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2004 The chick mbre yo amnion as an in vitro culture system for IVF and NT embryos Tonya Renea Davidson Louisiana State University and Agricultural and Mechanical College
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Recommended Citation Davidson, Tonya Renea, "The chick mbre yo amnion as an in vitro culture system for IVF and NT embryos" (2004). LSU Doctoral Dissertations. 1280. https://digitalcommons.lsu.edu/gradschool_dissertations/1280
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THE CHICK EMBRYO AMNION AS AN IN VITRO CULTURE SYSTEM FOR IVF AND NT EMBRYOS
A Dissertation
Submitted to the Graduate Faculty of Louisiana State University and Agricultural and Mechanical College In partial fulfillment of the Requirements for the degree of Doctor of Philosophy
in
The Interdepartmental Program in Animal and Dairy Sciences
by Tonya Renea Davidson B.S., Oklahoma State University, 1997 M.S., Oklahoma State University, 2000 December 2004
ACKNOWLEDGEMENTS
I would like to begin by dedicating this dissertation to my mom, Linda Millspaugh. Mom, you have always been my role model, my support system and my foundation. You taught me that I could be whatever I wanted to be, and you had faith in me even when I didn’t have faith in myself. I am so proud to be your daughter. I would also like to dedicate this dissertation to my brother, Casey Millspaugh. Thank you Casey, for always believing in me and looking up to me. That has been more encouragement than you will ever know. I would like to express me sincere gratitude to my major professor, Dr. Robert Godke. Thank you for allowing me to pursue my Ph.D. at LSU. You have provided me with a firm scientific background, but more importantly, you have encouraged me to stand up for that which I believe. You have been a wonderful mentor and I appreciate all that you have done for me. To Dr. Leibo, Dr. Lynn and Dr. Williams. Thank you so much for serving on my committee, and for all of the assistance you have provided to make my time at LSU such a success. It has been a great pleasure working with all of you. I have learned so much from each of you and I am a better scientist for having known you. To Trey Chapman, Kyle Hebert, Laurie Henderson, Andrea Huval, Angela “Beanie” Klumpp, Luke Lenard, Alison Mosian, Kyle Hebert, Dr. Masao Murakami and James “Par” Sterling. Thank you for all that you have done for me. You have each touched my life in a very special way and I will always remember you. To Kristy Bruce. Thank you for saying “Hello” that first day of class. Since that time, you have been a wonderful source of entertainment and support, but more importantly, you have been an exceptional friend. I am so much better for having known you. And finally, to Edward Ferguson. My words can not adequately express how I truly feel for you. You are my best friend and the love of my life. You have never left my side, no matter how hard things got, and for that I will always love you.
ii TABLE OF CONTENTS
ACKNOWLEDGEMENTS ...... ii
LIST OF TABLES...... vi
LIST OF FIGURES...... viii
ABBREVIATIONS USED IN THE DISSERTATION...... xi
ABSTRACT...... xii
CHAPTER I. INTRODUCTION ...... 1
CHAPTER II. LITERATURE REVIEW ...... 3 In Vitro Production of Embryos...... 3 Production of Bovine Embryos by In Vitro Fertilization...... 3 Production of Bovine Embryos by Nuclear Transfer...... 4 In Vitro Embryo Culture ...... 6 Influence of In Vitro Production on Embryo Development...... 11 Embryo Development Rates...... 11 Embryo Morphology ...... 12 Pregnancy and Calving Rates ...... 14 Large Offspring Syndrome and Associated Disorders...... 16 Apoptosis ...... 18 Chick Embryo Co-Culture...... 24 Biochemistry of the Avian Egg...... 24 Early Use of Chick Embryo Extracts in Mammalian Cell Culture ...... 25 In Vitro Culture of Chick Embryos ...... 26 Chick Embryo Co-Culture...... 26
CHAPTER III. CULTURING IN VITRO PRODUCED BOVINE EMBRYOS IN THE AMNIOTIC CAVITY OF THE DEVELOPING CHICKEN EMBYRO...... 28 Introduction...... 28 Materials and Methods...... 29 Experimental Design ...... 29 Experimental Methods...... 31 Statistical Analysis...... 36 Results ...... 36 Discussion...... 41
CHAPTER IV. USE OF THE AMNIOTIC CAVITY OF THE DEVELOPING CHICK EMBRYO FOR CULTURING BOVINE 8-CELL EMBRYOS DERIVED FROM SOMATIC CELL NUCLEAR TRANSFER ...... 46 Introduction ...... 46 Materials and Methods...... 47 Experimental Design ...... 47 Experimental Methods...... 49
iii Statistical Analysis...... 53 Results ...... 53 Discussion...... 54
CHAPTER V. THE EFFECT OF ALTERATIONS IN THE NORMAL ANGIOGENIC PATTERN OF THE CHICK CHORIOALLANTOIC MEMBRANE ON BOVINE EMBRYO DEVELOPMENT AFTER CHICK EMBRYO CO-CULTURE...... 60 Introduction ...... 60 Materials and Methods ...... 61 Experimental Design ...... 61 Experimental Methods...... 63 Statistical Analysis...... 69 Results...... 69 Discussion...... 78
CHAPTER VI. DEVELOPMENT OF 8-CELL BOVINE EMBRYOS IN CULTURE MEDIUM SUPPLEMENTED WITH CHICK AMNIOTIC FLUID (CAF).. 80 Introduction...... 80 Materials and Methods...... 81 Experimental Design ...... 81 Experimental Methods...... 81 Statistical Analysis...... 85 Results ...... 86 Discussion...... 86
CHAPTER VII. ESTABLISHMENT AND CULTURE OF CELLS DERIVED FROM DAY-6 CHICK EMBRYOS...... 93 Introduction...... 93 Materials and Methods...... 95 Experimental Design ...... 95 Experimental Methods...... 95 Statistical Analysis...... 101 Results ...... 102 Discussion...... 107
CHAPTER VIII. THE INCIDENCE OF APOPTOSIS IN BOVINE EMBRYOS AFTER IN VITRO CULTURE AND CHICK EMBRYO CO-CULTURE...... 112 Introduction...... 112 Materials and Methods...... 113 Experimental Design ...... 113 Experimental Methods...... 115 Statistical Analysis...... 122 Results ...... 122 Discussion...... 129
SUMMARY AND CONCLUSIONS...... 138
iv LITERATURE CITED...... 140
APPENDIX A: BO-A STOCK SOLUTION ...... 170
APPENDIX B: BO-B STOCK SOLUTION ...... 171
APPENDIX C: CR1aa STOCK SOLUTION ...... 172
VITA ...... 173
v LIST OF TABLES
3.1. Development of 1-cell bovine embryos cultured in CR1aa + 5% fetal bovine serum (control) or co-cultured in the chick amnion from day 0 to day 3 (CEC-I) or from day 3 to day 7 (CEC-II)...... 38
3.2. Development of 2-cell bovine embryos cultured in CR1aa + 5% fetal bovine serum (control) or co-cultured in the chick amnion from day 0 to day 3 (CEC-I) or from day 3 to day 7 (CEC-II)...... 40
4.1. Day 6 development of somatic cell nuclear transfer bovine 8-cell embryos co- cultured in the chick embryo amnion...... 55
5.1. Mortality of chick embryos after treatment application and subsequent 24 hour incubation...... 70
5.2. Effect of cytochalasin D, ethanol, PGE2 and PGF2α on capillary branching in the chorioallantoic membrane of the chick embryo. Treatment effects were quantified by counting the number of nodes in four random areas of each CAM ...... 71
5.3. Development of 8-cell bovine embryos cultured in CR1aa + 5% fetal bovine serum (control), co-cultured in the chick embryo amnion co-cultured in the amnion of PGE2-treated chicks ...... 75
6.1. Development of 8-cell bovine embryos cultured in CR1aa supplemented with 5% fetal bovine serum (control) or 20% chick amniotic fluid (CAF) ...... 87
7.1. Experimental design for Experiment 7.1 for growth and proliferation of fetal chick cells...... 96
7.2. Experimental design for Experiment 7.2 for growth and proliferation of chick amnion cells ...... 97
7.3. Growth characteristics of fetal chick cells during in vitro culture in mTCM and mDMEM culture media...... 103
7.4. Growth characteristics of chick amnion cells during in vitro culture in mTCM and mDMEM culture media...... 104
8.1. Response of cows to synchronization/superovulation protocol...... 123
8.2. Number, stage and quality grade of embryos recovered from superovulated cows...... 125
8.3. Effect of culture environment on mean number of apoptotic cells in morula, early blastocyst and blastocyst stage embryos ...... 126
vi 8.4. Effect of culture environment on mean number of apoptotic cells in morula, early blastocyst and blastocyst stage embryos...... 127
8.5. Effect of culture environment on apoptotic index in morula, early blastocyst, and blastocyst stage embryos...... 128
8.6. Effect of culture environment on mean number of apoptotic cells in grade 1, grade 2, grade 3, grade 4 and degenerate embryos...... 130
8.7. Effect of culture environment on mean number of apoptotic cells in grade 1, grade 2, grade 3, grade 4 and degenerate embryos...... 131
8.8. Effect of culture environment on apoptotic index in grade 1, grade 2, grade 3 and grade 4 embryos ...... 132
vii LIST OF FIGURES
3.1. Experimental design for the culture of 1-cell bovine embryos in the amnion of a developing chick embryo...... 30
3.2. Experimental design for the culture of 2-cell bovine embryos in the amnion of a developing chick embryo...... 32
3.3. Development of 1-cell bovine embryos cultured in CR1aa + 5% fetal bovine serum (control) or co-cultured in the chick amnion from day 0 to day 3 (CEC-I) or from day 3 to day 7 (CEC-II) on day 3 of development (A) and on day 7 of development (B)...... 39
3.4. Development of 2-cell bovine embryos cultured in CR1aa + 5% fetal bovine serum (control) or co-cultured in the chick amnion from day 0 to day 3 (CEC-I) or from day 3 to day 7 (CEC-II) on day 3 of development (A) and on day 7 of development (B)...... 42
3.5. Quality grade of bovine 2-cell embryos cultured in CR1aa + 5% fetal bovine serum (control) or co-cultured in the chick amnion from day 0 to day 3 (CEC-I) or from day 3 to day 7 (CEC-II) on day 7 of development ...... 43
4.1. Experimental design for the culture of bovine 8-cell embryos derived from somatic cell nuclear transfer. Culture was either in vitro in CR1aa supplemented with 5% fetal bovine serum (control) or co-cultured in the chick embryo amnion...... 48
4.2. Bovine somatic cell nuclear transfer embryos developed into morulae/blastocysts (day 6 of in vitro culture) after co-culture in the amniotic cavity of a developing chick embryo for 3 days. These embryos were agarose embedded (agar cylinders are indicated by arrows) at the 8-cell stage following in vitro culture in CR1aa medium from day 0 to day 3 of in vitro culture and then injected into the amnion of a 96-hour live chick embryo ...... 56
4.3. Ultrasound image of a fetus (day 30) derived from bovine somatic cell nuclear transfer embryos that were co-cultured in the amnion of a developing chick embryo from day 3 to day 6 following in vitro culture in CR1aa medium ...... 56
5.1. Experimental design to evaluate the effects of PGE2 and PGF2α on capillary development in the chick chorioallantoic membrane ...... 62
5.2. Experimental design for the culture of 8-cell bovine embryos cultured in CR1aa + 5% fetal bovine serum (control), co-cultured in the chick embryo amnion (CEC) or co-cultured in the amnion of PGE2-treated chicks (PGE) ...... 64
5.3. Restructuring of vasculature networks in CAMs treated with (A) chick Ringer’s solution (control), (B) ethanol (vehicle), (C) PGE2 and (D) PGF2α...... 72
viii
5.4. Effect of cytochalasin D, ethanol (EtOH), PGE2 (PGE) and PGF2α (PGF) on capillary branching in the chorioallantoic membrane of the chick embryo. The number on each bar indicates the total number of samples (four samples per CAM) in each treatment group ...... 73
5.5. Development of 8-cell bovine embryos cultured in CR1aa supplemented with 5% fetal bovine serum (control), cultured in the chick embryo amnion (CEC) or cultured in the amnion of PGE2-treated chicks (PGE) on day 7 of development (A), day 8 of development (B) and day 9 of development (C)...... 76
5.6. Quality grade of bovine embryos cultured in CR1aa supplemented with 5% fetal bovine serum (control), cultured in the chick embryo amnion (CEC) or cultured in the amnion of PGE2-treated chicks (PGE) on day 7 of develop- ment (A), day 8 of development (B) and day 9 of development (C)...... 77
6.1. Experimental design for culturing 8-cell bovine embryos in culture medium supplemented with chick amniotic fluid. In vitro culture was either in CR1aa supplemented with 5% fetal bovine serum (control group) or CR1aa supplemented with 20% chick amniotic fluid (CAF group) ...... 82
6.2. Development of 8-cell bovine embryos cultured in CR1aa supplemented with 5% fetal bovine serum (control) or 20% chick amniotic fluid (CAF) on day 7 of development (A), day 8 of development (B) and day 9 of development (C). .. 88
6.3. Quality grade of 8-cell bovine embryos cultured in CR1aa supplemented with 5% fetal bovine serum (control) or 20% chick amniotic fluid (CAF) on day 7 of development (A), day 8 of development (B) and day 9 of development (C)...... 89
7.1. Growth of chick amnion cells in vitro in mTCM culture medium...... 105
7.2. Growth of chick amnion cells in vitro in mDMEM culture medium...... 106
7.3. Cell populations of chick amnion cells cultured in vitro in mTCM during the first passage...... 108
7.4. Cell populations of chick amnion cells cultured in vitro in mTCM during the second passage ...... 109
7.5. Cell populations of chick amnion cells cultured in vitro in mTCM after freezing and thawing...... 110
8.1. Experimental design for the production and fluorescent labeling of in vivo- derived embryos...... 114
8.2. Experimental design for the production and fluorescent labeling of embryos cultured in vitro ...... 116
ix
8.3. Experimental design for the production and fluorescent labeling of embryos cultured in the chick embryo amnion...... 117
8.4. Localization of apoptosis staining to the ICM of a grade 2 frozen in vivo- derived bovine blastocyst...... 135
8.5. The presence of different degrees of fluorescence within an embryo. Pinpoint areas of fluorescence are thought to represent cells in the early stages of apoptosis with only small amounts of caspase present in the cytoplasm (A), whereas extensive fluorescence are thought to represent cells in more advanced stages of apoptosis with much more caspase present in the cell (B) ...... 136
x
ABBREVIATIONS USED IN THE DISSERTATION
AI - artificial insemination BSA - bovine serum albumin BVDV - bovine viral diarrhea virus CAF - chick amniotic fluid CAM - chorioallantoic membane CEC - chick embryo co-culture CIDR - control internal drug release CL - corpus luteum DMEM - Dulbecco’s modified eagle’s medium DPBS - Dulbecco’s phosphate buffered saline ET - embryo transfer EtOH - ethanol FBS - fetal bovine serum FCS - fetal calf serum FSH - follicle stimulating hormone ICM - inner cell mass IGF-I - insulin-like growth factor-I IGF-II - insulin-like growth factor-II IGF2R - insulin-like growth factor-II receptor IVF - in vitro fertilization IVM - in vitro maturation IVP - in vitro produced M-II - metaphase-II mBMOC - modified Brinster medium for ovum culture MOET - multiple ovulation embryo transfer NT - nuclear transfer P4 - progesterone PGE2 - prostaglandin E2 PGF2α - prostaglandin F2α PMSG - pregnant mare serum gonadotropin POEC - porcine oviduct epithelial cells SOFaa - synthetic oviduct fluid with amino acids TCM - tissue culture medium TE - trophoectoderm TGF-α - transforming growth factor-α TNF-α - tumor necrosis factor-α TNF-R1 - tumor necrosis factor receptor I TUNEL - TdT-mediated dUTP nick-end labeling WOW - well of the well
xi
ABSTRACT
Calving rates are significantly reduced following in vitro production of embryos. Thus, if a technique could be developed that would increase calving rates by as little as one viable offspring, significant research advances could be made. Therefore, in a series of experiments, the efficiency and quality of culturing IVP bovine embryos in the amnion of a domestic chicken egg was tested. In Experiment I, by culturing IVP bovine embryos in the chick amnion (day 4 to 7 of incubation) it was discovered that there was no significant difference in blastocyst rates compared with controls. In Experiment II, it was shown that nuclear transfer bovine embryos cultured in the chick amnion reached the blastocyst stage at rates equal to controls and were capable of producing pregnancies following transplantation into recipient females. In a subsequent experiment, a method of naturally improving the chick embryo co-culture (CEC) system was explored by treating the developing chick embryo with prostaglandin E2 (PGE2) or prostaglandin F2α (PGF2α).
It was determined that treatment with PGE2 increased angiogenesis within the developing chicken egg, while treatment with PGF2α decreased angiogenesis. When bovine embryos were cultured in PGE2-treated chicks, developmental rates were not increased. In Experiment IV, chick amniotic fluid (CAF) was evaluated as a media supplement to the control culture system. Although replacing fetal bovine serum (FBS) with CAF resulted in significantly fewer blastocysts on day 7 of culture, there were no significant differences in the number of grade 1 embryos between the two treatments. This finding was important because it demonstrated an ability to culture IVP bovine embryos in the absence of FBS, a medium component that has been implicated in numerous fetal and calf abnormalities. Another experiment was designed to develop a method of culturing cells derived from the chick embryo and surrounding amniotic membrane for later use as a co-culture system for bovine IVP embryos. In the final experiment, using a novel method to detect apoptotic cells, it was determined that CEC did not alter the number of apoptotic cells in the embryo when compared with in vivo- derived or in vitro-cultured bovine embryos.
xii
CHAPTER I
INTRODUCTION
With the advent of in vitro fertilization technologies in the 1980s came the need for methods of culturing embryos to later stages of development. As early as 1975, Dr. R.A. Godke envisioned the use of avian eggs for the culture of mammalian embryos (personal communication). Early attempts using unfertilized hen’s eggs proved largely unsuccessful (Blakewood et al., unpublished data), but after several months of trial and error a technique was developed that used shell-less chick embryos for the co-culture of mammalian embryos. In the initial report, pronuclear stage murine embryos were embedded in agarose and injected into the amniotic cavity of a 96-hour chick embryo. After 72 to 96 hours of incubation in the chick amnion, significantly more embryos had developed to the hatching blastocyst stage compared with those cultured in control medium alone (Blakewood et al., 1989a). Further studies demonstrated that the chick embryo co-culture system could support development of porcine (Ocampo et al., 1993), caprine (Blakewood et al., 1989a) and bovine (Blakewood et al., 1989b) embryos at rates equal to or better than controls. In vitro culture techniques have progressed significantly since these early experiments. Researchers have converted from culturing domestic animal embryos in modified Tissue Culture Medium (mTCM) and Ham’s F-10 to culturing embryos in media such as M2, CR1aa, CR2aa, NCSU-23, KSOM, SOFaa and even sequential mediums such as G1/G2 and PPM1/PPM2. Pregnancy rates and calving rates do not appear to be significantly different in sheep, pigs or cattle when embryos are cultured in these different media. These observations suggest that the culture media may not be a major factor in the increased abortion rates, increased placental problems and decreased calving rates often observed after the transfer of embryos produced in vitro. A more likely cause of the decreased calf production observed after transfer of in vitro fertilized and cultured embryos is the use of fetal bovine serum (FBS) as a media supplement. FBS is added to bovine embryo culture media (as with many other species) to stimulate formation of the blastoceol and to increase the rate of blastocyst development. Although there have been serum free media developed (known as defined culture media), the blastocyst rates obtained with these media are not sufficient enough
1
to merit the use of in vitro fertilization and culture to produce offspring (Pinyopunnintr and Bavister, 1991; Bavister, 1995). Although serum is necessary for the efficient production of bovine blastocysts for transfer, a number of studies have cited the use of serum as a cause for large offspring syndrome, a collection of fetal and calf abnormalities often observed following the transfer of in vitro-produced ovine and bovine embryos (Farin et al. 2000; Hasler et al., 2000). Among these reports it was suggested that the presence of serum in the culture medium resulted in significantly more embryo transfer recipients aborting during the later stages of pregnancy and a higher rate of calf mortality. This effect has also been reported when embryos were cultured in media supplemented with estrous cow serum or serum derived from other species (Kruip and den Daas, 1997; Agca et al., 1998; Hasler et al., 2000). Large offspring syndrome has also been reported following transfer of embryos co-cultured with different cell types, such as buffalo rat liver cells, oviduct epithial cells, uterine epithelial cells and fibroblast cells (Sinclair et al., 1997). Although co-culture was first used as a means to allow in vitro-produced embryos to develop beyond the developmental block stage, neither its use nor supplementing media with serum has improved calving rates compared with multiple ovulation and embryo transfer. These problems do not seem to be inherent only to embryos produced by in vitro fertilization and culture. Pregnancies produced following the transplantation of nuclear transfer (NT)-derived embryos to recipient females have an even higher rate of large offspring syndrome (Kruip and den Daas, 1997; Barnes, 2000). Although it has been suggested that manipulation of the genetic material of the oocyte may be the primary cause of large offspring syndrome with NT-derived embryos, the common link between those embryos and in vitro-produced embryos is the in vitro culture medium and the use of serum as a protein supplement. Collectively, these findings indicate a need for an alternative in vitro culture system that is capable of producing blastocyst rates at an acceptable level and that may reduce or alleviate the occurrence of large offspring syndrome following in vitro embryo production. Therefore, the objective of this dissertation is to re-evaluate the use of the chick embryo co-culture system as an in vitro culture system for bovine embryos.
