cell embryos or into the uteri of 6 ewes on Day 7 as expanded or hatched blastocysts to determine if early transfer would improve development beyond the blastocyst stage. Either 9 or 10 embryos were transferred per oviduct or uterine horn. Ewes were sacrificed and the embryos flushed from the uterine horns on Day 14 of the estrous cycle. The length and width of embryos was measured using an optical micrometer at constant magnification. The product of the two measurements (area) was considered a suitable index of surface area. Each embryo was inspected for the presence of an embryonic disk. Experiment

3

Developmental similarity of bovine in vitro produced embryos after residence in the sheep uterine environment in Experiment 2 was compared with bovine embryos recovered at a corresponding age from 3 primaparous Holstein cows that had received in vitro produced blastocysts by transcervical embryo transfer on Day 7 of the estrous cycle (10 blastocysts/hom) and from 2 superovulated primaparous Holstein cows. In the latter, superovulation was induced in luteal phase of the cycle by administering 32 mg FSH in 8 decreasing doses over 4 consecutive days, Prostaglandin F,a (25 mg) was administered concurrent with sixth dose of FSH to induce luteal regression. Cows were observed for estrus and inseminated with freshly collected bull semen. Embryos were recovered by nonsurgical flushing of the cows on Day 14 of the estrous cycle. Recovered embryos were measured as described in Experiment 2.

Theriogenology

355

Experiment 4 To determine if the presence of developing bovine embryos in the reproductive tract of the ewe would extend the estrous cycle, bovine in vitro produced Day 8 blastocysts were transferred by mid-ventral laparotomy into the uterine horns of ewes on Day 7 to 9 of the estrous cycle. When corpora lutea (CL) were present on both ovaries, 3 to 5 blastocysts were transferred into each horn. When CL were only present on 1 ovary, 7 to 10 blastocysts were transferred ipsilateral to CL. Ewes were observed for estrus twice daily and sacrificed on Day 21 to determine the status of the reproductive tract of those ewes that had not exhibited e&us. Experiment

5

To determine if prolonging the estrous cycle by MAP sponge would permit additional embryonal development, bovine in vitro produced Day 8 expanded and hatched blastocysts were transferred into the uterine horns of 3 ewes by mid-ventral laparotomy on Day 7 of the estrous cycle. On the day of transfer a MAP sponge was inserted into the vagina of each ewe. Ewes were sacrificed on Day 17, 18 and 23 of the prolonged estrous cycle, and the recovered embryos were assessed for elongation and the presence of an embryonic disk. Experiment 6 To assess if bovine embryos that were developing in the ovine uterine environment were able to produce bovine interferon-tau (boIFN-t), bovine in vitro produced Day 8 expanded and hatched blastocysts were transferred by mid-ventral laparotomy on Day 7, and a MAP sponge was inserted into the vagina of the ewe. Six ewes were sacrificed on Day 14 or 16, each uterine horn was flushed with 10 mL of DPBS with 10 % sheep serum, and antiviral assays of uterine flushings for boIFN-t were completed as described by Roberts (26). Fifty microliters of each sample were added to 100 PL of medium, and a 3-fold serial dilution was completed in a 96-well tissue culture plate. After addition and co-incubation with Madin-Darby kidney cells for 24 h, the cells were challenged with virus, and after 18 h viability of cells was determined by fixing the monolayers in methanol and staining with Gentian violet. The ability of all samples to prevent virus-induced cell lysis by 50% was compared with the laboratory standard, recombinant boIFNtl (5.4 x 10.’ IU/mg), which provides 50% inhibition from virus-induced cytolysis at 5.3 pM. Sensitivity of the assay for samples was 15.9 pM (0.36 ng/mL). The uteri of 2 control ewes, that had received MAP sponges on Day 7 but had not received bovine embryos were also flushed and assayed for the presence of boIFN-t. Statistical Analyses For Experiments 2 and 3, the recovery rate, area, and percentage of embryos with an embryonic disk were analyzed by the Proc Mixed procedure of SAS (29). The error term for analyses of transfer stage was based on the interaction of stage with recipient. The error term for recovery rate and percentage of embryos with an embryonic disk was the residual error. Data presented are least squares means.