2 CHAPTER II
LITERATURE REVIEW
IN VITRO PRODUCTION OF EMBRYOS Production of Bovine Embryos by In Vitro Fertilization Prior to the 1980s early attempts to produce live offspring from in vitro-fertilized bovine oocytes were largely unsuccessful. Initial success with bovine in vitro fertilization (IVF) came in 1977 when Iritani and Niwa published the first report of IVF using in vitro- matured (IVM) bovine oocytes. Much of the research in the following years was dedicated to the study of the mechanics of fertilization. For example, Brackett et al. (1980a) observed that sperm penetration (19 to 24 hours post-insemination) was favored by in vivo-matured ova when compared with immature ova (recovered from 2- to 5-mm follicles) matured in vitro for 18 to 25 hours prior to fertilization. In addition, Brackett et al. (1980b) observed similarities, such as loss of cortical granules and the presence of sperm remnants in oocytes and early embryos, between in vivo- and in vitro-fertilized oocytes. As with the development of IVF procedures in laboratory animals, preparation of bovine sperm, particularly capacitation, proved to be a major challenge (Brackett, 2001). Early methods of capacitating spermatozoa involved incubation of spermatozoa in the oviduct or uterus of estrual females (Iritani and Niwa, 1977). However, by 1984 fresh ejaculated spermatozoa could be capacitated in a chemically defined medium (Iritani et al., 1984). By 1986, proteooglycans and glycosoaminoglycans (GAGs), capacitating agents present at the site of fertilization in vivo (for review see Miller and Ax, 1990) were determined to effectively induce the acrosome reaction and capacitate spermatozoa in vitro (Parrish et al., 1985, 1986, 1988, 1989). Other agents used to capacitate spermatozoa in vitro have included hyaluronic acid (Fukui, 1990), a combination of heparin and caffeine (Niwa and Ohgoda, 1998), calcium ionophore A 23187 with or without caffeine (Hanada et al., 1985; Goto et al., 1988) and calcium-free Tyrodes (Ijaz and Hunter, 1989). These studies culminated in the production of the first IVF calf in June, 1981 and publication of the first repeatable protocol for bovine IVF in 1982 (Brackett et al., 1982). These results were confirmed in 1984 with the birth of twin calves following in vitro
3 fertilization of in vivo-matured oocytes (Brackett et al., 1984). In these reports oocytes were matured in vivo following hormonal stimulation of donor females with either pregnant mare serum gonadotropin (PMSG) or follicle stimulating hormone (FSH) and following IVF, presumptive zygotes were surgically transferred to the oviducts of recipient females. Because in vivo-matured oocytes were collected in the same manner (i.e., surgically) as early stage embryos, it became necessary to develop effective in vitro maturation techniques for IVF to be considered a practical approach to the production of bovine embryos. Two decades later, much research is still focused on the successful maturation of oocytes in vitro. Production of Bovine Embryos by Nuclear Transfer In 1997, Ian Wilmut et al. announced the birth of Dolly, the first live ‘clone’ of an adult animal (Wilmut et al., 1997). The concept of cloning, however, originated long before the birth of Dolly. As early as the 1880s, scientists sought the answer to one fundamental question of biology – how does the nucleus control cell specialization during embryogenesis (Di Berardina, 2001). To answer this question, scientists began to examine the development of early stage embryos. In 1892, Driesch (reviewed in Morgan, 1934; Speman, 1938; Di Berardina, 2001) separated 2-cell sea urchin embryos in calcium-free seawater and found that in most cases each separate cell could develop into a complete larva. Continued experiments in other species, including amphibians, fish and mammals, yielded similar results. In 1894, Loeb (reviewed in Di Berardina, 2001) conducted a primitive cloning experiment using sea urchin eggs. In this study, the sea urchin eggs were placed in a hypotonic solution, causing a break in the vitelline membrane and subsequent herniation of a portion of the zygote through the break. In this particular study, the herniated portion of the zygote lacked a nucleus. As cell division progressed in the nucleated portion of the zygote, a daughter nucleus migrated into the non-nucleated potion of the zygote, resulting in two whole and separate sea urchin embryos. Hans Spemann (1914; reviewed by Di Berardina, 2001) extended Loeb’s research into amphibians. In this study, Spemann (1914) bisected newt eggs with a baby hair from his own son’s head. As in Loeb’s study, only one portion of the constricted egg contained a nucleus. Development proceeded as normal in the nucleated side of the
4 egg, but the non-nucleated side failed to cleave. Once the nucleated portion had developed to the 8- to 16-cell stage, he removed the ligature, allowing a nucleus to cross over into the enucleated side and then severed the two halves. As development continued, he found that each half could develop into a complete larva, thus proving that an embryonic cell was capable of programming larval development. Soon thereafter Robert Briggs set out to determine if the nuclei of somatic cells could program development in the same manner that zygotic nuclei could. Many thought this to be a “harebrained” idea, but in 1952 Briggs and King successfully produced tadpoles from blastula nuclei injected into enucleated frog eggs, creating the prototype for today’s nuclear transfer procedure (Briggs and King, 1952). It was not for another 30 years after the initial successes of Briggs and King that the first mammal was successfully cloned. The first report of live offspring after nuclear transfer was that of Illmensee and Hoppe (1981) using inner cell mass (ICM) cells of mouse blastocysts injected into an enucleated murine zygote. However, these results could not be replicated for almost 17 years. Tsunoda and Kato (1998) were eventually able to produce live offspring from both ICM and trophoblast cells. However, to accomplish this they had to use different host cells and a serial nuclear transfer procedure. The first uncontested cloned offspring were produced by Willadsen in 1986. He produced three lambs by fusing enucleated metaphase-II (M-II) oocytes to a single blastomere derived from an 8- to 16-cell embryo. These results were verified by the production of live calves from blastomeres of 4- to 16-cell embryos (Prather et al., 1987). Following these initial successes, cloned offspring have been produced from embryonic cells of many different species including rabbits (Collas and Robl, 1991), pigs (Prather et al., 1989), sheep (Campbell et al., 1996; Wilmut et al., 1997), goats (Yong et al., 1991), cattle (Sims and First, 1993; Collas and Barnes, 1994; Keefer et al., 1994) and rhesus monkeys (Meng et al., 1997). After the success of nuclear transfer with embryonic cells, researchers began to question if cloned offspring could be produced from adult cells. Early attempts to clone an adult animal using differentiated cells were completely unsuccessful (Di Berardina, 2001). However with the birth of Dolly, the belief that it was ‘biologically impossible’ to clone a mammal by nuclear transfer was dispelled (McGrath and Solter, 1984).
5 Furthermore, the production of a live offspring from totipotent mammary tissue (as with Dolly) suggested that other types of somatic cells could be used for cloning adult animals (Edwards et al., 2003). These findings were confirmed in mice with cumulus cells (Wakayama et al., 1998), pigs with granulosa cells (Polejaeva et al., 2000), goats with cumulus granulosa cells (Zou et al., 2001) and cattle with cumulus and oviduct cells (Kato et al., 1998). Cloned offspring have since been produced from many different cell types including leukocytes (Galli et al., 1999), muscle (Shiga et al., 1999), mammary tissue (Zakhartchenko et al., 1999; Kishi et al., 2000), ear tissue (Zakhartchenko et al., 1999), skin cells (Hill et al., 2000a; Kubota et al., 2000) and sertoli cells (Ogura et al., 2000) and from many different species including mice (Wakayama et al., 1998), rabbits (Chesne et al., 2002), pigs (Polejaeva et al., 2000), sheep (Wilmut et al., 1997), goats (Zou et al., 2001), cattle (Kato et al., 1998; Zakhartchenko et al., 1999; Hill et al., 2000a; Kishi et al., 2000; Kubota et al., 2000; Kasinathan et al., 2001; Heyman et al., 2002) and cats (Shin et al., 2002). In Vitro Embryo Culture Attempts to culture embryos outside the protective environment of the uterus date as far back as 1912 when Brachet attempted to culture 5- to 7-day rabbit embryos in coagulated blood plasma (referenced in Kane, 2003) and in the later study of Lewis and Gregory (1929) who reported developing rabbit embryos in vitro. The first successful report of in vitro embryo development came when Hammond (1949) reported that 8-cell mouse embryos could develop to the blastocyst stage in vitro when cultured in a simple saline solution supplemented with chicken egg white and yolk. During this same period, Chang (1949) reported that heat inactivated serum could be used as a supplement in culture medium for 2-cell rabbit embryos. The first culture medium specifically designed for mammalian embryos was described by Whitten (1957). He observed that a simple Kreb’s Ringer’s bicarbonate solution supplemented with serum albumin could promote development of 2-cell mouse embryos to the blastocyst stage (Whitten, 1957). McLaren and Biggers (1958) later reported that these embryos could result in live offspring after transfer to a surrogate female. The most important outcome of this research, however, was the fundamental discovery that early stage mouse embryos required lactate as an energy source for continued development (Bavister, 1995; reviewed by Kane, 2003). This crucial
6 observation paved the way for future research to examine the precise substrate requirements of pre-implantation mammalian embryos (Bavister, 1995). Progress in in vitro embryo culture was relatively slow for the next 20 years as attempts to support in vitro development of embryos from species other than mice and rabbits met with limited success (Bavister, 1995). It became obvious that embryos of most species suffered from an ‘in vitro developmental block’ that prevented further pre- implantation development in vitro (Bavister, 1995). These blocks were observed to occur at a characteristic stage of development in each species. Early attempts at circumventing the ‘block’ initially involved culturing in vitro- produced embryos in the ligated oviduct of a rabbit. This technique has been used extensively for the culture of embryos of many different species including pigs (Polge et al., 1972), sheep (Averill et al., 1955; Lawson et al., 1972), cattle (Sreenan et al., 1968; Boland, 1984; Ectos et al., 1993) and horses (Allen et al., 1976). Similarly, hamster embryos have been cultured in the mouse oviduct (Minami et al., 1988), pig embryos have been cultured in the sheep oviduct (Prather et al., 1991) and cattle embryos have been cultured in the oviducts of sheep (Eyestone et al., 1987; Galli and Lazzari, 1996; Enright et al., 2000) and mice (Krisker et al., 1989; Sharif et al., 1991). It is interesting to note that despite the advances of embryo culture in recent years, current research indicates that modern culture media fail to produce embryos of the same quality as those cultured in the oviduct of an intermediary recipient. Lonergan et al. (2001) reported that IVM-IVF embryos cultured in the ewe oviduct survived vitrification and warming much better than IVM-IVF embryos cultured in vitro in SOF (88.0% and 5.6% hatched blastocysts, respectively). In a reciprocal experiment, in vitro culture of in vivo-derived presumptive zygotes had a significantly lower post-warming survival and hatching rate than did in vivo-derived presumptive zygotes cultured in the ewe oviduct (Lonergan et al., 2001). These findings have been corroborated in sheep (Tervit et al., 1994) and cattle (Holm et al., 1994; Galli and Lazari, 1996; Enright et al., 2000; Jimeneze et al., 2001; Pugh et al., 2001; Rizos et al., 2002). Although oviductal culture allowed development of in vitro-produced embryos through the ‘block’ it was both impractical and expensive; thus, there remained a need for a way to culture embryos completely in vitro. In 1965, Cole and Paul reported that a high percentage of mouse embryos could develop and hatch in vitro when cultured on
7 HeLa cells (Cole and Paul, 1965). This classic study provided a more efficient method for maintaining embryos in vitro for extended periods of time with minimal reductions in viability (Rexroad, 1989; Thibodeaux and Godke, 1995) and laid the foundation for many of the embryo co-culture systems still in use today (Thibodeaux and Godke, 1995). Since the classic study of Cole and Paul (1965) many different cell types have been found to support mammalian embryo development in vitro. One of the earliest reports of embryo co-culture was that of Kuzan and Wright (1982) who reported that a higher percentage of embryos developed to the hatched blastocyst stage when cultured on either uterine or testicular fibroblasts when compared with culture medium alone. Later, Weimer et al. found that fetal uterine fibroblasts could enhance development of bovine (1987a,b) and equine (1989a,b) embryos. Rexroad and Powell (1986, 1988) were among the first to report that oviduct epithelial cells could be used to culture early stage ovine embryos and soon thereafter, Gandolfi and Moor (1987) reported that culture of early stage ovine embryos on oviduct epithelial cells resulted in more expanded blastocysts and a higher pregnancy rate than when embryos were cultured on uterine fibroblasts. Similarly, Eyestone and First (1989) reported a higher percentage of morula and blastocyst stage bovine embryos when IVF-derived 1-cell embryos were cultured on oviduct cells than when cultured in medium alone (22% vs 3%, respectively). In 1988, Goto et al. reported the establishment of viable pregnancies following in vitro culture of IVF-derived bovine embryos on bovine granulosa cells. In a similar experiment, Fukuda et al. (1989) reported live offspring from both fresh and frozen- thawed bovine embryos cultured on cumulus cells. Interestingly, bovine cumulus cells have been used successfully to culture embryos from other species including pigs (Zhang et al., 1990) and horses (Rodriquez et al., 1991). These findings corroborate the early findings of Baird et al. (1990), who reported that culturing mouse embryos on hamster cumulus cells significantly increased in vitro development when compared with culture medium alone. Many other cells types have been evaluated for use in embryo co-culture in addition to the ones described above. The work of Cole and Paul (1965) led others to use different types of helper cells, such as L cells, liver cells, JLS-V11 cells and teratocarcinoma cells (Glass et al., 1979). Others have used hamster hepatocytes (Overskei and Cincotta, 1987), buffalo rat liver cells (Hu et al., 1989) bovine fetal spleen
8 cells (Kim et al., 1989; Kim et al., 1991), chick embryo fibroblasts (Kim et al., 1989; Kim et al., 1991) and kidney (VERO and MDBK) cells (Ouhibi et al., 1990). Despite the extensive research conducted in this area researchers have yet to explain how co-culture cells benefit embryo development. Three hypotheses seem to exist: 1) cells detoxify the culture medium by removing harmful contaminates, such as heavy metal ions, 2) cells reduce the concentration of components in the medium that inhibit embryo development (e.g., glucose) and 3) cells secrete embryotropic factors into the culture medium, such as amino acids, pyruvate, proteins and growth factors (Bavister, 1995). It seems more likely, however, that the benefits of co-culture are a combination of all three hypotheses (Bavister, 1995). In recent years, the use of co-culture systems has largely given way to culture in semi-defined or defined culture media supplemented to varying degrees with amino acids, vitamins, serum and/or other compounds. The first medium formulated specifically for in vitro embryo culture (synthetic oviduct fluid, SOF) was a simple medium based on the concentrations of ions and carbohydrates present in ovine oviduct fluid (Tervit et al, 1972). With this medium it was possible to develop 1-cell bovine embryos up to the 16- cell stage and 8-cell bovine embryos up the blastocyst stage (Tervit, et al., 1972). This approach to embryo culture was largely ignored by the research community, who favored the more physiological environment of the rabbit or ewe oviduct for embryo culture (Gardner, 1999). It has not been the formulation of exceptional new media, but rather the small observations along the way, that have led to the advances in embryo culture technologies. One of the major, but largely unappreciated, milestones in embryo culture was the realization that IVM and IVF should occur at a temperature equal to the core body temperature of the animal (39°C in cattle; Lenz et al., 1983); this was later applied to sheep (Gandolfi and Moor, 1987; Fukui et al., 1988) and cattle (Fukui and Ono, 1988; Fukuda et al., 1990) embryo culture. Another very significant milestone in the development of in vitro embryo culture was that of Thompson et al. (1990). These researchers reported that development was significantly improved when embryos were cultured in an O2 concentration between 5 and 10% as opposed to the 20% O2 content of air.
9 During that same time period, Bavister and Arlotto (1990) first reported the benefits of amino acids during in vitro culture. Soon thereafter, amino acids were incororporated into mSOF (Takahasi and First, 1992), CR1 (Rosenkrans and First, 1994) and KSOM (Liu and Foote, 1995) culture media. In all three of these reports, addition of amino acids to the culture media resulted in significantly more embryos developing to the blastocyst stage. In addition, transfer of embryos cultured in amino acid supplemented SOF resulted in a 55% calving rate (Takahashi and First, 1992). With the routine addition of amino acids to culture media, Gardner et al. (1994) observed that embryos were particularly sensitive to ammonia produced by the spontaneous deamination of the amino acids and/or amino acid metabolism. Based on this observation, they proposed that media be removed and replaced with fresh medium every 48 hours to optimize embryo development. In hindsight this was a critical finding. Recent research indicates that chronic exposure of embryos to ammonium in culture impairs development (Gardner and Lane, 1993; Gardner et al., 1994), retards fetal growth (Lane and Gardner, 1994) and can lead to abnormal fetal development (Lane and Gardner, 1994; McEvoy et al., 1997; Sinclair et al., 1998). One could not thoroughly discuss in vitro embryo culture without mentioning the use of serum in in vitro culture media. Serum has been used throughout the history of embryo culture, both alone and as a supplement (reviewed in Wright and Bondioli, 1981). Early attempts at embryo culture utilized pure serum as a medium, however little to no development was reported (Pincus, 1951; Brock and Rowson, 1952). Later studies utilized serum as a supplement to the culture medium. In 1963, Hafez et al. cultured 1- cell bovine embryos in serum supplemented saline, but again, failed to achieve high rates of embryonic development. However, when Moore (1970) cultured 2- to 8-cell ovine embryos in Brinster’s Salt Solution supplemented with 15% ovine serum, 38 of 57 (66%) of the embryos cleaved and 4 of 30 (13%) resulted in pregnancies. In 1976, Wright et al. compared embryo development in several different media. Of the media tested, Ham’s F-10 with 50% heat treated fetal calf serum (FCS) was the only medium evaluated that was able to support development from the 8-cell stage to the expanded blastocyst stage (Wright et al., 1976a,b,c). This was the first report of bovine embryos expanding and hatching in vitro. Unbeknownst to them, this was likely the first report of the biphasic effect of serum. In 1993, Kolver and MacMillan reported
10 that serum was inhibitory to embryos when added during early cleavage but was stimulatory when present at the initiation of compaction. Many current in vitro culture systems have taken advantage of this to maximize blastocyst yield by leaving serum out of the culture medium during early development but adding it later in development (Carolan et al., 1995; Massip et al., 1996; Pinyopmmintr and Bavister, 1996a,b; Van Langendonkt et al., 1996). In spite of the apparent benefits of serum supplementation, recent evidence suggests that prolonged culture of embryos in the presence of serum alters embryo morphology and biochemistry (Gardner et al., 1994; Shamsuddin and Rodriquez- Martinez, 1994; Thompson et al., 1995). Serum has been implicated in precocious blastoceol formation (Walker et al., 1992; Thompson et al., 1995), sequestration of lipids (Thompson et al., 1995), abnormal mitochondrial ultrastructure (Thompson et al., 1995), perturbations in metabolism (Gardner et al., 1995), altered embryo morphology (Gardner, 1999), abnormally large offspring (Willadsen et al., 1991; Keefer et al, 1994; Behboodi et al., 1995; Thompson et al., 1995), increased gestation lengths (Walker et al., 1996) and increased perinatal (Behboodi et al., 1995; Wilson et al., 1995; Garry et al., 1996; Schmidt et al., 1996; Kruip and den Daas, 1997; Hill et al., 1999) and neonatal mortality (Behboodi et al., 1995; Wilson et al., 1995; Garry et al., 1996; Schmidt et al., 1996; Kruip and den Daas, 1997; Hill et al., 1999). The effects of in vitro embryo production on embryo development will be discussed in detail later in this review. INFLUENCE OF IN VITRO PRODUCTION ON EMBRYO DEVELOPMENT Embryo Developmental Rates Historically, the most widely used method of evaluating embryo quality has been whether an embryo has attained an appropriate stage of development by a predetermined time point (Elsden et al., 1978; Schneider et al., 1980; Shea, 1981; Lidner and Wright, 1983). In a retrospective analysis of embryo collections at a commercial bovine embryo transfer center, Lindner and Wright (1983) reported that the highest proportion of morulae and compact morulae were recovered 5 to 6 days after estrus, whereas early blastocysts and blastocysts were more prevalent on day 7. On days 8 and 9, blastocysts, expanded blastocysts and hatched blastocysts were most often recovered. Similar findings have been reported with both superovulated females (Shea,
11 1981; Wright, 1981) and with single ovulating cows (Hamilton and Laing, 1946; Linares and King, 1980). In contrast, in vitro-produced bovine embryos exhibit delayed development when compared to their in vivo-derived counterparts (Sirard and Lambert, 1985; Hytel et al., 1989; Grisart et al., 1994). Early cleavage divisions (up to the 8- to 16-cell stage) of in vitro-produced embryos have been reported to occur at a rate similar to those of in vivo- recovered embryos (Hamilton and Laing, 1946; McGaugh et al., 1974; Christenson et al., 1975; Moore, 1975; Betteridge and Flechon, 1988; Barnes and Eyestone, 1990). However, Grisart et al. (1994) reported that the appearance of 8- to 16-cell embryos, morulae and blastocysts in vitro is delayed by 30 to 40 hours when compared to embryos produced in vivo. These findings are in agreement with those of McGaugh et al. (1974), Prather and First (1988) and Funahashi et al. (1994). Van Soom et al. (1992) suggested that this delay indicates that development past the 8- to 16-cell stage is extremely sensitive to culture environment. Moreover, developmental arrest is very common at this stage if culture conditions are inadequate (Van Soom et al., 1992), lending additional support to Van Soom’s theory. In addition, production of embryos in vitro is much less efficient than in vivo embryo production, with less than 40% of immature oocytes reaching the blastocyst stage (Wright and Ellington, 1995; Lonergan et al., 2001). It has been suggested that the first embryos to cleave are more likely to (up to 70%) reach the morula to blastocyst stage by day 8 (Plante and King, 1992; Grisart et al., 1994; Dinnyes et al., 1999; Lonergan et al., 1999). Similar results have been reported in other species (buffalo – Totey et al., 1996; rhesus monkey – Bavister et al., 1983; McKiernan and Bavister, 1994; human – Sakkas et al., 1998; Shouker et al., 1998). In addition, Van Soom et al. (1992) observed that embryos that cleaved at least once before 36 hours post-insemination were more likely to form compacted morulae. Similarly, Vergos et al. (1989) reported that embryos that had developed to at least the 4-cell stage by 44 to 48 hours post- insemination were 23% more likely to form blastocysts than those that had only reached the 1- to 3- cell stage by this time. Embryo Morphology Morphological characteristics of early bovine embryos have been described by several authors (Hamilton and Laing, 1946; Chang, 1952; Shea, 1981; Wright, 1981;
12 Lidner and Wright, 1983; Betteridge and Flechon, 1988), and numerous reports have demonstrated that embryos produced under in vitro conditions display distinct morphological differences when compared with in vivo-derived embryos (Plante and King, 1994; Thompson, 1997; Holm and Callesen, 1998; Abe et al., 1999). For example, embryos produced under in vitro conditions have been found to have darker cytoplasm, a grainy appearance, less perivitelline space and swollen blastomeres when compared with in vivo-derived embryos of similar degrees of development. Other observable differences in ultrastructural features of in vitro-produced embryos include the amount of lipid droplets contained in the cytoplasm, the development of junctional complexes between trophoblast cells and/or cells of the ICM, development of microvilli on the apical surface of trophoblast cells and alterations in morphological features of the zona pellucida (Abe et al., 1999). Interestingly, Abe et al. (1999) reported that morulae and blastocysts cultured in vitro in the presence of serum contained a higher number of lipid droplets, particularly within trophoblastic cells, than in vivo-derived morulae and blastocysts. These findings are in agreement with other reports (Brackett et al., 1980b; Mohr and Trounson, 1981; Shamsuddin et al., 1993; Plante and King, 1994; Crosier et al., 2000). Moreover, Abd El Razek et al. (2000) determined that the lipids observed in embryos produced in vitro were predominately triglycerides, with less lipids from other classes. Crosier et al. (2000) observed an increase in lipids in compact morulae produced in vitro compared with in vivo-derived morulae, regardless of the type of medium in which the embryos were cultured [TCM-199 with 10% estrous cow serum (ECS), TCM-199 with BSA or TCM-199 with BSA and ECS from 72 hours post-insemination (hpi) to 144 hpi or mSOF)]. Taken together, these findings suggest that the accumulation of lipid droplets within the cytoplasm is characteristic of in vitro-produced embryos (Abe et al., 1999). The reason for this lipid accumulation remains unknown, but it has been suggested that it is a result of membrane breakdown in response to an artificial culture environment (Crosier et al., 2000). Alternatively, it has been suggested that insufficient mitochondrial metabolism may cause the elevated lipid levels noted in in vitro-produced embryos (Dorland et al., 1994). Mitochondria, specifically the cristae, are the primary site of lipid metabolism. Immature mitochondria, lacking sufficient cristae, have been found in early cleavage and
13 morula stage bovine and primate embryos (Enders and Schlafke, 1981; Betteridge and Flechon, 1988). These embryos may have a decreased ability to metabolize lipids into ATP, thus causing a buildup of lipids within the embryo (Crosier et al., 2000). Pregnancy and Calving Rates In addition to producing lower quality embryos, in vitro maturation and fertilization of oocytes and/or in vitro culture of embryos results in other developmental problems. Generally, a bovine embryo will develop for ~12 to 14 days in vitro, but will fail to elongate as seen in vivo at this time. Therefore, in order to produce live offspring from in vitro-produced embryos, the embryos must be transferred to recipient females. Unfortunately, pregnancy rates achieved from the transfer of in vitro-produced embryos are lower than those achieved from in vivo-derived embryos. One of the first studies illustrating this difference was reported by Hasler et al. (1995). In this study 1884 fresh in vitro-produced embryos were transferred to recipient females resulting in a 56% pregnancy rate. In comparison, 320 in vivo-derived embryos were transferred, resulting in a 66% pregnancy rate. The pattern of decreased pregnancy rates to embryo transfer (ET) was consistent across embryo quality grades (grade 1, grade 2 and grade 3 embryos) and day of transfer (day 7 or day 8, post onset of estrus) when pregnancy rates for in vitro-produced embryos were compared with pregnancy rates for in vivo- derived embryos. However, as in vitro embryo culture systems have improved over time so have pregnancy rates resulting from the transfer of in vitro-produced embryos. Farin et al. (2001) reported no difference between pregnancy rates from the transfer of in vitro- produced (63%) and in vivo-derived embryos (65%). However, when serum-restricted in vitro-produced embryos were transferred, pregnancy rates were lower than those achieved from the transfer of serum supplemented in vitro-produced or in vivo-derived embryos. There was also a higher incidence of degenerate conceptuses resulting from the transfer of in vitro-produced embryos compared with their in vivo-derived counterparts. Taken together, these reports indicate that improvements to in vitro culture conditions have improved pregnancy rates achieved from the transfer of in vitro- produced embryos; however, many critical problems remain. This is predominantly a two part problem consisting of: (1) a higher incidence of abortion and (2) a higher incidence of post-natal loss. Hasler et al. (1995) reported that abortions occurring from the transfer
14 of in vitro-produced embryos were not randomly distributed. In a two year study, abortion occurred in 11% (range = 8% to 16% ) of 542 pregnancies resulting from the transfer of in vitro-produced embryos. In comparison, an abortion rate of only 5% has been reported for in vivo-derived ET pregnancies (Hasler et al., 1987). The timing of these abortions in regard to stage of gestation is also different from that of normally mated cattle. Ayalon (1978) and Diskin and Sreenan (1980) reported that the majority of pregnancy loss occurs during the embryonic period of gestation (day 1 to day 42). However, in pregnancies resulting from the transfer of in vitro-produced embryos, an abortion rate of 13 to 20% has been reported to occur during the last 2 trimesters of gestation (Hasler et al., 1995; Hasler et al., 2000; Agca et al., 1998). It has been suggested that an increase in placental abnormalities is a contributing factor in the increased incidence of abortion observed among ET pregnancies resulting from in vitro-produced embryos. Farin et al. (2000) reported that placental weight of fetuses resulting from in vitro-produced embryos were significantly heavier than those from fetuses resulting from in vivo-derived embryos. Although there were no differences in the number of placentomes and placental fluid volume, placentas of in vitro-produced embryos had a significantly smaller caruncular surface area compared with placentas from in vivo-derived embryos. Furthermore the fetal binucleate cell volume was significantly less for placentas resulting from in vitro-produced embryos compared with in vivo-derived pregnancies. In addition, Hasler et al. (1995) reported that hydrallantois occurred in 1 of every 200 pregnancies. This was significantly higher than the rate reported for normal pregnancies (1 of every 7500 pregnancies). Similar findings have been reported by Hill et al. (1999) and Mello et al. (2003). In addition to problems associated with placental malformations and increased abortion rates, in vitro-produced ovine and bovine fetuses that survive to term often experience significantly longer gestation periods (Walker et al., 1992; Sinclair et al., 1995; Kruip and den Daas, 1997). The mean increase in gestation length for cattle has been reported to be 3.6 days longer for in vitro-produced embryos compared with gestation lengths from normal mating, artificial insemination (AI) or ET (Sinclair et al., 1995; Kruip and den Daas, 1997). Because of the increased incidence of abortion, the calving rate for cows transplanted with in vitro-produced embryos is significantly lower than for cows receiving
15 in vivo-derived embryos (Behboodi et al., 1995; Hasler et al., 1995). Furthermore, of the cows that calve, there is an increase in dystocia in pregnancies achieved from in vitro- produced embryos (Behboodi et al., 1995; Schmidt et al., 1996). Increases in dystocia for in vitro-produced embryo established pregnancies have been reported to range from 20% (Kruip and den Daas, 1997) to 62% (Behboodi et al., 1995). This increase is a six- fold increase over that observed with natural mating, AI or ET. In addition to lower calving rates a significant increase in perinatal morality has been reported for sheep and cattle (Walker et al., 1992; Behboodi et al., 1995; Hasler et al., 1995). Kruip and den Daas (1997) reported a two-fold increase in perinatal death (5.6% to 6.7% occurrence) from in vitro-produced embryos compared with in vivo- derived embryos. However, one of the original reports showed a 50% calf mortality following parturition in calves resulting from in vitro-produced embryos (Behboodi et al., 1995). In respect to the perinatal death rate increase there has been a report of an increase in congenital abnormalities as a result of the transplantation of in vitro-produced embryos (Schmidt et al., 1996). Large Offspring Syndrome and Associated Disorders One of the first reported incidences of ‘large offspring syndrome’ was described by Walker et al. (1992) as possibly resulting from transplantation of in vitro-produced ovine embryos. Lambs resulting from these in vitro-produced embryos were significantly heavier at birth (mean = 5.6±0.31 kg) compared with lambs produced from in vivo- derived embryos (mean = 4.8±0.27 kg). Since this time there have been many descriptions of large offspring syndrome occurring in sheep and cattle after transfer of in vitro-produced embryos (Behboodi et al., 1995; Hasler et al., 1995; Schmidt et al., 1996). In cattle, birth weights from in vitro-produced embryos have been reported to deviate from that of naturally produced calves by as little as 1.9 kg to as much as 15.9 kg (Kruip and den Daas, 1997). These adverse affects were attributed to factors within the culture environment. Since this time there have been many articles describing and attempting to explain the causes of large offspring syndrome. It has been hypothesized that in vitro culture is an abnormal environment and may be a leading cause of this problem. Others have attributed the increased birth weight to the use of co-culture systems, and still
16 others believe that the addition of bovine serum albumin (BSA) or serum to the culture medium is the primary cause. In the absence of in vitro culture there are two reported methods to increase embryo and/or conceptus size in utero. One of these methods is to transfer an embryo into a more advanced stage uterus. Wilmut and Sales (1981) demonstrated this by transferring early stage sheep embryos (4- to 8-cell and blastocyst stage) into uteri that were -72 to +72 hours in synchrony with the transferred embryos. They concluded that transfer of embryos into a uterus that is developmentally behind that of the embryo resulted in rapid degeneration of the embryo. However, when an embryo was transferred into a uterus that is developmentally advanced compared with the embryo the embryo experienced a rapid increase in development, so that it matched the stage of the uterus within a few days. A second method to increase the rate of development and size of the fetus is progesterone (P4) treatment to early pregnant females. Kleemann et al. (1994) reported that if a pregnant ewe was administered P4 during the 3 days following ovulation fetal crown-rump size and weight would be significantly larger compared with nontreated controls. Later, Kleemann et al. (2001) found that the increase in fetal length and weight were a response to increased heart, skeletal and muscle growth of the conceptus. These findings are similar to reported increases in fetal size following in vitro culture. Farin and Farin (1995) reported that following the transfer of in vitro-produced embryos the resulting fetuses had significant increases in body weight, heart girth, long-bone lengths and heart weight compared with fetuses produced from the transfer of in vivo- derived embryos. There are conflicting reports regarding the incidence of large offspring syndrome following culture of in vitro-produced embryos with co-culture cells. The first incidence of large offspring syndrome was reported to occur in sheep as a result of in vitro culture; this was coincidentally one of the first reports of successful in vitro production of embryos in the absence of co-culture (Walker et al., 1992). But one of the first reports of large offspring syndrome occurring in in vitro-produced embryo pregnancies used an oviduct epithelium co-culture system (Behboodi et al., 1995). Even more puzzling are the reports of a very low incidence of large offspring syndrome as a result of the transplantation of in vitro-produced embryos (Massip et al., 1996; Breukelman et al.,
17 2002). However, these two reports have two things in common, low numbers and low pregnancy rates that may be the cause of these contradictory results. When ovine pregnancies were established from in vitro-produced embryos cultured in SOF supplemented with human serum or BSA there was a significant reduction in birth weight from embryos cultured in SOF supplemented with BSA compared with human serum (Thompson et al., 1995). Furthermore, when in vivo- derived embryos were cultured in SOF supplemented with BSA the lambing weights were not different from the in vitro-produced embryos cultured in the same medium (Thompson et al., 1995). This finding demonstrated early that regardless of the origin of the embryo, in vitro culture could result in large offspring syndrome at parturition, but the addition of serum significantly magnified this effect. Later, however, Thompson et al. (1998) reported that calves resulting from embryos cultured in SOF supplemented with FCS where lighter at birth compared with calves produced from embryos cultured in SOF supplemented with BSA or charcoal striped FCS, although the differences were not significant. However, culturing embryos in the presence of serum still results in significantly higher birthweights in sheep (Sinclair et al., 1998; Brown and Radziewic, 1996) and in cattle (Farin and Farin, 1995; Kruip and den Daas, 1997) compared with naturally mated, AI or MOET produced control calves. In addition, pregnancies resulting from the transfer of embryos cultured in the presence of serum or co-culture systems resulted in significantly greater fetal liver and heart sizes compared with those of fetuses derived from embryos cultured in vitro without serum or co-culture (Sinclair et al. 1997). Apoptosis There are two distinct forms of cell death, necrosis and apoptosis, which can be distinguished by specific morphological criteria (Wyllie et al., 1980; Majno and Joris, 1995). Necrosis is generally considered to be caused by accidental injury to the cell initiated by environmental insults such as severe hypoxia, ischemia, reactive oxygen species, heat shock or exposure to metabolic toxins or infections (Wyllie et al., 1980; Betts and King, 2001). It affects large groups of cells or tissues, causing cellular swelling and membrane rupture (Wyllie et al., 1980). Necrosis will not be discussed in detail in this dissertation.
18 In contrast, apoptosis is a self-directed form of cell suicide that characteristically affects single cells (Hardy, 1997; Betts and King, 2001). Initially, the cell shrinks, and the chromatin condenses and migrates against the inner nuclear membrane, causing gross indentions in the membrane (Hardy, 1997; Betts and King, 2001). The nucleus fragments, and the DNA is cleaved between nucleosomes forming oligonucleosomal- sized fragments (~185 bp; Hardy, 1997; Betts and King, 2001). During the final phases of apoptosis, the cytoplasm condenses and the nucleus breaks up (karyohexis) into pyknotic nuclear fragments (Betts and King, 2001). The fragmenting cellular components can be found in budding membrane processes (called blebs) that break off into apoptotic bodies, which are then either extruded into intracellular tissue spaces or phagocytized by macrophages or neighboring tissue cells (Wyllie et al., 1980; Majno and Joris, 1995). Cell death during early embryo development Characteristic events of apoptosis have been documented in blastocysts of several species including mice (El-Shershaby and Hinchliffe, 1974; Copp, 1978; Handyside and Hunter, 1986), cattle (Mohr and Trouson, 1982; Plante and King; 1994), baboons (Enders et al., 1990), rhesus monkeys (Hurst et al., 1978; Enders and Schlafke, 1981; Enders et al., 1982) and humans (Mohr and Trouson, 1982; Hardy, 1999; Hardy et al., 1989; Hardy et al., 1996; Sathananthan et al., 1982). There is little evidence, however, that apoptosis occurs prior to compaction in developing embryos of good quality (Hardy et al., 1989). For example, nuclear DNA fragmentation has been observed in situ using the TdT-mediated dUTP nick-end labeling (TUNEL) method in in vitro-produced bovine oocytes, 8-cell embryos, morulae and blastocysts, but not in zygotes or early cleavage-stage embryos (Byrne et al., 1999; Matwee et al., 2000), nor in in vivo-derived embryos recovered from superovulated cows (referenced in Betts and King, 2001). Similar results have been observed in pigs in which DNA fragmentation was detected in in vitro-produced oocytes (Tatemoto et al., 2000) and blastocysts (Long et al., 1998), but not 4- to 8-cell embryos. Normal developing human embryos have also failed to show signs of apoptosis from the 2-cell to uncompacted morula stage (Jurisocova et al., 1996). It must be kept in mind, however, that it is difficult to estimate the full extent of apoptosis in the embryo, as current techniques provide only a ‘snapshot’ of the embryo at a specific point in time (Hardy, 1997). The initial stages of apoptosis appear to occur
19 very rapidly, with cytoplasmic fragmentation occurring within minutes (Wyllie et al., 1980). In addition, phagocytosis of cell remnants can occur within as little as 12 to 18 hours (Wyllie et al., 1980). These findings suggest that observation of only a moderate number of dead cells by current techniques may in all actuality represent extensive cell death (Hardy, 1997). Apoptosis is a critical process during pre-implantation development for the disposal of excess cells, cells that are in the way, abnormal or potentially dangerous (Gjorret et al., 2003). Moreover, elevated levels of apoptotic cells have been correlated with abnormal embryo morphology (Hardy et al., 1989) and it has been suggested that when apoptosis surpasses a certain ‘threshold level’ further embryonic development is compromised (Tam, 1988; Juriscova et al., 1998; Hardy, 1999) and nonviable offspring are eliminated (Byrne et al., 1999). Apoptosis appears to be a normal part of embryonic development as in vivo-derived embryos from mice (El-Shershaby and Hinchliffe, 1974; Copp, 1978; Handyside and Hunter, 1984; Brison and Schultz, 1997; Devreker and Hardy, 1997), rats (Pampfer et al., 1990a,b), baboons (Enders et al., 1990), rhesus monkeys (Enders and Schlafke, 1981) and humans (Hertig et al., 1954) have exhibited characteristics of apoptosis immediately upon recovery from the reproductive tract. Apoptosis appears to predominate in the ICM (El-Shershaby and Hinchliffe, 1974; Copp, 1978; Mohr and Trouson, 1982; Handyside and Hunter, 1986; Hardy and Handyside, 1996), with much lower levels detected in the trophoectoderm (Copp, 1978; Hardy and Handyside, 1996; Brison and Schultz, 1997) and it has been suggested that apoptosis occurs naturally in vivo as a means to help eliminate ICM cells that still have the potential to form trophoectoderm, thus decreasing the chance of inappropriate trophoectoderm expression during germ layer differentiation (Handyside and Hunter, 1986; Pierce et al., 1989). In support of this theory, a wave of cell death is apparent in mouse embryos recovered from the reproductive tract 96 hours post-hCG administration coincident with the time that ICM cells loose the ability to differentiate into trophoectoderm cells (Handyside and Hunter, 1986). In contrast, in vivo-derived rabbit (Giles and Foote, 1995) and porcine (Papaioannou and Ebert, 1988) embryos show no evidence of apoptotic activity upon recovery from the reproductive tract. Although, the technique (differential labeling) used in these reports may not be adequate to detect apoptotic activity in these species.