Theriogenology RESULTS Experiment

1

Transfer of bovine blastocysts into the uteri of ewes resulted in continued development of the embryos, with most of the embryos becoming large spheres or undergoing elongation (Table 1). Most of the recovered embryos (68%) had developed an embryonic disk (Figure la). Of the recovered embryos, approximately 3 times as many were elongated as were spherical, indicating extensive remodeling of the embryo. Table 1. Development of Day 8 bovine in vitro produced blastocysts transferred into the uteri of ewes on Day 7 or 9 in the spring or on Day 7 in the fall season. Embryos

Season Ewes Transfer n Day

Recovery Day

Transferred Recovered n n (%)

Degenerated %

Spherical or elongated n (%)

With disk n (“XI)

Spring

3

9

15

81

55 (68)

25

41 (75)

37 (67)

Spring

3

7

14

79

70 (89)

0

70 (100)

48 (69)

7

14

65

44 (68)

0

44(100)

ND*

Fall 5 a Not determined.

Experiment 2 Transfer of 4-cell embryos at Day 3 of the ovine estrous cycle resulted in a significantly lower embryo recovery rate compared with transfer of blastocysts on Day 7 (Table 2). When the embryos were recovered on Day 14, a higher percentage of those that had been transferred at the 4-cell stage had embryonic disks than those transferred as blastocysts. Moreover, the embryos transferred at the 4-cell stage appeared to be larger than those transferred as blastocysts, but the variability was sufficiently large that the calculated areas were not significantly different (Table 2).

357

Theriogenology Table 2.

Recovery, length, area, and embryonic disk development in bovine embryos recovered on Day 14 after transfer of IVP embryos into ewes on Day 3 or 7 or into cows on Day 7, and for in vivo fertilized oocytes in superovulated cows. Embrvos Transfer stage/in vivo

Transferred n

Recovered (“~6f SEM)a

Length (mm f SEM)a

Area (mm’ f SEM)a

With disks (“x7f SEM)a

Ewe (6)

4-cell

118

47 * 5.9b

1.8 +~.43~ (.2 - 13S)d

2.1 f .63b (.02 16.2)d

71 f 6.4b

Ewe (6)

Blastocyst

120

80 * 5.9"

1 .Of .42b (.2 - 3.0)d

1.O * .62b (.03 - 8.1)d

50 f 6.0'

Blastocyst

60

Q&&) -21(35)

1.3 f .38b (.5 - 4.5)d

1.2 f .44b (.27 - 3.8)d

80*4.1b

in vivo

NA’

Host (n)

Exueriment Cow(3)

3

Cow(2)

1.7* .42b 1.2 f .39b 79+ 4.6b (.12 - 4.7)d (.6 - 5.1)d a Values are least squares means and standard errors of means. bc Means within experiment and column lacking a common superscript differ, (P
Experiment

24

3

The calculated areas and percentage of embryos with embryonic disks were similar for embryos recovered on Day 14 Tom cows that were superovulated and fertilized in vivo and cows that received in vitro produced Day 8 blastocysts (Table 2). While the recovery rate for the latter 3 cows appeared low (35%), and was considerably lower than the recovery rate for cow embryos recovered from ewes, this may be largely due to the difference in flushing methods used to recover the embryos. The gross morphology appeared to be the same for cow embryos recovered from cows in this experiment and from sheep in Experiment 2. The largest embryos were similar in size in each group.

358

Theriogenology

Experiment 4 Transfer of bovine embryos into ewes did not extend the length of their estrous cycles; 3 of 6 ewes returned to estrus on Days 15 to 17. Estrus was not detected in the other 3 ewes; however, one or both ovaries from these 3 ewes contained recent ovulation sites when they were examined on Day 21. At that time, no trace of bovine embryos was evident in the uterine lumen or in fluid that was flushed through the uteri. Experiment 5 Treatment of recipient ewes with MAP sponges was successful in permitting some of the bovine embryos to continue developing. While only 12 embryos were recovered of the 46 blastocysts that were transferred, 10 bovine embryos had embryonic disks (Figure lb), and 7 of the embryos were filamentous (Figure lc).

Theriogenology

c Figm ‘e 1 a) Typical appearance of bovine embryos Day 14, after 7 days in an ovine uterus. Embryonic disc is evident, indicated by the black arrow (x 30). b) Typical appearance of bovine embryos on Day 16, nine days after transfer to a ewe that had been treated with a MAP sponge. Embryonic disk is indicated by the black arrow (x 70). c) Typical appearance of bovine embryos on Day 18, eleven days after transfer to a ewe that had been treated with a MAP sponge (X 7).

359

360

Theriogenology

Experiment 6 The uterine flushings recovered on Days 14 and 16 contained elevated levels of boIFN-t in all 6 ewes bearing bovine embryos compared with flushings from 2 control ewes (Table 3). Although boIFN-t concentration tended to be higher on Day 16 than on Day 14, the mean concentration did not differ. Recovery of embryos was similar for Day 14 (27 of 38) and Day 16 (29 of 37).