20 Apoptosis may also function to remove abnormal cells from the embryo (Hardy, 1997; Hardy, 1999). Nuclear and chromosomal abnormalities have been identified in both human and bovine embryos. Specifically, ~ 20% of human embryos exhibit gross chromosomal abnormalities and more than 40% of embryonic cells are mosaic (i.e., some cells within the same embryo are diploid while others are aneuploid, haploid or triploid; Harper et al., 1995). Similarly, 25% of in vivo- and 72% of in vitro-produced bovine embryos were identified as mosaic (Viuff et al., 1999). These data are in agreement with those of Jacobs (1990) and King (1991) in that the in vitro production environment appears to increase nuclear and chromosomal abnormalities and consequently reduce their developmental potential. Effect of in vitro culture in the incidence of apoptosis In vitro embryo production has been correlated with alterations of normal apoptotic patterns; however, conflicting data does exist. In one study, the incidence of apoptotic cells in murine blastocysts was similar to those seen in in vivo-derived embryos (Devreker and Hardy, 1997). In a separate study, the percentage of apoptotic cells was significantly higher in murine embryos produced in vitro as compared to their in vivo-derived counterparts (Brison and Schultz, 1997). Similarly, in vitro-produced bovine blastocysts had a significantly higher number of apoptotic cells than those that developed completely in vivo (Byrne et al., 1999; Gjorret et al., 2001). It is generally accepted that maternal- and embryonic-derived growth factors significantly influence pre-implantation development (Kaye, 1997; Watson et al., 2000) and it has been suggested that the effect of in vitro culture may be a result of the presence or absence of these factors in the culture medium (Brison and Schultz, 1997; Hardy, 1997; Moley et al., 1998; Gjorret et al., 2001). Both transforming growth factor α (TGF-α) and insulin-like growth factor I (IGF-I) increase blastocyst formation and blastocyst cell number, presumably by reducing blastomere cell death (Brison and Schultz, 1997; Lighten et al., 1998). Similarly, the incidence of apoptosis is elevated in blastocysts from diabetic rats (Pampfer et al., 1990a), but can be reduced by maternal treatment with insulin (de Hertogh et al., 1992). In contrast, both in vitro- and in vivo- derived murine blastocysts had a significant increase in nuclear fragmentation in ICM cells with a concomitant reduction in the number of ICM cells and embryo viability after treatment with tumor necrosis factor-α (TNF-α; Wuu et al., 1999). Byrne et al. (1999)
21 suggested that the addition of TBS to embryo culture media, while beneficial, may also be detrimental in that it may contain ‘death’ factors such as TNF-α that could trigger apoptosis within the embryo. Regulation of apoptosis and expression of apoptotic genes during early embryo development
The Bcl-2 family of genes are key regulators of apoptosis in mammalian cells (Betts and King, 2001). The Bcl-2 family of proteins can be subdivided into two distinct groups: (1) the anti-apoptotic proteins or survival proteins, including Bcl-2, Bcl-XL and
Bcl-W and (2) the pro-apoptotic proteins or death proteins, including Bax, Bcl-XS, Bak, Bas, Bik and Bid (Krajewski et al., 1993). It is believed that the ratio of pro- and anti- apoptotic proteins, rather than the expression levels of individual Bcl-2 genes, determine a cell’s susceptibility to apoptosis (Rheostat hypothesis; Oltvai et al., 1993). There are several mechanisms through which apoptosis can be induced in cells; however, there are thought to be two general pathways: the receptor or extrinsic pathway and the mitochondrial or intrinsic pathway (Lawen, 2003). In the receptor mediated or extrinsic pathway, extracellular ligands bind to specific death receptors bound to the plasma membrane of the targeted cell. The most common death receptors are Fas (also called Apo-1 or CD95) and tumor-necrosis factor receptor-1 (TNF-R1; Lawen, 2003). Activation of a death receptor results in activation of an initiator caspase, caspase-8 or -10 in particular. The initiator caspase then activates other caspases in the cascade eventually leading to the activation of effector caspases, such as caspase-3 or - 6 (Lawen, 2003). In contrast to the receptor-mediated induction pathway, the mitochondrial or intrinsic pathway is activated by intracellular stresses, such as withdrawal from growth factors (Betts and King, 2001), oxidative stress (Lawen, 2003) or exposure to DNA damaging agents (Betts and King, 2001; Lawen, 2003). In this pathway, pro-apoptotic members of the Bcl-2 family, usually Bax, undergo a conformational change that allows them to integrate themselves into the outer mitochondrial membrane. Once incorporated into the membrane they interact with voltage-dependent anion channels, causing the formation of pores in the membrane and allowing release of cytochrome c into the cytosol (Shimizu et al., 1999). Within the cytosol, cytochrome c binds to Apoptotic Protease Factor-I (APF-I) to form an apoptosome. The apoptosome contains the initiator
22 caspase procaspase-9 (Waterhouse et al., 2002) that, when activated, initiates a cascade of events leading to activation of an effector caspase and cellular breakdown (Lawen, 2003). It has been suggested the fate of the cell via the intrinsic pathway is influenced by the ratio of death factors to survival factors (Wyllie, 1995). Specifically, survival factors, such as Bax and Bcl-XL, may act to prevent the release of cytochrome c from the mitochondria by binding to Bax or by closing the pores in the mitochondrial membrane (Shimizu et al., 1999). As indicated, regardless of the type of induction, both signaling pathways converge at the level of the caspases (Lawen, 2003), a family of cysteine-dependent aspartate-specific proteases that cleave key cellular proteins and cause disassembly of the cell (Barinaga, 1998; Mehmet, 2000). To date, 14 mammalian caspases have been identified (Strasser et al., 2000) and divided into three groups based on function (Grutter, 2000): (1) the initiator caspses (caspase-2, -8, -9 and -10) recognize specific receptors in the cell and ‘declare the death sentence’ (Betts and King, 2001), (2) the executioner or effector caspases (caspase-3, -6 and -7) which ‘carry out the death sentence’ by breaking down cellular proteins (Betts and King, 2001) and (3) the caspases involved in cytokine maturation rather than apoptosis (caspase-1, -4, -5, -11, -12, -13 and -14). Briefly, initiator caspases cleave and activate effector caspases, which then cleave cellular proteins causing the typical morphological changes observed in cells undergoing apoptosis (i.e., membrane blebbing, disassembly of the cell structure and DNA fragmentation; Lawen, 2003). Transcripts for apoptosis related genes have been detected in early stage embryos of mice (Exley et al., 1999), cattle (Kolle et al., 2002; Yang and Rajamahendran, 2002; Rizos et al., 2003) and humans (Warner et al., 1998). Specifically, transcripts for both Bax and Bcl-2 have been identified in 2- to 12-cell human embryos (Warner et al., 1998). Moreover, levels of Bcl-2 were higher in blastocysts of good morphology; whereas, Bax transcript levels were higher in blastocysts with extensive fragmentation (Hardy, 1999). Similar results have been observed in mice in that fragmented blastocysts contained lower levels of Bcl-2 (Exley et al., 1999) adding support to the hypothesis that alteration in the ratio of Bcl-2 to Bax determines susceptibility to apoptosis (Oltvai et al., 1993). In cattle, Bcl-2 expression has been detected from the 4-cell stage throughout development and in both the TE and ICM
23 of blastocysts and hatched blastocysts (Kolle et al., 2002; Neuber et al., 2002), with apoptosis being proportionally higher in the ICM than the TE (Neuber et al., 2002). Similar to results reported in mice, Bcl-2 expression varied between embryos and was suggested to be related to degree of fragmentation of the embryo (Kolle et al., 2002). Similarly, Bax expression has been detected from the 8-cell stage throughout development (Byrne et al., 1999; Matwee et al., 2000; Paula-Lopes and Hansen, 2002; Lonergan et al., 2003). In addition, Yang and Rajamahendran (2002) found that Grade I bovine embryos had a much higher expression of Bcl-2 compared with Grade IV embryos; whereas Bax expression was much more prominent in Grade IV embryos. These authors suggest that Grade IV embryos are in an advanced stage of apoptosis, as indicated by the elevated levels of Bax expression (Yang and Rajamanhendran, 2002). Furthermore, these authors concur with the concept proposed by Oltvai et al. (1993) that the ratio of Bcl-2 to Bax determined cell fate (Yang and Rajamanhendran, 1993). Similar results have been observed with bovine embryos produced in the presence of serum (Rizos et al., 2003). CHICK EMBRYO CO-CULTURE Biochemistry of the Avian Embryo The avian egg is a complete environment capable of sustaining embryonic development from the earliest stages of blastulation through hatching. At the time of oviposition, the chicken egg is composed of ~ 56% albumen, 32% yolk and 12% shell (Romanoff and Romanoff, 1949). At this time, the albumen contains ~ 88% water, 10% protein, 0.05% lipid and 0.8% ash (Runge, 1982). In contrast, the yolk is composed of ~ 49% water, 17% protein, 32% lipid and 2% ash (Runge, 1982). The yolk is the main source of nutrients for the developing chick during early development. The nutrients within the yolk are quite concentrated and because fat is much more concentrated calorically than sugar, the egg is rich in fat rather than carbohydrates (Romanoff, 1967). High energy lipids make up almost 64% of the dry weight of the egg, including lecithin, cephalin, cholesterol, carotenoids and ergosterol (Romanoff, 1967). In addition, egg yolk contains glucose, glycogen, lactic acid and high energy adenosine triphosphate, as well as many amino acids, vitamin A, ascorbic acid, vitamin B12, thiamine and minerals such as chloride, calcium, sodium, potassium and iron (Romanoff, 1967).
24 During early embryo development, the embryo receives nutrients from the yolk via the vitelline circulation (Runge, 1982). It is believed that the amniotic fluid that bathes the developing embryo is a result of the metabolism of these nutrients (Runge, 1982). However, the regulation of the volume and contents of the amniotic fluid are poorly understood (Pierce, 1933; Romanoff, 1960). Recent reports indicate that there are three barriers (the blood/amnion barrier, the blood/allantois barrier and the allantois/amnion barrier) that specifically control amino acid concentration within these compartments (ten Busch et al., 1997b). In birds, the allantois and amnion lack innervation (Pierce, 1933; Romanoff, 1960), so it is believed that the amniotic and allantoic fluids are controlled largely by hormones (Schmidek et al., 2001). The components of avian amniotic fluid still remain largely undefined; however, ions of chloride, sodium, potassium, phosphorus, magnesium, calcium, iron and sulfur are present as well as low levels of proteins and carbohydrates (Smoczkiewiczowa, 1959). In addition, 47 free amino acids and related compounds have been identified in the plasma, amniotic fluid and/or allantoic fluid of the chick embryo (Epple et al., 1992; Gill et al., 1994; ten Busch et al., 1997a,b; Epple et al., 1997; Piechotta et al., 1998) and IGF-I has also been reported to be widespread in the tissues of the chick (McFarland et al., 1992; De Pablo et al., 1993; Yang et al., 1993; Tanaka et al., 1996; McMurtry et al., 1997). Early Use of Chick Embryo Extracts in Mammalian Cell Culture Extracts obtained from developing chick embryos have been used throughout the history of mammalian cell culture. In 1913, Alexis Carrel reported that when extracts from 20-day old chick embryos were added to in vitro cultures of canine connective tissue, there was a dramatic increase in growth rate. In a second experiment he noted that a 30-fold increase in growth rate could be obtained if the chick embryos were stored for 20 days in cold storage prior to extraction (Carrel, 1913). Later, Willmer and Jacoby (1936) reported that extracts from day 7 chick embryos could stimulate the development of quiescent avian cells. Interestingly, the rate of cell proliferation was proportional to the amount of extract in the medium. Since these early experiments, chick embryo extract has been used as a supplement for many different cell types including HeLa cells (Takano, 1957), heart endothelial cells (Namikawa, 1969), retinal pigmented epithelium cells (Chader et al.,
25 1975), rat pancrease cells (McEvoy and Hegre, 1976), chick epidermis (Matsuhashi and Sugihara, 1984), dorsal root ganglion (Xue et al., 1992) and hematopoietic cells (Nicolas-Bolnet et al., 1995) and it is commonly used for modern neural crest (Hou and Takeuchi, 1992; Wu et al., 2001) and muscle (Slater, 1976; Scarpini et al., 1982; Ii et al., 1982; Bischoff, 1986; Smith and Schroedl, 1992; Nicolas-Bolnet and Dieterlen-Lievre, 1995) cell cultures. In Vitro Culture of Chick Embryos The earliest report of the culture of whole egg contents outside of the egg shell were those of Preyer (1885; referenced in Rowlett, 1990). With this system, embryo survival was limited to only 2 days. Since that time, many researchers have attempted to improve embryo viability following shell-less chick culture. The most successful method has been that of Dunn (1974) in which he suspended the egg contents in commercial plastic wrap placed in culture vessels made of modified Playtex® nurser holders. More importantly, he determined that incubation in a 5% CO2 humidified environment improved embryo viability with chicks surviving up to 22 to 23 days, but were unable to hatch. This technique has since been modified (Dunn and Boone, 1976, 1977, 1978; Dunn et al., 1981; Dunn et al., 1987, Blakewood and Godke, 1989). The advantages to using this method of chick culture is that development of the chick embryo is easily observed and reasonable survival rates can be obtained. More important to this dissertation, this culture system allows manipulations of the chick embryo and access to the amnion for injection of mammalian embryos. The main drawback to culturing chick embryos in the absence of a shell is that the shell provides 70 to 80% of the calcium required by the developing chick (Johnston and Comar, 1955; Romanoff, 1960, Crooks and Simnkiss, 1974). This is a problem in that it appears to prevent the chick from “hatching” from this shell-less culture system. In addition, because the embryo is maintained on plastic film, the egg contents assume abnormal spatial relationships due to the shape of the plastic film sac and excessive evaporation may occur from the surface of the cultures. All of these factors have significant effects on the growth and development of chick embryos in shell-less culture (Rowlett, 1990). Chick Embryo Co-Culture The use of avian eggs for the culture of mammalian embryos was envisioned by Dr. R.A. Godke in 1975 (personal communication). However, it took more than 15 years
26 for this idea to come to fruition. Early attempts using unfertilized hen’s eggs proved largely unsuccessful (Blakewood et al., unpublished), but after much trial and error a technique using shell-less chick embryos for the co-culture of mammalian embryos was developed (Blakewood et al., 1989a,b). With this technique, mammalian embryos were embedded in agar and injected into the amnion of a live chick embryo maintained in shell-less culture. After 72 to 96 hours of culture, the mammalian embryos could be recovered from the amnion and transferred to recipient females. In the initial report, pronuclear stage murine embryos from two separate strains, one of which characteristically showed a 2-cell developmental block in vitro, were placed in the amniotic cavity of 96-hour chick embryos and cultured for 72 hours. Upon recovery, there were significantly more hatched blastocysts resulting from culture in the chick embryo in both strains of mice (Blakewood et al., 1989a) when compared with Whitten’s control medium. Later studies were conducted to determine if the chick embryo amnion could support development of caprine embryos. Blakewood et al. (1990) reported that culture of 2- to 8-cell caprine embryos in the chick amnion allowed development past the in vitro developmental block. Furthermore, these embryos developed to blastocysts at rates equal to co-culture of similar stage embryos on uterine fibroblasts. When culture was extended to 96 hours, significantly more embryos developed to the blastocyst stage after culture in the chick amnion. Furthermore, 12 of the resulting embryos were surgically transferred to recipient females and resulted in 6 live offspring. Further studies indicated that the chick amnion could promote development of pre-compaction stage bovine morulae to the expanded blastocyst stage at rates significantly greater than embryos cultured on uterine fibroblasts or in medium alone. In addition, chick embryo co-culture prior to freezing improved post-thaw viability when compared with embryos cultured in medium alone prior to freezing.
27 CHAPTER III
CULTURING IN VITRO-PRODUCED BOVINE EMBRYOS IN THE AMNIOTIC CAVITY OF THE DEVELOPING CHICKEN EMBRYO
INTRODUCTION With the advent of in vitro fertilization technologies in the 1980s came the need for methods of culturing embryos to later stages of development. Early methods included in vitro co-culture with feeder cells (Cole and Paul, 1965), fibroblast monolayers (Kuzan and Wright, 1982; Voelkel et al., 1985) or trophoblastic vesicles (Camous et al., 1984; Heyman et al., 1987; Pool et al., 1988) and in vivo culture in the ligated oviducts of sheep (Willadsen, 1979). As early as 1975, Dr. R.A. Godke envisioned the use of avian eggs for the culture of mammalian embryos (personal communication). Early attempts using unfertilized hen’s eggs proved largely unsuccessful (Blakewood et al., unpublished data), but after several months of trial and error a technique was developed that used shell-less chick embryos for the co-culture of mammalian embryos. Chick embryo co- culture improved development of murine (Blakewood et al., 1989a), caprine (Blakewood et al., 1989b) and bovine (Blakewood et al., 1989b) embryos compared with the culture media of the time, with embryos consistently passing thru the in vitro developmental block and a greater percentage reaching the blastocyst stage compared to control methods. Since that time, in vitro embryo culture techniques have progressed and there is no longer a need for in vitro co-culture or culture in the oviducts of sheep. However, many problems still remain. With current embryo culture methods, only 30 to 40% of in vitro-matured and fertilized ova routinely reach the blastocyst stage (Krisher and Bavister, 1998; Lonergan et al., 2003). In addition, in vitro-produced embryos show marked differences in morphology (Greve et al., 1995), timing of development (Van Soom et al., 1997), embryonic metabolism (Khurana and Niemann, 2000) and pregnancy rates following transfer (Enright et al., 2000) when compared with embryos derived in vivo. It has been hypothesized that exposure of embryos to in vitro culture conditions, particularly exposure to fetal bovine serum (FBS), may result in abnormal fetal and neonatal development and growth, known collectively as large calf or large offspring
28 syndrome (reviewed by Kane, 2003). Specifically, offspring produced from embryos cultured in vitro have exhibited increased birth weights (Willadsen et al., 1991; Keefer et al, 1994; Behboodi et al., 1995), longer gestation lengths (Walker et al., 1996), increased placental anomalies (Hasler, et al., 1995; Hill et al., 1999, 2001, 2002) and increased incidences of abortion (Walker et al., 1996; Hill et al., 1999) and perinatal (Behboodi et al., 1995; Wilson et al., 1995; Garry et al., 1996; Schmidt et al., 1996; Kruip and den Daas, 1997; Hill et al., 1999) and neonatal mortality (Schmidt et al., 1996; Kruip and den Daas, 1997; Hill et al., 1999). Breathing difficulties (Young et al., 1998) and disruptions in the growth and development of body organs are also common (Kane, 2003). The problems associated with modern in vitro embryo culture systems have led us to re-visit the chick embryo co-culture system. In the present study, the development of bovine 1-cell and 2-cell embryos produced by in vitro fertilization was assessed after culture in the chick embryo amnion. This study was conducted as a preliminary evaluation of the use of this novel culture technique as a means to improve in vitro development of bovine embryos. MATERIALS AND METHODS Experimental Design Experiment 3.1 The design for Experiment 3.1 is illustrated in Figure 3.1. This experiment was conducted to determine if IVF-derived 1-cell embryos could develop to morulae and blastocysts at rates similar to IVF-derived embryos cultured in CR1aa. Upon removal from in vitro insemination, 1-cell embryos were randomly allotted to one of four treatment groups for in vitro culture. Bovine 1-cell embryos allotted to Treatment A (control) were cultured in CR1aa from day 0 to day 3 post-insemination and CR1aa containing 5% FBS from day 3 to day 7. Bovine 1-cell embryos allotted to Treatment B (agarose control) were embedded in agarose immediately upon removal from insemination and cultured in CR1aa from day 0 to day 3 post-insemination and CR1aa containing 5% FBS from day 3 to day 7. Bovine 1-cell embryos allotted to Treatment C (CEC-I) were embedded in agarose immediately upon removal from insemination and then injected into the amniotic cavity of 96-hour chick embryos. After 72 hours of co-culture, the bovine embryos were recovered from the amnion and placed in CR1aa containing 5% FBS for the remaining
29
Control Groups Media Change IVF IVC Addition of FBS Evaluation
5 h
48 h ~24 h 72 h 96 h
Onset of Preparation of Injection of Recovery of CEC-I Recovery of CEC-II Egg Incubation Shell-less CEC-I Injection of CEC-II Evaluation Cultures
Chick Culture Treatment Groups
Figure 3.1. Experimental design for the culture of 1-cell bovine embryos in the amnion of a developing chick embryo.
30 culture period. Bovine 1-cell embryos allotted to Treatment D (CEC-II) were embedded in agarose immediately upon removal from insemination and cultured in CR1aa from day 0 to day 3 post-insemination. On day 3, agar-embedded embryos were injected into the amniotic cavity of 96-hour chick embryos. After 96 hours of culture, the bovine embryos were recovered from the amnion and evaluated. Experiment 3.2 The design for Experiment 3.2 is illustrated in Figure 3.2. This experiment was conducted to determine if IVF-derived 2-cell embryos could develop to morulae and blastocysts at rates similar to IVF-derived embryos cultured in CR1aa. Presumptive zygotes were purchased from a commercial source and upon arrival were randomly allotted to one of four treatment groups for in vitro culture. Bovine 2-cell embryos allotted to Treatment A (control) were cultured in CR1aa from day 0 to day 3 post-insemination and CR1aa containing 5% FBS from day 3 to day 7. Bovine 2-cell embryos allotted to Treatment B (agar control) were embedded in agarose immediately upon arrival and cultured in CR1aa from day 0 to day 3 post-insemination and CR1aa containing 5% FBS from day 3 to day 7. Bovine 2-cell embryos allotted to Treatment C (CEC-I) were embedded in agarose immediately upon arrival and then injected into the amniotic cavity of 96-hour chick embryos. After 72 hours of co-culture, the bovine embryos were recovered from the amnion and placed in CR1aa containing 5% FBS for the remaining culture period. Bovine 2-cell embryos allotted to Treatment D (CEC-II) were embedded in agarose immediately upon arrival and cultured in CR1aa from day 0 to day 3 post- insemination. On day 3, agar-embedded embryos were injected into the amniotic cavity of 96-hour chick embryos. After 96 hours of culture, the bovine embryos were recovered from the amnion and evaluated. Experimental Methods Production of In Vitro Embryos Media Preparation Brackett-Oliphant (BO) media were prepared as previously described by Brackett and Oliphant (1975). Briefly, BO-A stock solutions contain 150.2 mM NaCL (S-5886,
MO), 3.0 mM CaCl2·2H2O (C-7902, Sigma-Aldrich Inc., St. Louis, MO), 1.0 mM
NaH2PO4·H2O (S-9638, Sigma-Aldrich Inc., St. Louis, MO), 0.6857 mM MgCl2·6H2O
31
Control Groups Shipment of Presumptive Media Change IVF Zygotes IVC Addition of FBS Evaluation
18 h
48 h ~24 h 72 h 96 h
Onset of Preparation of Injection of Recovery of CEC-I Recovery of CEC-I Egg Incubation Shell-less Cultures CEC-I Injection of CEC-II Evaluation
Chick Culture Treatment Groups
Figure 3.2. Experimental design for the culture of 2-cell bovine embryos in the amnion of a developing chick embryo.
32 (M-2393, Sigma-Aldrich Inc., St. Louis, MO) and 0.1 ml of 0.5% phenol red (P-0290, Sigma-Aldrich Inc., St. Louis, MO) per 500 ml of sterile water. BO-B stock solutions contain 154.0 mM NaHCO3 (S-8875, Sigma-Aldrich Inc., St. Louis, MO) and 0.04 ml of 0.5% phenol red per 200 ml of sterile water. CR1aa medium was prepared as previously described by Rosenkrans and First (1994). Briefly, CR1aa stock solution was prepared by adding 114.7 mM NaCl, 1.6 mM
KCl, 26.2 mM NaHCO3, 0.3999 mM pyruvic acid (P-4562, Sigma-Aldrich Inc., St. Louis, MO), 2.5 mM L(+) lactic acid (L-4388, Sigma-Aldrich Inc., St. Louis, MO), 0.5195 mM glycine (G-8790, Sigma-Aldrich Inc., St. Louis, MO), 0.5051 mM L-alanine (A-7469, Sigma-Aldrich Inc., St. Louis, MO) and 0.2 ml of 0.5% phenol red per 100 ml of sterile water. Fertilization stock medium (BO-AB) was prepared by adding 1.2 mM pyruvic acid, 0.05 ml of gentamicin (50 mg/ml; 15750-060, Gibco, Grand Island, NY) and 0.018 ml of heparin sodium (10,000 IU/ml; Elkins-Sinn, Cherry Hill, NJ) per 38 ml of BO-A stock solution and 12 ml of BO-B stock solution. Fertilization medium (BO-caffeine) was prepared by adding 0.0243 g of caffeine sodium (C-4144, Sigma-Aldrich Inc., St. Louis, MO) per 25 ml of BO-AB. Sperm washing medium (0.6% BSA-BO) was prepared by adding 0.03 g of bovine serum albumin (BSA; Fraction V, A-4503, Sigma-Aldrich Inc., St. Louis, MO) to 5 ml of BO-AB. Oocyte washing medium (0.3% BSA-BO) was prepared by adding 0.03 g of BSA (Fraction V) to 10 ml of BO-AB. TCM oocyte washing medium was prepared by adding 0.03 g of BSA (Fraction V) to 10 ml of Tissue Culture Medium-199 (Eagle’s salt) with 25 mM HEPES Buffer (TCM-199; Gibco Laboratories, Grand Island, NY). CR1aa culture medium for day 0 to day 3 was prepared by adding 0.2 ml of Basal Medium Eagle (BME) amino acid solution (50X, B-6766, Sigma-Aldrich Inc., St. Louis, MO), 0.1 ml of Modified Eagle Medium (MEM) amino acid solution (10X, 11140- 050, Gibco Laboratories, Grand Island, NY), 0.01 ml of gentamicin, 1.1 mM L-glutamine (G-5763, Sigma-Aldrich Inc., St. Louis, MO) and 0.03 g of BSA (fatty acid free, A-7511, Sigma-Aldrich Inc., St. Louis, MO) to 9.5 ml of CR1aa stock solution. CR1aa culture medium for day 3 to day 7 was prepared by adding 0.2 ml of BME amino acid solution, 0.1 ml of MEM amino acid solution, 0.5 ml of FBS (SH30070.02; HyClone, Logan, UT),
33 0.01 ml of gentamicin, 1.1 mM L-glutamine and 0.03 g of BSA (fatty acid free) to 9.5 ml of CR1aa stock solution. In Vitro Fertilization Oocytes were purchased weekly from a commercial supplier (BoMed, Inc., Madison, WI) and matured in a portable incubator (~38.5°C) during the overnight travel interval (20 to 22 hours). Upon arrival at the laboratory, shipping vials were removed from the portable incubator and stored in a humidified atmosphere of 5% CO2 in air at 39°C until insemination. Semen was prepared by thawing a straw of frozen semen from a Holstein bull (CSS 7H5188, Genex Cooperative Inc., Shawano, WI) of proven fertility in a 39° to 40°C water bath for ~45 seconds. After thawing, the contents of the straw were deposited into a 15-ml plastic conical tube (Corning, New York, NY). Then, 9 ml of BO-caffeine was added to the tube and the suspension was centrifuged for 6 minutes at 200 x g. After centrifugation, the supernatant was removed and the remaining pellet was resuspended in an additional 9 ml of BO-caffeine. The resulting suspension was centrifuged for 6 additional minutes at 200 x g. The supernatant was removed again and the pellet resuspended in 4 ml of BO-caffeine and 4 ml of 0.6% BSA-BO and placed in a 38°C incubator for ~10 minutes to equilibrate. After equilibration, four 80-µl insemination drops were created in a preheated 35 x 10-mm Falcon® plastic petri dish (Becton Dickinson, Lincoln Park, NJ) and covered with warmed mineral oil (M-8140, Sigma-Aldrich Inc., St. Louis, MO). After the 20 to 22 hour maturation period, cumulus oocyte complexes were removed from the shipping vials and washed through two 35 x 10 mm plastic petri dishes containing 2 to 3 ml of 0.3% BSA-BO. Then, 10 to 20 oocytes were placed in each of the 80-µl insemination drops previously prepared and were co-incubated for 5 hours at 38°C and 5% CO2 in air. After fertilization, oocytes were washed through a single 35 x 10-mm plastic petri dish of TCM oocyte washing medium. Cumulus cells were removed by vortexing for 3 minutes in 1 ml of TCM-199 containing 0.0010 g of hyaluronidase (H-3506, Sigma- Aldrich Inc., St. Louis, MO). After vortexing, oocytes were washed through two 35 x 10- mm plastic petri dishes containing TCM oocyte washing medium and one 35 x 10-mm plastic petri dish of CR1aa. Then, 10 to 15 oocytes per drop were placed in 50-µl drops
34 of CR1aa covered with warmed mineral oil and incubated in a modular incubator containing 5% CO2, 5% O2 and 90% N2 at 39°C. Chick Embryo Co-Culture The basic procedure for the culture of bovine 8-cell IVF embryos using the chick embryo amnion was modified from that previously described by Blakewood and Godke (1989). Briefly, fertilized domestic chicken eggs were collected on the day of oviposition and stored at 10°C until the onset of incubation. Four days prior to the introduction of bovine embryos, the eggs were removed from cold storage and allowed to come to room temperature (25°C) before placing them in a 37°C commercial egg incubator. After 72 hours of incubation, the eggs were rinsed with a solution of 70% ethanol (Aaper Alcohol and Chemical Company, Shelbyville, KY) and then coated with 7.5% providone iodine solution (Betadine®, Purdue Fredrick Company, Stanford, CT) and allowed to air dry in a horizontal position to ensure proper positioning of the chick embryo. To remove the egg contents, a crack was made in the shell by striking the center of the egg against the rim of a sterile beaker. The egg contents were then released into a 100-ml plastic embryo collection dish (Veterinary Concepts, Spring Valley, WI) covered by a piece of cellophane kitchen wrap (Saran®, Dow Chemical, Midland, MI). The dish, including the egg contents, was loosely capped with a plastic lid (Veterinary Concepts, Spring Valley, WI) and maintained in an incubator at 38°C for an additional 24 hours before introducing the bovine embryos into the amniotic cavity of the chick embryo. Prior to embedding and injection of bovine embryos, suitable chick embryos were selected for injection based on the following criteria: 1) round intact yolk, 2) good vascularity in the chorioallantoic membrane, 3) vigorous heartbeat and 4) no apparent abnormalities. The procedure for embedding embryos in agarose was modified from that of Willadsen (1979) for the culture of ovine embryos in the ligated oviducts of sheep. On day 0 of culture, two or three 8-cell bovine IVF embryos were placed in a 50-µl drop of low-melting agarose solution [1.2% agarose (A-9799, Sigma-Aldrich Inc., St. Louis, MO) dissolved in Dulbecco’s Phosphate-Buffered Saline (DPBS; D1408, Sigma-Aldrich Inc., St. Louis, MO)] at ~37oC and then aspirated into a beveled glass pipette (200 µm outer diameter). After holding in air for ~30 seconds, the agar (containing the embryos) within
35 the pipette was expelled as a gel cylinder into room temperature TL-HEPES (BioWittaker, Walkersville, MD). The agarose cylinder, including the bovine embryos, was then aspirated back into the beveled pipette and carefully expelled into the amniotic cavity of a 96-hour chick embryo. After introducing the embedded embryos into the amniotic cavity, the shell-less culture systems were returned to incubation for an additional 72 hours at 39oC. Thus, a single agarose cylinder containing two or three 8-cell bovine IVF embryos was co- cultured in the amnion of each developing chick embryo for 3 to 4 days. On day 3 or day 7 of in vitro culture, the amnion containing the agarose cylinder was dissected away from the rest of the egg components, placed in a 35 x 10-mm plastic petri dish and rinsed several times with DPBS containing 1% FBS. After rinsing, the petri dish was placed under a stereomicroscope (Nikon SMZ-2B, Tokyo, Japan) equipped with an overhead light source and the agarose cylinder was removed from the amniotic cavity by using two 22-gauge, 37.5-mm needles (Kendall, Mansfield, MA) to make a small opening in the amniotic membrane. The agarose cylinder was carried from the amnion with the escaping amniotic fluid, settling on the bottom of the dish. The cylinder was then washed twice in CR1aa medium and dissected using two 22-gauge needles to isolate and remove the developing embryos. Statistical Analysis For both experiments, data were analyzed using PROC GLM of the SAS Statistical Package (v. 8e, 1999, SAS Institute, Inc., Gary, NC). Significant differences between embryo developmental rates (Experiment 3.1 and Experiment 3.2) and embryo quality grade (Experiment 3.2) in each group were tested using a post hoc least significant difference (LSD) test. Differences with a probability level (P) of 0.05 or less were considered significant. RESULTS In Experiment 3.1, 60 chicken eggs were initially placed in incubation. After 72 hours of incubation, the eggs were removed from their shell and placed in shell-less culture. At this time, six (10%) of the eggs were found to be unfertilized, four (7%) contained embryos that failed to develop and one (2%) contained an embryo that was nonviable. The remaining 49 eggs were placed in shell-less culture and returned to
36 incubation. After 24 hours of incubation in shell-less culture, 21 eggs were selected for injection of bovine embryos. In the CEC-I treatment group, 29 of 36 (81%) embryos were recovered from the amniotic cavity of nine live chicks after 72 hours of culture. Of the 29 embryos recovered, 16 (55%) had successfully cleaved with seven (24%) of those appearing to be viable 6- to 8-cell embryos (Table 3.1). Overall cleavage rate of the CEC-I treatment group was significantly lower (P<0.05) than the control (34/36), agar (33/36) and CEC-II treatment groups (29/36). Similarly, there were significantly fewer (P<0.05) 6- to 8-cell embryos in the CEC-I treatment group compared with the control (25/36), agar (23/36) or CEC-II (21/36) treatment groups. At this time, the CEC-II embryos (n=36) were injected into the amniotic cavity of nine chick embryos (four embryos per chick amnion). These embryos were recovered after 96 hours of culture. On day 7 of culture, 32 of the 36 (89%) embryos originally injected on day 3 were successfully recovered from the chick amnion. Of these, six (20%) had developed to the blastocyst stage. This was not significantly different (P>0.05) from the total number of blastocysts in the other treatment groups [control = 16/36 (44%); agar = 17/36 (47%); CEC-I = 2/29 (7%)]. However, there were significantly fewer (P<0.05) total blastocysts in the CEC-I treatment group when compared with the control or agar treatment groups (Table 3.1). There was no significant difference (P>0.05) in the number of morulae, early blastocysts, blastocysts or expanded blastocysts between the four treatment groups (Figure 3.3). In Experiment 3.2, 50 chicken eggs were initially placed in incubation. When the eggs were prepared for shell-less culture, two (4%) eggs were unfertile, six (12%) contained embryos that failed to develop and three (6%) contained embryos that were nonviable. The remaining 36 eggs were placed in shell-less culture and returned to incubation. After 24 hours of incubation in shell-less culture, 13 eggs were selected for injection of bovine embryos. In the CEC-I treatment group, 28 of 40 (70%) embryos were recovered from the amniotic cavity of nine live chicks after 72 hours of culture. Of the 28 embryos recovered, 17 (61%) appeared to be viable 6- to 8-cell embryos and seven (25%) had developed past the 8-cell stage (Table 3.2). There were no significant differences (P>0.05) in development between the four treatment groups. At this time, the CEC-II
37
Table 3.1. Development of 1-cell bovine embryos cultured in CR1aa + 5% fetal bovine serum (control) or co-cultured in the chick amnion from day 0 to day 3 (CEC-I) or from day 3 to day 7 (CEC-II)
a Number of 1-cell embryos allotted to each treatment. The number in parenthesis indicates the number of embryos successfully recovered from live chick embryos on day 3 (CEC-I) or day 7 (CEC-II). b The percentage of embryos developing in the chick co-culture groups (CEC-I and CEC-II) were based on the number of embryos successfully recovered from live chick embryos. c MORL = morulae. d BLST = blastocysts. e,f Within a column, numbers without a common superscript are significantly different (P<0.05).
38
A Control Agar CEC-I CEC-II
100 a a 90 a 80 a 70
s a
o a y
r 60 b b m 50 b E
f 40 % o a a a 30 a b 20 a a 10 a 0 1-cell 2-4 cell 6-8 cell cleavage
B Control Agar CEC-I CEC-II
50
40
s a o y 30 br a a
m a E 20 a of a % a a 10 a a a a a a a a 0 MORL Early BLST BLST Expanded BLST
Figure 3.3. Development of 1-cell bovine embryos cultured in CR1aa + 5% fetal bovine serum (control) or co-cultured in the chick amnion from day 0 to day 3 (CEC-I) or from day 3 to day 7 (CEC-II) on day 3 of development (A) and on day 7 of development (B). Within a developmental stage, columns without a common superscript are significantly different (P<0.05).
39
Table 3.2. Development of 2-cell bovine embryos cultured in CR1aa + 5% fetal bovine serum (control) or co-cultured in the chick amnion from day 0 to day 3 (CEC-I) or from day 3 to day 7 (CEC-II)
Day 3 Day 7
Treatment Total Grade 1 group na % 2-4 cellb % 6-8 cellb % >8 cellb % MORLbc % BLSTbd % BLSTbd
Control 39 21 ± 7e 54 ± 8e 3 ± 5e 5 ± 4e 64 ± 8e 26 ± 7e
Agar 39 21 ± 7e 62 ± 8e 15 ± 6e 5 ± 4e 38 ± 8e 23 ± 7e
CEC-I 40 (28) 7 ± 5e 61 ± 9e 25 ± 8e 7 ± 5e 42 ± 10e 31 ± 12e
CEC-II 39 (19) 23 ± 7e 46 ± 8e 21 ± 7e 11 ± 7e 37 ± 11e 16 ± 9e
a Number of 2-cell embryos allotted to each treatment. The number in parenthesis indicates the number of embryos successfully recovered from live chick embryos on day 3 (CEC-I) or day 7 (CEC-II). b The percentage of embryos developing in the chick co-culture groups (CEC-I and CEC-II) were based on the number of embryos successfully recovered from live chick embryos. c MORL = morulae. d BLST = blastocysts. e Within a column, numbers without a common superscript are significantly different (P<0.05).
40 embryos (n=39) were injected into the amniotic cavity of 13 chick embryos (three embryos per chick). These embryos were recovered after 96 hours of culture. On day 7 of culture, 19 of the 39 (49%) embryos originally injected on day 3 were successfully recovered from the chick amnion. Of these, seven (37%) had developed to at least the early blastocyst stage. This was not significantly different (P>0.05) from the total number of blastocysts in the other treatment groups [control = 25/39 (64%); agar = 15/39 (38%); CEC-I = 12/28 (25%)]. There was no significant difference (P>0.05) in the number of embryos developing to the morula or early blastocyst stage of development between the four treatment groups (Figure 3.4). More embryos developed to the blastocyst stage in the CEC-I group than in the CEC-II group (P<0.05). However, there were more expanded blastocysts in the CEC-II group than in either the agar control group or the CEC-I group (P<0.05). The number of embryos developing to the hatching or hatched blastocyst stage did not differ (P>0.05) between treatments. There was no significant difference (P>0.05) in the number of grade1, grade 2 or grade 4 embryos when embryos in the four treatments were evaluated for quality (Figure 3.5). There were significantly more (P<0.05) grade 3 embryos in the control group when compared with the agar group; however, neither group was significantly different (P>0.05) from either chick treatment. DISCUSSION In the early 1990s, the preferred method of culturing bovine embryos in vitro involved the inclusion of co-culture cells in the embryo production system (Gandolfi and Moor, 1987; Eyestone and First, 1989). In vitro embryo production systems have progressed, and co-culture is no longer required for embryos to progress through the in vitro developmental block and develop to the blastocyst stage. However, despite the progress that has been made, current in vitro embryo culture conditions are unable to mimic the in vivo conditions of the oviducts and uterus. Moreover, in vitro embryo culture has been implicated in problems such as delayed embryo development (Van Soom et al., 1997), reduced embryo quality (Duby et al., 1997; Boni et al., 1999; Viuff et al., 1999), reduced pregnancy rates (Enright et al., 2000) and numerous fetal and/or calf abnormalities (Thompson et al., 1995; Walker et al., 1996; Sinclair et al., 1999). These problems have led researchers to develop and test alternative culture methods such as perifusion culture (Lim et al., 1996; McGowan and Thompson, 1997), microfluidic
41
A Control Agar CEC-I CEC-II
100
90
80
70 s a a
yo 60 a
br a m 50
E
of 40 % 30 a a a a a 20 a a a 10 0 2-4 cell 6-8 cell >8-cell
B Control Aga r CEC-I CEC-II
50
40
s o y r 30 a b a a m
E
f a ab 20 ab ab % o a a a 10 a a a b a b a b a a 0 MORL Early BLST BLST Expanded Hatching or BLST Hatched BLST
Figure 3.4. Development of 2-cell bovine embryos cultured in CR1aa + 5% fetal bovine serum (control) or co-cultured in the chick amnion from day 0 to day 3 (CEC-I) or from day 3 to day 7 (CEC-II) on day 3 of developmen t (A) and on day 7 of development (B). Within a developmental stage, columns without a common superscript are significantly different (P<0.05).