Table 3. Bovine interferon-tau (boIFN-t) measured in ovine uterine flushings of control ewes or after the transfer of Dav 8 in vitro oroduced bovine blastocvsts. Embryos Recovery day

Transfer day

Ewes n

Transferred n

Recovered n (%)

boIFN-t ng/mL f SEM

14 - 16

N A”

2

NA

NA

CO.36

7

3

37

29 (78)

166* 118

3

38

27 (71)

3733 f 1995

14

16 7 a NA = not applicable.

DISCUSSION The results demonstrated that bovine in vitro produced embryos can undergo continued development in the reproductive tract of ewes when transferred either as 4-cell embryos or as expanded or hatched blastocysts. Development continued with extreme elongation (filamentous) when ovine estrous cycles were extended with MAP sponges. These results appear to be novel and go beyond findings that the ovine oviduct maintains the development of bovine embryos from the l-cell to the blastocyst stage (6, 13). These findings suggest that the uterine environment of the ewe provides most of the nutrition or growth factors needed to stimulate the early development of the post-hatching bovine embryo. Development of in vitro produced bovine embryos in the reproductive tract of ewes was similar to that of in vivo fertilized superovulated bovine embryos or of in vitro produced embryos transferred to the bovine uterus based on their size distribution and the presence of embryonic disks on most of the embryos. Extending the ovine estrous cycle with exogenous MAR allowed bovine embryos to develop to the tilamentous stage. The average calculated area of bovine embryos recovered on Day 14 was 1 and 2.1 mm* in Experiment 2, which was similar to the 1.2 mm* for embryos recovered from cows in Experiment 3. The embryos recovered in our experiments appeared to be smaller than those of Grealy et al. (18), who reported 7.8 x 1.1

Theriogenology

361

mm embryos recovered on Day 14 from superovulated cows. Our means included the 53 to 60% of embryos that were smaller than 1 mm*. Previous research has shown that in vitro produced bovine embryos at the blastocyst stage produce fewer cell lines when cultured than in vivo produced embryos (33). Likewise, the morphology of in vitro produced blastocysts is somewhat abnormal compared with in vivo embryos (25). Communication between the ovine uterus and the bovine embryo is probably not the same as for the ovine embryo and ovine uterus. Bovine in vitro produced bovine embryos failed to extend the estrous cycle of recipient ewes. These findings are unlike those of Heyman et al. (20), who found that bovine trophoblastic vesicles from cattle extend the estrous cycle of sheep, probably due to the production of boIFN-r, which, like oIFN-t, extends the life span of the bovine CL (24). The design of the present study may not have been adequate for determining the effect of bovine embryos on the lifespan of the ovine CL, as there was considerable variation in the development of the transferred embryos. On the other hand, the development of the embryos may have precluded an effect. In sheep, the presence of an embryo on Day 13 to 14 is critical for extending the estrous cycle, while in the cow Day 16 is critical (5, 14). The timing of maximal production of embryonic antiluteolytic factors by bovine embryos may not be concurrent with the time that the ovine CL requires the presence of an antiluteolytic factor to maintain the life-span of the CL. Our findings of a massive increase in boIFN-t from Days 14 to 16 substantiates this difference (2). On the other hand, when the cycle was extended by exogenous MAP, embryos were capable of elongating into a filamentous structure, suggesting that the sheep uterus provides appropriate nutrition or signals for pre-implantation development of the bovine embryos. The impetus for this study was the need for an economical method to produce bovine embryonic disks for transfer into SCID mice (34). This was done with the purpose of assaying the xenograft for formation of teratomas or teratocarcinomas in the kidney capsule or testicular capsule of the mice. Twenty-eight bovine embryonic disks (- 13 d) produced in sheep uteri were transferred into SCID mice with little or no resulting tumor formation. In comparison, blastocysts developed in vivo in cow uteri produced no tumors in 15 xenografis and 6 tumors in 27 xenografts of 7- and 14-d blastocysts, respectively (unpublished observations; 34). This result was similar to that obtained by Anderson et al. (l), who found tumor formation rates to be low or ml from bovine embryonic disks and inner cell masses of less than 14-d developmental age. It is possible, therefore, that the embryonic disks from in vitro produced/sheep-incubated bovine blastocysts were developmentally retarded or were not qualitatively equivalent to those produced entirely in vivo in the cow. The embryonic disks from the filamentous blastocysts (Figure lb) were not tested for tumor formation in SCID mice. This might have compared similarly to the in vivo 14-d embryonic disks because of their more advanced development and because the pattern of increasing teratoma formation with later developmental stage was also observed in the pig and sheep (1,34).