42
Control Agar CEC-I CEC-II
50
40
s a a yo r 30 a a b
m a a E f 20 a a % o a 10 ab b ab a a a a 0 Grade 1 Grade 2 Grade 3 Grade 4
Figure 3.5. Quality grade of bovine 2-cell embryos cultured in CR1aa + 5% fetal bovine serum (control) or co-cultured in the chick amnion from day 0 to day 3 (CEC-I) or from day 3 to day 7 (CEC-II) on day 7 of development. Within a quality grade, columns without a common superscript are significantly different (P<0.05).
43 “microchannel” systems (Glasgow et al., 1998) and the “well-of-well” (WOW) method (Vajta et al., 2000) for in vitro embryo culture. In the early 1990s, one such alternative culture system was developed in which mammalian embryos were cultured in the amnion of a living chicken embryo (Blakewood and Godke, 1989). The trials and tribulations of current in vitro embryo culture and the need to develop alternative culture environments have led us back to the chick embryo co-culture system. Results of the present study indicate that bovine embryos can develop in the environment of the chick amnion at rates similar to those of embryos cultured in a semi-defined culture medium (CR1aa in the present study). Culture of bovine embryos in the chick amnion from the 1-cell stage resulted in significantly fewer 6- to 8-cell embryos on day 3 of development and a significantly lower total blastocyst (early blastocyst, blastocyst and expanded blastocyst) rate on day 7. In contrast, there was no significant difference in development on day 3 or in total blastocysts on day 7 when chick embryo co-culture was initiated at the 2-cell stage. These findings are interesting and may indicate that embryos are particularly sensitive to culture environment during the first 24 hours of development. If this is in fact the case, inadequate conditions during this time could have profound impacts on embryo developmental potential. The low number of observations in the present study must be taken into account. However, further investigation into the effects of culture environment on the first 24 hours of embryo development is warranted. In theory, the chick embryo co-culture system should provide a more physiological “normal” environment for embryo development. Because the live chick embryo actively regulates the pH, osmolarity and O2 content of its own environment, the amnion should provide a more stable environment for mammalian embryo development than current “static” in vitro culture systems (Blakewood et al., 1989a). In addition, the chick embryo continuously synthesizes growth factors needed for its own development. These growth factors could provide a stimulus to mammalian embryo development that is currently unavailable in current embryo culture systems without the addition of serum or other exogenous growth factors (Blakewood et al., 1989a). In Experiment 3.2, there was no difference in the number of grade 1, grade 2 or grade 4 embryos when embryos in the four treatments were evaluated for quality. Numerically, the number of grade 3 embryos was greater in controls than in either chick
44 treatment. The lack of statistical differences is likely due to the low number of observations in each chick treatment group. These results are promising and suggest that chick embryo culture may in fact increase embryo quality over current culture conditions. As stated above, the low number of observations must be taken into account, but it is apparent that further research may be warranted. One major drawback of the chick embryo co-culture system, however, is the inherent mortality rate of shell-less chick embryos. Dunn and Boone (1976) reported that chick survival rates of 90 to 95% appear to be as high as can be expected with shell-less culture and a rapid degeneration of mammalian embryos has been reported following chick embryo death (Blakewood and Godke, 1989). The exact cause of the decline in mammalian embryo viability following chick embryo death is not known, but it can be speculated that with the death of the chick comes alterations in amniotic fluid pH, osmolarity and temperature, as well as build-up of waste products. In conclusion, although bovine embryos can develop in the embryotropic environment of the chick embryo amnion, the use of this novel culture system does not appear to be advantageous for embryo development when compared with the semi- defined culture medium, CR1aa. This approach, however, should not be ruled out. Further research should be conducted to determine the effects of the avian amniotic fluid on embryo quality, embryo metabolism and pregnancy and calving rates.
45 CHAPTER IV
USE OF THE AMNIOTIC CAVITY OF THE DEVELOPING CHICK EMBRYO FOR CULTURING BOVINE 8-CELL EMBRYOS DERIVED FROM SOMATIC CELL NUCLEAR TRANSFER (NT)*
INTRODUCTION Somatic cell NT has been utilized in cattle for production of transgenic animals as well as to produce copies of elite animals, but success rates are low (Cibelli et al., 1998; Hill et al., 1999; Shiga et al., 1999; Wells et al., 1999; Hill et al., 2000b). It remains to be determined whether this inefficiency is a result of the NT procedure itself or conditions in which the embryos are cultured following the NT procedure. Despite sustained efforts in laboratories throughout the world, present in vitro culture conditions are unable to fully replicate in vivo conditions. In vitro culture conditions are routinely enhanced by the addition of serum to the culture medium (Bavister, 1995). It has been suggested that serum may contain components advantageous for blastoceol development, such as growth factors or extracellular matrix components like fibronectin (Pinyopummintr and Bavister, 1994). However, it is hypothesized that the components of serum may also contribute to fetal and calf abnormalities (Thompson et al., 1995; Walker et al., 1996; Sinclair et al., 1999). Calves produced from embryos cultured in vitro often exhibit increased birth weight (Willadsen et al., 1991; Keefer et al., 1994; Behboodi et al., 1995), longer gestation lengths (Walker et al., 1996), increased placental anomalies (Hasler et al., 1995; Hill et al., 1999, 2001, 2002) and increased incidences of abortion (Walker et al., 1996; Hill et al., 1999) and perinatal (Behboodi et al., 1995; Wilson et al., 1995; Garry et al., 1996; Schmidt et al., 1996; Kruip and den Daas, 1997; Hill et al., 1999) and neonatal mortality (Schmidt et al., 1996; Kruip and den Daas, 1997; Hill et al., 1999). Furthermore, bovine viral diarrhea virus (BVDV) may be introduced into in vitro production systems via the addition of serum, extracted components of serum or somatic cell lines (Stringfellow et al., 2000). BVDV is a ubiquitous pathogen that can cause abortion resulting in severe economic losses, and undetected association of BVDV in
* Used with permission of Molecular Reproduction and Development, Wiley-Liss,Inc., Hoboken, NJ
46 transferred in vitro-produced embryos may result in early embryonic death or abortion. Shin et al. (2000) have reported that the rate of pregnancy loss between 30 and 40 days of gestation was significantly greater in bovine fetuses derived from BVDV- infected somatic cells than in those from noninfected somatic cell lines. Total elimination of materials of bovine origin, including serum and serum albumin, from the standard oocyte maturation and embryo or somatic cell culture systems could reduce the risk of virus introduction. Previous reports indicate that factors within the amniotic cavity of developing chick embryos are capable of supporting the maturation of porcine oocytes (Ocampo et al., 1994) and development of embryos in several mammalian species. In these studies, researchers found significantly more expanded and hatched blastocysts compared with controls in mice (control = Whitten's medium), goats (control = Ham's F-10), cattle (control = Ham's F-10; Blakewood et al., 1989a, 1989b, 1990) and pigs (control = mBMOC + POEC) (Ocampo et al., 1993). Unfortunately, most of these studies used a relatively low number of observations per treatment group (mean of 40 observations per treatment across studies). In addition, the media used in these studies are now considered somewhat obsolete by today’s standards. Even with these drawbacks, this technique warrants further investigation as an alternative in vitro embryo culture system that is free from materials of bovine origin for the culture of in vitro-produced mammalian embryos. In the present study, the development of bovine 8-cell embryos produced by somatic cell NT was assessed after culture in the chick embryo amnion. The viability of bovine embryos developing into morulae or blastocysts after culture in the chick amnion was further investigated by transferring a representative sample of these embryos into recipient females. This study was conducted as a preliminary trial in evaluating the use of this unique co-culture system as an alternative to present in vitro embryo culture systems used for the production of somatic cell NT calves. MATERIALS AND METHODS Experimental Design The design for this experiment is illustrated in Figure 4.1. After reconstruction (day=0), all embryos were cultured in vitro in CR1aa medium for 3 days. Viable 8-cell embryos were then randomly allotted to one of two treatment groups for in vitro culture.
47
Onset of Egg Incubation
72 hours Nuclear Transfer In Vitro Culture in CR1aa
72 hours Preparation of Shell-less Cultures
Selection of
8-cell Embryos 24 hours
Culture in Injection of 8-cell Embryos CR1aa + 5% FBS into Chick Amnion
72 hours 72 hours
Recovery Evaluation Evaluation Embryo Transfer
Figure 4.1. Experimental design for the culture of bovine 8-cell embryos derived from somatic cell nuclear transfer. Culture was either in vitro in CR1aa supplemented with 5% fetal bovine serum (control) or co-cultured in the chick embryo amnion.
48 Embryos allotted to Treatment A (control) were cultured in CR1aa supplemented with 5% fetal bovine serum (FBS) from day 3 to day 6 post-reconstruction. Embryos allotted to Treatment B were cultured in the amniotic cavity of 96-hour chick embryos from day 3 to day 6 post-reconstruction. Embryos that had reached the morula and blastocyst stages after culture in the chick embryo amnion on day 6 of culture were transferred to recipient females. Experimental Methods Media Preparation CR1aa medium was prepared according to Rosenkrans and First (1994). First, CR1aa stock solution was prepared by adding 114.7 mM NaCl (S-5886, Sigma-Aldrich Inc., St. Louis, MO), 1.6 mM KCl (P-5405, Sigma-Aldrich Inc. St. Louis, MO), 26.2 mM
NaHCO3 (S-8875, Sigma-Aldrich Inc., St. Louis, MO), 0.3999 mM pyruvic acid (P-4562, Sigma-Aldrich Inc., St. Louis, MO), 2.5 mM L(+) lactic acid (L-4388, Sigma-Aldrich Inc., St. Louis, MO), 0.5195 mM glycine (G-8790, Sigma-Aldrich Inc., St. Louis, MO), 0.5051 mM L-alanine (A-7469, Sigma-Aldrich Inc., St. Louis, MO) and 0.2 ml of 0.5% phenol red per 100 ml of sterile water. For culture from day 0 to day 3, CR1aa culture medium was prepared by adding 0.2 ml of Basal Medium Eagle (BME) amino acid solution (50X, B-6766, Sigma-Aldrich Inc., St. Louis, MO), 0.1 ml of Modified Eagle Medium (MEM) amino acid solution (10X, 11140-050, Gibco Laboratories, Grand Island, NY), 0.01 ml of gentamicin, 1.1 mM L- glutamine (G-5763, Sigma-Aldrich Inc., St. Louis, MO) and 0.03 g of bovine serum albumin (BSA; fatty acid free, A-7511, Sigma-Aldrich Inc., St. Louis, MO) to 9.5 ml of CR1aa stock solution. For culture from day 3 to day 7, CR1aa culture medium was prepared by adding 0.2 ml of BME amino acid solution, 0.1 ml of MEM amino acid solution, 0.5 ml of FBS (SH30072.02; HyClone, Logan, UT), 0.01 ml of gentamicin, 1.1 mM L-glutamine and 0.03 g of BSA (fatty acid free) to 9.5 ml of CR1aa stock solution. Preparation of Adult Bovine Donor Cells The donor cells used for the NT procedure were established from skin biopsy samples of a 3-year-old Brahman cow. Briefly, skin biopsies were obtained from the dorsal aspect of the shoulder directly over the supraspinatus muscle using a 6-mm diameter biopsy punch (Miltex Instrument Co, Betpage, NY). Using heat sterilized iris scissors, the dermal portion of the biopsy was finely minced into pieces measuring
49 ~1 mm2. Culture medium (mDMEM) (Cibelli et al., 1998; Shiga et al., 1999) consisting of Dulbeco’s Modified Eagle Medium (DMEM; Gibco Laboratories, Grand Island, NY), 10% FBS and 1 µl/ml of gentamicin was added and the tissue suspension transferred to a 15- ml conical tube (Corning, New York, NY). The tissue suspension was washed by swirling and the pieces of tissue were allowed to settle to the bottom of the conical tube. The supernatant was discarded and 3 ml of fresh mDMEM was added. This washing process was repeated two additional times. After the last wash, the tissue was resuspended in 3 ml of mDMEM and transferred to a 35 x 10-mm Falcon® plastic petri dish (Becton and
Dickinson, Lincoln Park, NJ). All cultures were maintained at 39°C and 5% CO2 in a humidified atmosphere. Once a fibroblast cell layer was established, the cells were maintained in 25-cm2 tissue culture flasks (CostarTM, Cambridge, MA) until they reached ~95% confluency. Cells were then trypsinized (Trypsin EDTA, 1x; T3924, Sigma-Aldrich Inc., St. Louis, MO), counted on a hemocytometer and reseeded at a density of 100,000 cells per 25- cm2 tissue culture flask. The cells were passaged twice prior to cryopreservation in DMEM supplemented with 20% FBS and 5% dimethylsulfoxide (DMSO; D128-500, Fischer Scientific, Fairlawn NJ) and then frozen in aliquots of 500,000 cells per cryovial. To prepare adult fibroblasts as donors for NT, cells were thawed and cultured in mDMEM in Nunc® 4-well plates (Apogent Technologies, Rochester, NY). Nuclear Transfer Oocytes were obtained on a weekly basis from a commercial source (BoMed, Madison, WI). Oocytes were matured in a portable incubator (~38.5oC) during the overnight travel interval for 20 to 22 hours. After arrival, the cumulus cells were removed from the oocytes by vortexing in 1 ml of TCM-199 (Eagle’s salt) with 25 mM HEPES buffer (TCM-199, Gibco, Grand Island, NY) containing 0.001 g of hyaluronidase (H-3506, Sigma-Aldrich Inc., St. Lois, MO) for 3 minutes. The denuded oocytes were washed twice in modified TCM (mTCM: 19 ml of TCM-199 supplemented with 1 ml of FBS and 0.02 ml of gentamicin) and were selected for the presence of a first polar body. Micromanipulation was performed in a 200-µl drop of manipulation medium [4.0 ml of Dulbecco’s Phosphate Buffered Saline (DPBS; D-1408, Sigma-Aldrich Inc., St. Louis, MO) containing 1 ml of FBS, 0.005 ml of gentamicin and 0.005 ml of cytochalasin D (C-8273, Sigma-Aldrich Inc., St. Louis, MO)] overlaid with warmed mineral oil (M-8140,
50 Sigma-Aldrich Inc., St. Louis, Mo). All manipulations were conducted on an inverted microscope (Nikon Diphot, Tokyo Japan) equipped with a pair of Lietz micromanipulators. The NT procedure was modified from that previously described by Shiga et al. (1999). Briefly, oocytes were held in place by negative pressure applied to a holding pipette (120 µm outside diameter). After brief exposure (<10 seconds) to ultraviolet light, the first polar body and adjacent cytoplasm containing the nuclear material, as visualized by Hoechst-33342 (5 µg/ml; bisBenzimide; B-2261; Sigma-Aldrich Inc., St. Lois, MO) staining, were manipulated out of a rent in the zona pellucida using a flexible glass needle. A pipette (25 µm outer diameter) containing the donor cell (passages 2 to 8) was introduced through the rent in the zona pellucida made during enucleation, and the cell was wedged between the zona and vitelline membrane to ensure close membrane contact required for subsequent fusion. The reconstructed cytoplast-karyoplast couplets were induced to fuse in a microslide fusion chamber with stainless steel electrodes separated by a 1-mm gap. The chamber was filled with fusion buffer consisting of 0.3 M mannitol (D-3651, Sigma-
Aldrich Inc., St. Louis, MO), 0.05 mM CaCl2·2H2O (C-7902, Sigma-Aldrich Inc., St. Louis,
MO) and 0.1 mM MgSO4 (M-7774, Sigma-Aldrich Inc., St. Louis, MO) and the couplets were fused with two direct current pulses of 2.25 kV/cm for 15 microsecconds, delivered by a BTX Electrocell Manipulator 200 unit (Genetronics, San Diego, CA). The fused couplets were further activated with 10 µg/ml of cyclohexamide (C-7698, Sigma-Aldrich Inc., St. Louis, MO) in CR1aa medium for 4 hours. After activation, embryos were cultured in 50-µl drops of CR1aa medium in a modular incubator containing 5% CO2, 5% O2 and 90% N2 (activation = day 0). On day 3 of culture, good quality (as indicated by equal size, well defined blastomeres), 8-cell NT embryos were randomly selected for in vitro culture in CR1aa supplemented with 5% FBS (control group) or co-culture in the amniotic cavity of a developing chick embryo as described below. Chick Embryo Co-Culture A technique modified from that originally described by Blakewood and Godke (1989) was used for the shell-less culture of chick embryos. Fertilized domestic chicken eggs were collected on the day of oviposition and stored at 10°C until the onset of
51 incubation. Four days prior to introduction of bovine embryos into the chick embryo amnion the eggs were removed from cold storage and allowed to come to room temperature (25°C). They were then placed in a 37°C commercial egg incubator. After 72 hours of incubation, the eggs were rinsed with a solution of 70% ethanol (Aaper Alcohol and Chemical Company, Shelbyville, KY) and then coated with 7.5% providone iodine (Betadine®, Purdue Fredrick Company, Stanford, CT) and allowed to air dry in a horizontal position to ensure proper positioning of the chick embryo. To remove the egg contents, the egg shell was cracked against the rim of a sterile beaker and the egg contents were released into a 100-ml plastic embryo collection dish (Veterinary Concepts, Spring Valley, WI) covered by a piece of cellophane kitchen wrap (Saran®, Dow Chemical, Midland, MI). The dish, including the egg contents, was loosely capped with a plastic lid (Veterinary Concepts, Spring Valley, WI) and maintained in an incubator at 38°C for an additional 24 hours before introducing the bovine embryos into the amniotic cavity of the chick embryo. Prior to embedding and injection of bovine embryos, suitable chick embryos were selected for injection based on the following criteria: 1) round intact yolk, 2) good vascularity in the chorioallantoic membrane, 3) vigorous heartbeat and 4) no apparent abnormalities. A technique modified from that originally described by Willadsen (1979) was used for embedding embryos in agarose. On day 3 of culture, two or three 8-cell bovine NT embryos were placed in a 50-µl drop of low-melting agarose solution [1.2% agarose (A- 9799; Sigma-Aldrich Inc., St. Louis, MO) dissolved in DPBS] at ~37oC. The embryos were then aspirated into a beveled glass pipette (200 µm outer diameter). After holding in air for ~30 seconds, the agar (containing the embryos) within the pipette was expelled as a gel cylinder into room temperature TL-HEPES (BioWittaker, Walkersville, MD). The agarose cylinder, including the bovine embryos, was then aspirated back into the beveled pipette, and carefully expelled into the amniotic cavity of a 96-hour chick embryo. After introducing the embedded embryos into the amniotic cavity the shell-less culture systems were returned to incubation for an additional 72 hours at 39oC. Thus, a single agarose cylinder containing two or three 8-cell bovine NT embryos was co- cultured in the amnion of each developing chick embryo for 3 days.