362

Theriogenology

The transfer of bovine in vitro produced embryos into the sheep uterus may provide a new and relatively economical model for the development of bovine embryos during the critical phase of elongation and during the period of maternal recognition of pregnancy. The application of somatic cell nuclear transfer for creating transgenic animals was recently demonstrated in both cattle and sheep, and homologous recombination to knock out or modify specific genes in the bovine or ovine embryo is a possibility (10,30). Thus, the effects of experimental manipulations of bovine embryo developmental genes might be usemlly assayed in the sheep uterus.

REFERENCES 1. Anderson GB, BonDurant RI-I, Goff L, Groff J, Moyer AL. Development of bovine and porcine embryonic teratomas in athymic mice. Animal Reprod Sci 1996;45:23 l-240. 2. Ashworth CJ, Fuller WB Changes in ovine conceptus and endometrial function following asynchronous embryo transfer or administration of progesterone. Biol Reprod 1989;40:425434. 3. Bazer FW, Spencer TE, Ott TL. Interferon tau: a novel pregnancy recognition signal. Am J Reprod hmmmol 1997;37:412-420. 4. Betteridge KJ, FlCchon JE. The anatomy and physiology of pre-attachment bovine embryos. Theriogenology 1988;29: 155-187. 5. Bet&ridge KJ, Eaglesome MD, Randall GCB, Mitchell D. Collection, description, and transfer of embryos from cattle lo-16 days after oestrus. J Reprod Fertil 1980;59:205-216. 6. Bondioli KR, Beiry KA, Hill KG, Jones KB, De Mayo FJ. Production of transgenic cattle by microinjection. In: First NL and Haseltine FP (eds), Transgenic Animals. Boston: ButterworthHeinemann, 1991;265-273. 7. Butler JE, Hamilton, WC, Sasser RG, Ruder CA, Hass GM, Williams RJ. Detection and partial characterization of two bovine pregnancy-specific proteins. Biol Reprod 1982;26:925-933. 8. Cassida LE. Fertilization failure and embryonic death in domestic animals. In: Engle E (ed), Pregnancy Wastage. Springfield IL: Charles C. Thomas, 1953;27-37. 9. Chang MC. Development of bovine blastocyst with a note on implantation. Anat Ret 1952;113:143-161. 10. Cibelli JB, Stice SL, Golueke PJ, Kane JJ, Jerry J, Blackwell C, Ponce de Leon A, Rob1 JM. Cloned transgenic calves produced from nonquiescent fetal fibroblasts. Science 1998;280:1256-1258. 11. Cross JC, Werb Z, Fisher SJ. Implantation and the placenta: key pieces of the developmental puzzle. Science 1994;266:1508-1517. 12. Edwards JL, Ealy AD, Monterroso VH, Hansen PJ. Ontogeny of temperature-regulated heat shock protein 70 synthesis in preimplanation bovine embryos. Mol Reprod Dev 1997; 48:2533. 13. Eyestone WH, Leibtiied-Rutledge ML, Northey DL, Gilligan BG, First NL. Culture of oneand two-cell bovine embryos to the blastocyst stage in ovine oviducts. Theriogenology 1987;28:1-7. 14. Farin CE, Imakawa K, Hansen TR, McDonnell JJ, Murphy CN, Farin PW, Roberts RM. Expression oftrophoblastic interferon genes in sheep and cattle. Biol Reprod 1990;43:210-218.