52 On day 6 of in vitro culture, the amnion containing the agarose cylinder was dissected away from the rest of the egg components, placed in a 35 x 10-mm plastic petri dish and rinsed several times with DPBS containing 1% FBS. Under a stereomicroscope (Nikon SMZ-2B, Tokyo, Japan), the agarose cylinder was removed from the amniotic cavity by using two 22-gauge, 37.5 mm needles (Kendall, Mansfield, MA) to make a small opening in the amniotic membrane. The agarose cylinder was carried from the amnion with the escaping amniotic fluid and allowed to settle on the bottom of the dish. The cylinder was then washed twice in CR1aa medium and the developing embryos were dissected out of the agar using two 22-gauge needles. Embryos that had reached the morula and blastocyst stages after culture in the chick embryo amnion (day 6) were nonsurgically transferred to naturally cycling beef females (age = 2 to 10 years; BCS = 5 to 7) on days 6 to 7 of their estrous cycle (estrus = day 0) when recipients were available. Pregnancy status of recipient females was diagnosed by ultrasonography (Aloka, SSD-500V, Japan) on day 30 of gestation. Statistical Analysis Data are expressed as means and were analyzed using PROC GLM of the SAS Statistical Package (v. 8e, 1999, SAS Institute, Inc., Gary, NC). Significant differences between means for embryo development rates in each group were tested using a post hoc, Fisher’s protected least significant difference test (PLSD test). Differences with a probability (P) value of 0.05 or less were considered significant in this study. RESULTS For preparation of shell-less chick embryos, 210 chicken eggs were initially placed in incubation. After 72 hours of incubation, the eggs were removed from their shell and placed in shell-less culture. At this time, 25 (12%) of the eggs were found to be unfertilized, 21 (10%) contained embryos that failed to fully develop and 11 (5%) contained an embryo that was nonviable. The remaining 153 eggs were returned to incubation. After 24 hours of incubation in shell-less culture, 39 eggs were selected for injection of bovine embryos. Of 485 cytoplast-karyoplast couplets reconstructed, 387 (79.8%) successfully fused during the NT procedure. Of the 372 viable fused embryos, 232 (62.4%) developed to the 8-cell stage by day 3 of culture in CR1aa; 12 of these embryos were discarded due to poor quality. Of the 220 8-cell embryos remaining, 113 embryos were
53 randomly allotted for culture in CR1aa + 5% FBS (control). The remaining 107 8-cell embryos were embedded in agar and injected into the amniotic cavity of chick embryos. Of the embryos injected, 54 (50.5%) were successfully recovered for evaluation. Embryo loss occurred either during the manual recovery procedure or was due to chick embryo death. Ten of the 39 (25.6%) chick embryos originally injected did not survive shell-less incubation, accounting for the majority of loss of the bovine embryos. Developmental rates for bovine 8-cell embryos co-cultured in the chick-embryo amnion or cultured in CR1aa medium containing 5% FBS (control) are summarized in Table 4.1. There was no effect (P>0.05) of culture system on the percentage of 8-cell embryos that developed to the morula (control = 54.2% and chick embryo co-culture = 42.6%) or blastocyst (control = 19.8% and chick embryo co-culture = 17.2%) stage on day 6 of culture. A total of 21 morula to blastocyst stage embryos from the chick embryo co-culture group (Figure 4.2) were transferred nonsurgically to 7 synchronized recipient females (2 to 4 embryos per female) on day 6 or day 7 of their estrous cycle (estrus = day 0) corresponding to day 6 of culture. Of those recipients, three females (42.9%) were diagnosed (via ultrasonography and visualization of fetal heartbeats) pregnant with singletons on day 30 of gestation (Figure 4.3). By day 120 of gestation, however, the recipients had lost their pregnancies. DISCUSSION Previous reports have shown that an embryo co-culture system using domestic chicken egg shell-less culture is capable of supporting the development of pre-hatching mammalian embryos (Blakewood and Godke, 1989; Blakewood et al., 1989a, 1989b), and live transplant offspring have been produced following chick embryo co-culture of 2- to 8-cell caprine embryos. (Blakewood et al.,1990). Results of the present study indicate that the chick embryo amnion is also capable of supporting development of bovine 8-cell embryos derived from somatic cell NT at rates comparable with embryos cultured in vitro in CR1aa medium containing 5% FBS. Supplementation of bovine embryo culture medium with serum from the initiation of compaction, as performed here in the control group, is widely practiced to stimulate blastoceol development, despite the undefined and variable nature of its components (Pinyopummintr and Bavister, 1994; Thompson et al., 1998). However, addition of serum has been implicated in several fetal and/or calf abnormalities including increased birth
54
Table 4.1. Day 6 development of somatic cell nuclear transfer bovine 8-cell embryos co-cultured in the chick embryo amnion
Treatment No. embryos for No. (%) a c fh gh fgh group culture recovered % MORL % BLST % MORL & BLST
Control culture medium 113 -- 54.2f 19.8f 74.1f
Chick embryo d e f f f co-culture 107 54 (51) 42.6 17.2 59.9
a Seven replications of nuclear transfer were performed in this experiment. b Control 8-cell nuclear transfer embryos were cultured in vitro in CR1aa medium supplemented with 5% FBS. c Number of 8-cell embryos allotted to each treatment . d Number of 8-cell embryos embedded in agar and injected into the amniotic cavity of live chick embryos. e Number (%) of bovine embryos successfully recovered from the amniotic cavities of surviving chick embryos on day 6 of culture. f MORL = morulae. g BLST = blastocysts. h Embryo development was assessed on day 6 of culture. The percentage was based on the number originally cultured and number recovered in the control groups and chick embryo co-culture, respectively. There was no significant difference (P>0.05) between the mean number of morulae and blastocysts in the in vitro culture control and chick embryo co-culture groups.
55
Figure 4.2. Bovine somatic cell nuclear transfer embryos developed into morulae/blastocysts (day 6 of in vitro culture) after co-culture in the amniotic cavity of a developing chick embryo for 3 days. These embryos were agarose embedded (agar cylinders are indicated by arrows) at the 8-cell stage following in vitro culture in CR1aa medium from day 0 to day 3 of in vitro culture and then injected into the amnion of a 96- hour live chick embryo.
Figure 4.3. Ultrasound image of a fetus (day 30) derived from bovine somatic cell nuclear transfer embryos that were co-cultured in the amnion of a developing chick embryo from day 3 to day 6 following in vitro culture in CR1aa medium.
56 weight and abortion of NT-derived conceptuses (Walker et al., 1996). Also, Stringfellow et al. (2000) indicated that materials of bovine origin, including serum and extracted components of serum, used in bovine in vitro embryo production systems could possibly originate from cattle that were exposed to BVDV. The BVDV infection has been implicated as a cause of early embryonic death or abortion in cattle (Shin et al., 2000). Although the embryotropic properties of avian amniotic fluid remain largely undefined, amniotic fluid may serve as a beneficial embryo culture medium. Because the living chick embryo actively regulates the pH, osmolarity and O2 content of its own environment, it may serve as a more stable physiological system for mammalian embryo development than current in vitro culture systems (Blakewood et al., 1989a). In addition, the chick embryo actively synthesizes growth factors and other materials that may stimulate growth and development of mammalian embryos, a benefit unavailable in current embryo culture systems without supplementation of serum or bovine serum albumin (Blakewood et al., 1989a). The use of the chick embryo amnion for culturing bovine NT embryos may also be a way of circumventing potential risks associated with present bovine in vitro embryo culture conditions. In particular, the risk of transmitting BVDV or other pathogens into reconstructed embryos may be lowered when chick amniotic fluid is used for donor cell culture, as well as for both oocyte maturation and embryo culture. In the present study, 42.9% of the recipient females that received NT embryos co-cultured in the chick embryo amnion were pregnant on day 30 post estrus. This percentage was similar to contemporary day-30 pregnancy data for NT embryos cultured in vitro (in CR1aa medium for the first 72 hours and in CR1aa medium plus 5% FBS for the second 72 hours) using the same methods described herein (8/21 for 38.1%, unpublished data). All three recipient females lost their pregnancies prior to 120 days of gestation in the present study, while 37.5% (3/8) of the recipients carrying NT-derived fetuses remained pregnant in a corresponding study at this laboratory. It is widely acknowledged that NT in cattle results in increased rates of abortion throughout pregnancy, particularly during the first trimester. Since the timing of early fetal loss coincides with the timing of placentome formation in normal conceptuses, failure of placental development has been considered a contributing factor in pregnancy loss for NT embryos derived from embryonic cell lines (Stice et al., 1996). The effects appear
57 more extreme with somatic cell NT (Hill et al., 2000b) and may relate to a deficiency or a combination of deficiencies either in the NT procedure itself, leading to incomplete nuclear reprogramming of the cultured donor cells, or in the in vitro maturation and/or embryo culture systems used (Wells et al., 1999). The lack of placentome development could be the result of diminished or aberrant production of key regulatory hormones, such as insulin-like growth factors (IGFs), or abnormalities in maternal-placental cell communication (Hill et al., 2000a). It has been postulated that these anomalies, including fetal overgrowth, may reflect epigenetic changes, especially in the patterns of methylation, resulting from cloning and/or culture conditions (Niemann and Wrenzycki, 2000; Niemann et al., 2002; Young et al., 2001). Niemann and Wrenzycki (2000) have shown, in their studies of mRNA expression in viable in vitro-produced bovine embryos, that extended exposure to in vitro conditions leads to significant divergence from normal gene expression patterns. They suggested that alterations of gene expression during the pre-implantation period may affect fetal development of bovine embryos and could be relevant to the large offspring phenomenon. These alterations may be an attempt by the embryo to compensate for suboptimal conditions and further investigation of current in vitro embryo production systems is needed to determine the cause(s). One drawback of the chick embryo co-culture system is the embryo loss that occurs when the chick embryo dies. This death might contribute to a decrease in overall viable embryo yield, unless this co-culture system would produce embryos that were more viable at transfer and subsequently produce more live offspring than a standard culture system. In conclusion, results in the present study suggest that somatic cell NT bovine 8-cell embryos co-cultured in the chick embryo amnion are capable of developing into transferable stage embryos with a pregnancy rate comparable with embryos cultured in vitro in serum-supplemented medium. However, the problem of early pregnancy loss has not been ameliorated at this time. Several factors must be considered when determining the efficacy of this procedure for production of live offspring. First, embryos were only transferred when recipients were available, resulting in a low number of embryos transferred in this study. In addition, these transfers occurred at the end of May through July, 2002, a suboptimal time of year due to climate at this location. Further studies,
58 such as the use of the chick embryo co-culture system for both oocyte maturation and embryo culture, as well as transfer of additional embryos, must be conducted to determine the potential benefits of this culture system in bovine NT.
59 CHAPTER V
THE EFFECT OF ALTERATIONS IN THE NORMAL ANGIOGENIC PATTERN OF THE CHICK CHORIOALLANTOIC MEMBRANE ON BOVINE EMBRYO DEVELOPMENT AFTER CHICK EMBRYO CO-CULTURE
INTRODUCTION In vitro embryo production techniques are continuously being reinvestigated and reinvented (Wheeler et al., 2004). Despite much progress, however, in vitro-produced embryos still lag behind their in vivo-derived counterparts with respect to morphology (Greve et al., 1995), timing of development (Van Soom et al., 1997), embryonic metabolism (Khurana and Niemann, 2000) and pregnancy rates (Enright et al., 2000). In addition, a significant percentage of offspring resulting from embryos produced in vitro have been reported to exhibit distinct developmental abnormalities, known collectively as large calf or large offspring syndrome (Behboodi et al., 1995; Farin and Farin, 1995; Sinclair et al., 1995; Walker et al., 1996; Kruip and den Daas, 1997; McEvoy et al., 1998; van Wagtendonk-de Leeuw et al., 1998, 2000). Abnormalities associated with large offspring syndrome include increased birth weight (Behboodi et al., 1995; Sinclair et al., 1995; van Wagtendonk-de Leeuw et al., 1998, Jacobsen et al., 2000; van Wagtendonk- de Leeuw et al., 2000), hydroallantois (Hasler et al., 1995; van Wagtendonk-de Leeuw et al., 1998, 2000), placental abnormalities (Hasler et al., 1995; Farin and Farin, 1995) and perinatal and neonatal mortality (Behboodi et al., 1995; Schmidt et al., 1996). The inadequacies of current in vitro culture conditions have led researchers to develop alternative culture systems such as perifusion culture (Lim et al., 1996; McGowan and Thompson, 1997), microfluidic “microchannel” systems (Glasgow et al., 1998) and the “well-of-well” (WOW) method (Vajta et al., 2000) for culturing embryos in vitro. Previously, a culture system was developed in which mammalian embryos could be cultured in the amnion of a developing chick embryo. In early studies, researchers found that chick embryo co-culture resulted in significantly more expanded and hatched blastocysts compared with controls in mice (Blakewood et al., 1989a), goats (Blakewood et al., 1989a) and pigs (Ocampo et al., 1993). More recent research indicates that bovine embryos cultured in the chick amnion develop into blastocysts at rates similar to those of embryos cultured in a semi-defined medium (Davidson et al., unpublished data). It is believed that the chick amnion may serve as a more physiologically “normal” culture
60 environment compared with current “static” in vitro culture systems. Because of this the chick embryo culture system should not be abandoned without further investigation. Therefore the present study was conducted in an effort to improve the chick embryo co-culture system. It is hypothesized that by increasing blood flow to the chick embryo, chick embryo viability and growth factor production may be improved, thus potentially improving development of the bovine embryos contained within the amnion. Thus, the purpose of the first experiment was to determine if normal blood vessel patterns could be altered with angiogenic compounds. To accomplish this, angiogenesis was examined in chick chorioallantoic membranes (CAM) treated with prostaglandin E2
(PGE2) and prostaglandin F2α (PGF2α). Although many angiogenic compounds exist,
PGE2 and PGF2α were chosen for this experiment because they are similar in chemical composition, but have very opposite physiological actions. Specifically, PGE2 is a vasodialator involved in maintenance of the corpus luteum (CL) and implantation. PGF2α, in contrast, is a vasoconstrictor involved in regression of the CL and parturition. Based on the findings of the first experiment, a second experiment was conducted to determine if PGE2 treatment prior to or during bovine embryo co-culture could improve bovine embryo development over traditional culture techniques. MATERIALS AND METHODS Experimental Design The design for Experiment 5.1 is illustrated in Figure 5.1. Fertilized chicken eggs were obtained from a commercial supplier and incubated at 37°C and 85 to 90% relative humidity. On day 4 of incubation, eggs were prepared for treatment by creating a window in the egg shell. The window was sealed with transparent tape, and the eggs were returned to incubation for 24 hours. On day 5, eggs allotted to Treatment A (control) were treated with chick ringers solution, eggs allotted to Treatment B (EtOH) were treated with ethanol (vehicle), eggs allotted to Treatment C (cytochalasin) were treated with cytochalasin-D (negative control), eggs allotted to Treatment D (PGE) were treated with PGE2 and eggs allotted to Treatment E (PGF) were treated with PGF2α. All eggs were then returned to incubation. On day 6, CAMs were evaluated for treatment effects. The design for Experiment 5.2 is illustrated in Figure 5.2. Presumptive zygotes were purchased from a commercial supplier and upon arrival were placed in CR1aa for
61
EXPERIMENTAL DESIGN
Prepare eggs for assay Incubate eggs at 37°C by removing albumin and Apply treatments and 85 to 90% relative creating a window and return to Quantify humidity for 96 hours Return to incubation incubation results
0 1 2 3 4 5 6
Time (Days)
Figure 5.1. Experimental design to evaluate the effects of PGE2 and PGF2α on capillary development in the chick chorioallantoic membrane.
62 48 hours. On day 3 post-insemination, good quality 8-cell embryos were randomly allotted to one of three treatment groups. Embryos allotted to Treatment A (control) were cultured in CR1aa supplemented with 5% fetal bovine serum (FBS) from day 3 to day 7 post-insemination. Embryos allotted to Treatment B (CEC) were embedded in agar and injected into the amnion cavity of 96-hour chick embryos. After 96 hours of culture, the bovine embryos were recovered from the chick amnion and evaluated. Embryos allotted to Treatment C (PGE) were embedded in agar and injected into the amniotic cavity of
96-hour chick embryos that had been treated with PGE2. After 96 hours of culture, the bovine embryos were recovered from the chick amnion and evaluated. Experimental Methods Evaluation of the Effects of Prostaglandins on Angiogenesis Media Preparation Chick Lactated Ringers solution was prepared by addition of 123.2 mM NaCl (S-
5886, Sigma-Aldrich Inc., St. Louis, MO), 1.6 mM CaCl2·2H2O (C-7902 Sigma-Aldrich Inc., St. Louis, MO), 2.5 mM KCl (P-5404, Sigma-Aldrich Inc., St. Louis, MO), 10 mM HEPES (49329, Sigma-Aldrich Inc., St. Louis, MO) per 1 L of distilled water. The pH was then adjusted to 7.4. CAM Assay Angiogenesis was evaluated using the CAM assay modified from that previously described by Martins-Green and Feugate (1998). Fertilized chicken eggs were collected on the day of oviposition and stored at 10°C until incubation. Incubations were conducted at 37°C and 85 to 90% relative humidity throughout the duration of the experiment. On the fourth day of incubation, 3 to 4 ml of albumin was aspirated from the pole adjacent to the air pocket using an 18-gauge, 37.5-mm needle (Kendall, Mansfield, MA) and 3-ml disposable syringe (Sherwood, Davis and Geck, St. Louis, MO) and sealed with transparent tape (Gould QUIKSTIK; Gould Packaging, Inc. Vancouver, WA). This was done to allow the embryo to drop away from the egg shell and to allow the CAM to develop in a way that is accessible to treatments. A window (~25 mm in diameter) was opened in each egg to allow subsequent access to the CAM and then sealed with transparent tape. On day 5, 0.2 ml of chick ringers (control), 100% EtOH (vehicle), cytochalasin D (negative control; 0.99 µM; C-8273, Sigma-Aldrich Inc., St.
Louis, MO), PGE2 (2 µM; P-4172, Sigma-Aldrich Inc., St. Louis, MO) or PGF2α (2 µM;
63 Control Group
Shipment of Presumptive Media Change Zygotes IVC IVF Addition of FBS Evaluation
18 h ~24 h 48 h 96 h
CEC Treatment Group
Shipment of Onset of Presumptive Preparation of Injection of Recovery Eg g Incubation IVF Zygotes IVC Shell-less Cultures Bovine Embryos Evaluation
18 h ~24 h 24 h 24 h 96 h
Onset of IVF Shipment of IVC Preparation of Injection of Recovery Egg Incubation Presumptive Shell-less Cultures Bovine Embryos Evaluation Zygotes Application of PGE2
PGE Treatment Group
Figure 5.2. Experimental design for the culture of 8-cell bovine embryos cultured in CR1aa + 5% fetal bovine serum (control), co- cultured in the chick embryo amnion (CEC) or co-cultured in the amnion of PGE2-treated chicks (PGE).
64 P-0424, Sigma-Aldrich Inc., St. Louis, MO) were applied to the surface of each CAM by inserting a 22-gauge, 37.5-mm needle (Kendall, Mansfield, MO) through the transparent tape and delivering treatment solutions in a back and forth motion to ensure coverage of the entire CAM. On day 6, CAMs were prepared for evaluation as described below. Quantification of CAM Results Control and experimental CAMs were prepared for quantification by removing an additional 3 to 4 ml of albumin as described above and enlarging the window to expose as much of the CAM as possible. Digital images of each CAM were taken with an Olympus C-2100 digital camera from a distance of 31.3 cm with the addition of close-up lenses with diopters of +1, +2 and +4 for macro-imaging. All digital images were processed using Adobe Photoshop (v 6.0). Degree of capillary branching was quantified by placing a 256 pixel x 256 pixel box on each CAM image at random. The chosen area was then pasted into a new blank image and processed using the brightness/contrast option to contrast enhance the blood vessels. The nodes or branch points were then counted. This process was repeated three additional times to obtain node counts for a total of four random areas of each CAM. Data from the four boxes were combined to produce a mean number of nodes for each CAM. Production of In Vitro Embryos Media Preparation Brackett-Oliphant (BO) media were prepared according to Brackett and Oliphant (1975). Briefly, BO-A stock solutions were prepared by addition of 150.2 mM NaCL (S- 5886, Sigma-Aldrich Inc., St. Louis, MO), 2.7 mM KCl (P-5405, Sigma-Aldrich Inc., St.