Theriogenology

363

15. Gharib-Hamrouche N, ChiZne N, Guillomot M, Martal J. Localization and characterization of EGF/TGF-cl receptors on perimplautation trophoblast in sheep. J Rcprod Fertil 1993;98:385392. 16. Glover MD, Seidel GE, Jr. Allograft inflammatory factor-l mRNA increases in bovine embryos between Days 15 and 17 of gestation. Theriogenology 1998;49:271 abstr. 17. Guillomot M. Cellular interactions during implantation in domestic ruminants. J Rcprod Fertil 1995; 49:39-51. 18. Grealy M, Diskin MG, Sreenan JM. Protein content of cattle oocytes and embryos from the two-cell to the elongated blastocyst stage at Day 16. J Reprod Fertil 1996;107:229-233. 19. Hare WCD, Singh EL, Betteridge KJ, Eaglesome MD, Randall GCB, Mitchell D, Bilton RJ Trounson AO. Chromosomal analysis of 159 bovine embryos collected 12 to 18 days after estrus. Can J Genet Cytol 1980;22:615-626. 20. Heyman Y, Camous S, Fevre J, Meziou W, Martal J. Maintenance of the corpus luteum after uterine transfer of trophoblastic vesicles to cyclic cows and ewes. J Rcprod Fertil1984,70:533. 21. Imakawa K, Anthony RV, Kazemi M, Marotti KR, Polites HG, Roberts RM. Interferon-like sequence of ovine trophoblast protein secreted by embryonic trophectoderm. Nature 1987;330:377-379. 22. Martal J, Chene N, Camous S, Huynh L, Lantier F, Hermier P, L’Haridon R, Charpigny G, Charlier M, Chaouat G. Recent developments and potentialities for reducing embryo mortality in ruminants: the role of IFN-r and other cytokines in early pregnancy. Reprod Fertil Dev 1997; 9:355-380. 23. Mathialagan N, Bixby JA, Roberts RM. Expression of intcrleukin-6 in porcine, ovine, and bovine preimplantation conceptuses. Mol Reprod Dev 1992;32:324-330. 24. Meyer MD, Hansen PJ, Thatacher WW, Drost M, Badinga L, Roberts RM, Li J, Ott TL, Bazer FW. Extension of corpus luteum life span and reduction of uterine secretion of prostaglandin, Fz= of cows in response to recombinant interferon-tau. J Dairy Sci 1995;78:1921-1931. 25. Plante L, King WA. Light and electron microscopic analysis of bovine embryos derived by in vitro and in vivo fertilization. J Assist Reprod Genet 1994; 11:5 15-529. 26. Roberts RM, Imakawa K, Niwano Y, Kazemi M, Malathy PV, Hansen TR, Glass AA, Kronenbert LH. Interferon production by the preimplantation sheep embryo. J Interferon Res 1989;9:175-187. 27. Rob1 JM, Prather R, Barnes FL, Eyestone WI-I, Northay D., Gilligan B, First N. Nuclear transplantation in bovine embryos. J Anim Sci 1987;64:642-647. 28. Rosekrans CF, First NL. Culture of bovine zygotes to the blastocyst stage: effects of amino acids and vitamins. Thcriogcnology 1991; 35:266 abstr. 29. SAS Institute, Inc., SAS/STAT@ Software: Changes and Enhancements, Release 6.10 Cary NC: SAS Institute Inc., 1994. 30. Schnieke AE, Kind AJ, Ritchie WA, Mycock K, Scott AR, Ritchie M, Wihnut I, Colman A, Campbell KHS. Human factor IX transgenic sheep produced by transfer of nuclei from transfected fetal tibroblasts. Science 1997;278:2130-2133.

364

Theriogenology

3 1. Stewart HJ, McCann SHE, Barker PJ, Lee KE, Lamming GE, Flint APE. Interferon sequence homology and receptor binding activity of ovine trophoblast antiluteolytic protein. J Endocrinol 1987;115:R13-R15. 32. Schultz GA, Hogan A, Watson AJ, Smith RM, Heyner S. Insulin, insulin-like growth factors and glucose transporters: temporal patterns of gene expression in early murine and bovine embryos. Reprod Fertil Dev 1992;4:361-371. 33. Talbot NC, Powell AM, Rexroad CE Jr. In vitro phrripotency of epiblast derived from bovine blastocysts. Mol Reprod Dev 1995;42:35-52. 34. Talbot NC, Powell AM, Rexroad CE Jr, Schuster M. Xenografis of livestock embryonic cells into mice with severe combined immunodeficiency. Theriogenology 1996;45:239 abstr. 35. Thatcher WW, Meyer MD, Danet-Desnoyers G. Maternal recognition of pregnancy. J Reprod Fertil 1995;49 (Suppl):15-28. 36. Watson AJ, Hogan A, Hahnel A, Wiemer KE, Schultz GA. Expression of growth factor ligand and receptor genes in the preimplantation bovine embryo. Mol Reprod Dev 1992;3 1:87-95. 37. Wrenzycki C, Herrmann D, Carnwath JW, Niemamr H. Expression of RNA from developmentally important genes in preimplantation bovine embryos produced in TCM supplemented with BSA. Reprod Fertil 1998;112:387-398. 38. Willadsen, SM. Micromanipulation of embryos of the large domestic species. In: Adams CE (ed), Mammalian Egg Transfer. Boca Raton FL CRC Press, 1982;185-210. 39. Willadsen SM, Polge C. Attempts to produce monozygotic quadruplets in cattle by blastomere separation Vet Ret 1981;108:211-213. 40. Xie S, Low BG, Nagel RJ, Kramer KK, Anthony RV, Zoli AP, Beckers JF, Roberts RM. Identification of the major pregnancy-specific antigens of cattle and sheep as inactive members of the aspartic proteinase family. Proc Nat1 Acad Sci (USA) 1991;88:10247-10251.