Louis, MO), 3.0 mM CaCl2·2H2O (C-7902, Sigma-Aldrich Inc., St. Louis, MO), 1.0 mM
NaH2PO4·H2O (S-9638, Sigma-Aldrich Inc., St. Louis, MO), 0.6857 mM MgCl2·6H2O (M- 2393, Sigma-Aldrich Inc., St. Louis, MO) and 0.1 ml of 0.5% phenol red (P-0290, Sigma- Aldrich Inc., St. Louis, MO) per 500 ml of sterile water. BO-B stock solutions were prepared by addition of 154.0 mM NaHCO3 (S-8875, Sigma-Aldrich Inc., St. Louis, MO) and 0.04 ml of 0.5% phenol red per 200 ml of sterile water. Fertilization stock medium (BO-AB) was prepared from the stock solutions by adding 1.2 mM pyruvic acid, 0.05 ml of gentamicin (50 mg/ml; 15750-060, Gibco, Grand Island, NY) and 0.018 ml of heparin sodium (10,000 IU/ml; Elkins-Sinn, Cherry Hill, NJ)
65 per 38 ml of BO-A stock solution and 12 ml of BO-B stock solution. Fertilization medium (BO-caffeine) was prepared by adding 0.0243 g of caffeine sodium (C-4144, Sigma- Aldrich Inc., St. Louis, MO) per 25 ml of BO-AB. Sperm washing medium (0.6% BSA- BO) was prepared by adding 0.03 g of bovine serum albumin (BSA; Fraction V, A-4503, Sigma-Aldrich Inc., St. Louis, MO) to 5 ml of BO-AB. Oocyte washing medium (0.3% BSA-BO) was prepared by adding 0.03 g of BSA (Fraction V) to 10 ml of BO-AB. TCM oocyte washing medium was prepared by adding 0.03 g of BSA (Fraction V) to 10 ml of Tissue Culture Medium-199 (Eagle’s salt) with 25 mM HEPES Buffer (TCM-199; Gibco Laboratories, Grand Island, NY). CR1aa medium was prepared according to Rosenkrans and First (1994). CR1aa stock solution was first prepared by adding 114.7 mM NaCl, 1.6 mM KCl, 26.2 mM
NaHCO3, 0.3999 mM pyruvic acid (P-4562, Sigma-Aldrich Inc., St. Louis, MO), 2.5 mM L(+) lactic acid (L-4388, Sigma-Aldrich Inc., St. Louis, MO), 0.5195 mM glycine (G-8790, Sigma-Aldrich Inc., St. Louis, MO), 0.5051 mM L-alanine (A-7469, Sigma-Aldrich Inc., St. Louis, MO) and 0.2 ml of 0.5% phenol red per 100 ml of sterile water. For embryo culture from day 0 to day 3, CR1aa culture medium was prepared by adding 0.2 ml of Basal Medium Eagle (BME) amino acid solution (50X, B-6766, Sigma- Aldrich Inc., St. Louis, MO), 0.1 ml of Modified Eagle Medium (MEM) amino acid solution (10X, 11140-050, Gibco Laboratories, Grand Island, NY), 0.01 ml of gentamicin, 0.00146 g of L-glutamine (G-5763, Sigma-Aldrich Inc., St. Louis, MO) and 0.03 g of BSA (fatty acid free, A-7511, Sigma-Aldrich Inc., St. Louis, MO) to 9.5 ml of CR1aa stock solution. For embryo culture from day 3 to day 7, CR1aa culture medium was prepared by adding 0.2 ml of BME amino acid solution, 0.1 ml of MEM amino acid solution, 0.5 ml of fetal bovine serum (FBS; SH30070.02, HyClone, Logan, UT), 0.01 ml of gentamicin, 0.00146 g of L-glutamine and 0.03 g of BSA (fatty acid free) to 9.5 ml of CR1aa stock solution. In Vitro Fertilization Oocytes were purchased weekly from a commercial supplier (BoMed, Inc, Madison, WI) and shipped in a portable incubator (~39°C). Upon arrival at the laboratory, shipping vials were removed from the portable incubator and stored in a humidified atmosphere of 5% CO2 in air at 39°C for completion of maturation (maturation time = 20 to 22 hours).
66 To prepare the semen for in vitro fertilization, a straw of frozen semen from a Holstein bull (CSS 7H5188, Genex Cooperative Inc., Shawano, WI) of proven fertility was thawed in a 39° to 40°C water bath for ~45 seconds. After thawing, the contents of the straw were deposited into a 15-ml plastic conical tube (Corning, New York, NY). Then, 9 ml of BO-caffeine was added to the tube and the suspension was centrifuged for 6 minutes at 200 x g. After centrifugation the supernatant was removed and the remaining pellet was resuspended in an additional 9 ml of BO-caffeine. The resulting suspension was centrifuged for 6 additional minutes at 200 x g. The supernatant was removed again and the pellet resuspended in 4 ml of BO-caffeine and 4 ml of 0.6% BSA- BO. The semen preparation was then allowed to equilibrate in a 39°C incubator for ~10 minutes. After equilibration, four 80-µl insemination drops were created in a preheated 35 x 10-mm Falcon® plastic Petri dish (Becton and Dickinson, Lincoln Park, NJ) and covered with warmed mineral oil (M-8140, Sigma-Aldrich Inc., St. Louis, MO). After 20 to 22 hours of maturation, cumulus oocyte complexes were removed from the shipping vials and washed through two 35 x 10-mm plastic petri dishes containing 2 to 3 ml of 0.3% BSA-BO. Then, 10 to 20 oocytes were placed in each of the 80-µl insemination drops previously prepared and were co-incubated for 5 hours at 39°C and 5% CO2 in air. After the 5-hour fertilization period, oocytes were removed from insemination drops and washed through a single 35 x 10-mm plastic petri dish of TCM oocyte washing medium. Cumulus cells were then removed by vortexing for 3 minutes in 1 ml of TCM-199 containing 0.001 g of hyaluronidase (H-3506, Sigma-Aldrich Inc., St. Louis, MO). After vortexing, oocytes were washed through two 35 x 10 mm plastic petri dishes containing TCM oocyte washing medium and one 35 x 10-mm plastic petri dish of CR1aa. Then, 10 to 15 oocytes per drop were placed in 50-µl drops of CR1aa covered with warmed mineral oil and incubated in a modular incubator containing 5% CO2, 5%
O2 and 90% N2 at 39°C. Chick Embryo Co-Culture The procedure for the culture of bovine 8-cell IVF embryos using the chick embryo amnion was modified from that of Blakewood and Godke (1989). Briefly, fertilized domestic chicken eggs were collected on the day of oviposition and stored at 10°C until the onset of incubation. Four days prior to the introduction of bovine embryos,
67 the eggs were removed from cold storage and allowed to come to room temperature (25°C) before placing them in a 37°C commercial egg incubator. After 72 hours of incubation, the eggs were sterilized with 70% ethanol (Aaper Alcohol and Chemical Company, Shelbyville, KY) and 7.5% providone iodine (Betadine®, Purdue Fredrick Company, Stanford, CT). The eggs were then allowed to air dry in a horizontal position to ensure proper positioning of the chick embryo. To remove the egg contents, the egg was broken against the rim of a sterile beaker and the contents released into a 100-ml plastic embryo collection dish (Veterinary Concepts, Spring Valley, WI) covered by a piece of cellophane kitchen wrap (Saran®, Dow Chemical, Midland, MI). The dish, including the egg contents, was then loosely capped with a plastic lid (Veterinary Concepts, Spring Valley, WI) and placed in a 38°C incubator for an additional 24 hours before introducing the 8-cell bovine embryos into the amniotic cavity of the chick embryo. Prior to embedding and injection of bovine embryos, suitable chick embryos were selected for injection based on the following criteria: 1) round intact yolk, 2) good vascularity in the chorioallantoic membrane, 3) vigorous heartbeats and 4) no apparent abnormalities. Bovine embryos were embedded in agarose according to the procedure set forth by Willadsen (1979) for the culture of ovine embryos in the ligated oviducts of sheep. On day 3 of culture, two or three 8-cell bovine IVF embryos were placed in a 50- µl drop of low-melting agarose solution [1.2% agarose (A-9799, Sigma-Aldrich Inc., St. Louis, MO) dissolved in Dulbecco’s Phosphate-Buffered Saline (DPBS; D1408, Sigma- Aldrich Inc., St. Louis, MO)] at ~37oC. The embryos were then aspirated into a beveled glass pipette (200 µm outer diameter) and held in air for ~30 seconds to allow the agarose to harden. Then, the agar (containing the embryos) within the pipette was expelled as a gel cylinder into room temperature TL-HEPES (BioWittaker, Walkersville, MD). The agarose cylinder, including the bovine embryos, was then aspirated back into the beveled pipette, and carefully expelled into the amniotic cavity of a 96-hour chick embryo. After introducing the embedded embryos into the amniotic cavity the shell-less culture systems were returned to incubation for an additional 72 hours at 39oC. Thus, a single agarose cylinder containing two or three 8-cell bovine IVF embryos was co- cultured in the amnion of each developing chick embryo for 4 days.
68 On day 7 of in vitro culture, the amnion containing the agarose cylinder was dissected away from the rest of the egg components and placed in a 35 x 10-mm plastic petri dish. The amnion was then rinsed several times with DPBS containing 1% FBS. After rinsing, the petri dish was placed under a stereomicroscope (Nikon SMZ-2B, Tokyo, Japan) equipped with an overhead light source and the agarose cylinder was removed from the amniotic cavity using two 22-gauge, 37.5-mm needles (Kendall, Mansfield, MA) to make a small opening in the amniotic membrane. The agarose cylinder was carried from the amnion with the escaping amniotic fluid, settling on the bottom of the dish. The cylinder was then washed twice in CR1aa medium and dissected using two 22-gauge needles to isolate and remove the developing embryos. Statistical Analysis For Experiment 5.1, data were analyzed using PROC GLM in the SAS Statistical Package (v. 8e, 1999, SAS Institute, Inc., Gary, NC). When significance was detected (P<0.05), post-hoc comparisons were made using Duncan’s Multiple Range test. For Experiment 5.2, data were analyzed using PROC GLM of SAS. Significant differences between embryo development rates in each group were tested using a post hoc least significant difference test (LSD test). Differences with a probability (P) value of 0.05 or less were considered significant in this study RESULTS In Experiment 5.1, chick embryo mortality was 41%, with the highest mortality rate (86%) occurring in CAMs treated with cytochalasin D (Table 5.1). To determine the affect of prostaglandins on blood vessel development, CAMs were exposed to treatments at an early stage of blood vessel development (day 5) and then evaluated 24 hours later. Average node counts per CAM are provided in Table 5.2. Capillary branching was not affected (P>0.05) when CAMs were treated with cytochalasin D (negative control), but was significantly greater (P<0.05) when CAMs were treated with
EtOH (carrier), PGE2 or PGF2α (Figure 5.3). Both PGE2 and PGF2α stimulated (P<0.05) capillary branching when compared with control and cytochalasin-treated CAMs, and
PGE2 increased (P<0.05) capillary branching over that of EtOH. PGF2α did not differ
(P>0.05) from either EtOH or PGE2 (Figure 5.4). In Experiment 5.2, 60 chicken eggs were initially placed in incubation. After 72 hours of incubation, the eggs were removed from their shell and placed in shell-less
69
Table 5.1. Mortality of chick embryos after treatment application and subsequent 24 hour incubation
Treatment n No. viable chick embryos (%) No. dead chick embryos (%)
Control 6 3 (50) 3 (50)
Cytochalasin D 7 1 (14) 6 (86)
Ethanol 7 5 (71) 2 (29)
a PGE2 7 4 (57) 3 (43)
PGF2α 7 7 (100) 0 (0)
a One CAM was damaged while enlarging the shell for quantification of capillary development.
70
Table 5.2. Effect of cytochalasin D, ethanol, PGE2 and PGF2α on capillary branching in the chorioallantoic membrane of the chick embryo. Treatment effects were quantified by counting the number of nodes in four random areas of each CAM
Treatment n Average no. nodes per CAM ±SD Control 20 8.4a 1.28
Cytochalasin D 4 10.5a 1.04
Ethanol 20 16.3b 1.89
c PGE2 28 24.1 1.26
bc PGF2α 16 19.1 1.22 a,b,c Values without a common superscript are significantly different (P<0.05).
71
A B
C D
Figure 5.3. Restructuring of vasculature networks in CAMs treated with (A) chick Ringer’s solution (control), (B) ethanol (vehicle), (C) PGE2 and (D) PGF2α.
72
30
c 25
bc 20 b
15
a
10 a Number of Nodes
5
n = 12 n = 4 n = 20 n = 28 n = 16 0 Control Cytocha lasin EtOH PGE PGF
Figure 5.4. Effect of cytochalasin D, ethanol (EtOH), PGE2 (PGE) and PGF2α (PGF) on capillary branching in the chorio- allantoic membrane of the chick embryo. The number on each bar indicates the total number of samples (four samples per CAM) in each treatment group. a,b,c Columns without a common superscript are significantly different (P<0.05).
73 culture. At this time, four (6%) of the eggs were found to be unfertilized, four (6%) contained embryos that failed to fully develop and one (2%) contained an embryo that was nonviable. The remaining 51 eggs were returned to incubation. After 24 hours of incubation in shell-less culture, 34 eggs were selected for injection of bovine embryos. For chick embryo co-culture, 2-cell bovine embryos (n=102) were embedded and placed into the amniotic cavities of chick embryos, and 51 (50%) were successfully recovered on day 7 for evaluation. Loss of embryos occurred during recovery or was due to chick embryo mortality. Death of chick embryos was the primary reason for the loss, as six of the 17 (35%) chick embryos in the CEC treatment group, and four of the 17 (24%) chick embryos in the PGE2 treatment group did not survive injection and subsequent shell-less culture. On day 7 of culture, a significantly greater (P<0.05) percentage of morulae were noted following culture in PGE2-treated chick embryos than in untreated chicks. There was no significant difference (P<0.05) between the CEC treatment group and the control treatment group (Table 5.3). Although there was no difference detected in the number of early blastocysts or blastocysts among the three treatments, there were significantly more expanded blastocysts after culture in control medium alone than in either chick embryo co-culture treatment group (Figure 5.5). The number of hatching blastocysts did not differ (P>0.05) across treatments (Figure 5.5). Although there were no differences in the percentage of embryos developing to the early blastocyst, hatching blastocysts or hatched blastocyst stage on day 8 of development, there were significantly more expanded blastocysts in the control treatment group than the CEC treatment group.
Embryo development to the expanded blastocyst stage in the PGE2 treatment group was not significantly different (P>0.05) from the other two groups. On day 9 of development, there were more blastocysts in the CEC and PGE2 treatment groups compared with controls (P<0.05). However, embryo development to the expanded blastocyst, hatching blastocyst or hatched blastocyst stage did not differ with treatment (P>0.05). Embryo quality grade did not differ (P>0.05) between the three treatments on day 7 or day 8 of culture, but there were significantly more (P<0.05) grade 1 blastocysts in the CEC treatment group than in the PGE2 or control treatment groups on day 9 of development (Figure 5.6).
74
Table 5.3. Development of 8-cell bovine embryos cultured in CR1aa + 5% fetal bovine serum (control), co-cultured in the chick embryo amnion or co-cultured in the amnion of PGE2-treated chicks
a Embryos allotted to the control group were cultured in CR1aa + 5% FBS from day 3 to day 7 post-insemination. Embryos allotted to the CEC group were culture in the amnion of a developing chick embryo from day 3 to day 7 post insemination.
Embryos cultured in the PGE2 group were cultured in the amnion of PGE2 treated chicks from day 3 to day 7 post insemination. b The number of 8-cell embryos allotted to each treatment. The number in parenthesis indicates the number of embryos successfully recovered from live chick embryos. c The percentage of embryos developing in the chick co-culture group and the PGE2 treated chick co-culture group were based on the number of embryos successfully recovered from live chick embryos. d MOR = morulae. e BLST = blastocysts. f HT/HD BLST = hatching and/or hatched blastocysts. g,h Within a column, values without a common superscript are significantly different (P<0.05).
75 A Control 40 b CEC a PGE
s 30 ab
o a
y a r b
m 20 a
E a aa f a % o 10 b b a a a 0 MORL Early BLST Expand Hatching BLST BLST BLST
B Control 40 CEC PGE
s 30
o a y a r b
m 20 a a a E
f a a a a ab % o 10 a a b a a a a a 0 MORL Early BLST Expanded Hatching Hatche d BLST BLST BLST BLST
C Control CEC 20 b b PGE 15 s o y
br a a
m 10 E a of a a % 5 a
a a a a 0 BLST Expand Hatching Hatche d BLST BLST BLST Figure 5.5. Development of 8-cell bovine embryos cultured in CR1aa supplemented with 5% fetal bovine serum (control), cultured in the chick embryo amnion (CEC) or cultured in the amnion of PGE2-treated chicks (PGE) on day 7 of development (A), day 8 of development (B) and day 9 of development (C). Within a developmental stage, columns without a common superscript are significantly different (P<0.05).
76
A Control 50 CEC PGE a a 40 a a s o
y 30
br a m E 20 a
% of a
10 a a a a a 0 Grade 1 Grade 2 Grade 3 Grade 4
B Control 50 CEC PGE
a 40 a s o
y 30 a br
m E 20 a of a a % 10 a a a a a a 0 Grade 1 Grade 2 Grade 3 Grade 4
C Control CEC 50 PGE b 40
s
o a y 30
br m E 20 a of
% a a 10 a a a a a a a 0 Grade 1 Grade 2 Grade 3 Grade 4 Figure 5.6. Quality grade of bovine embryos cultured in CR1aa supplemented with 5% fetal bovine serum (control), cultured in the chick embryo amnion (CEC) or cultured in the amnion of PGE2-treated chicks (PGE) on day 7 of development (A), day 8 of development (B) and day 9 of development (C). Within a quality grade, columns without a common superscript are significantly different (P<0.05).
77 DISCUSSION Angiogenesis is the formation of new blood vessels by branching and growth, and migration of pre-existing blood vessels (Rissau, 1995; Rissau and Flamme, 1995; Rissau, 1997). This capillary proliferation is vital during embryogenesis (Wilting et al., 1995; Flamme et al., 1997), wound healing (Arbiser, 1996; Martins-Green and Hanafusa, 1997) and bone fracture repair (Klagsburn and D’Amore, 1991). Rapid or persistent capillary growth also has been associated with tumors, retinopathies, haemangiomas, fibrosis and rheumatoid arthritis (for review see Folkman and Klagsburn, 1987; Klagsburn and D’Amore, 1991). In most adult tissues, angiogenesis is rare (Denekamp, 1984; Klagsbrun and D’Amore, 1991). Exceptions can be found in the reproductive organs of adult females, where angiogenesis occurs cyclicly in the ovary and the uterus, as well as in the uterine endometrium and placenta during pregnancy (Augustin et al., 1995; Reynolds and Redmer, 1995; Redmer and Reynolds, 1996). The purpose of the first experiment of this series was to determine if normal blood flow patterns to the chick embryo could be altered by the application of prostaglandins. To investigate this hypothesis, angiogenesis was examined in chick chorioallantoic membranes (CAM) treated with either PGE2 or PGF2α. The CAM forms between day 4 and day 5 of development by fusion of the allantois and chorion. The resulting membrane consists of the chorion (ectoderm), mesoderm with blood vessels and allantois (endoderm) (DeFouw et al., 1989). The CAM is of physiologic importance to the chick because it serves as the major respiratory organ for gaseous exchange until hatching, and it provides a bladder into which waste products can be delivered (Romanoff, 1967). Because the chorioallantoic membrane is easily accessible for in vivo work and is highly vascularized, it has proven to be an excellent model for the study of the angiogenic process (Ribatti et al., 1996).
Findings of the first experiment suggest that PGF2α may directly inhibit angiogenesis. Based on visual observation, blood vessels in CAMs treated with PGF2α appeared thicker and larger with fewer main vessels than those in controls or vehicle- treated CAMs. The response of CAMs to PGF2α was not dramatic, however, and quantification of branch nodes was not significantly different than vehicle. It is interesting to note that although blood vessel development was visually impaired in PGF2α -treated chicks, 100% of the treated chicks survived shell-less culture. These findings are
78 counterintuitive to our original hypothesis, but the small number of observations should be taken into account. It is possible that these findings would not hold true with additional observations.
In contrast, CAMs treated with PGE2 showed a significant increase in capillary development compared with controls and vehicle-treated CAMs, suggesting that PGE2 directly influences angiogenesis. It is well known that increased blood flow to a tissue results in an increase of function (Guyton, 1971). Should PGE2 in fact increase angiogenesis, as indicated here, then it is possible to infer that the increase in blood vessel formation would increase blood flow to the chick embryo, resulting in increased chick viability. The results of the first study were encouraging because the inherent mortality rate of chick embryos maintained in shell-less culture poses a major drawback to the efficacy of this procedure for use as an alternative embryo culture technique. Dunn and Boone (1976) reported that chick survival rates of 90 to 95% appear as high as can be expected with shell-less culture. More importantly, Rowlett and Simkiss (1987) have reported losses as high as 27% between day 4 and day 7 of shell-less culture, the time period in which bovine embryos are maintained in the chick amnion. If chick viability could be improved with the application of prostaglandins, loss of valuable bovine embryos upon chick death could be minimized.
In the second experiment, CAMs were treated with PGE2 24 hours before introduction of bovine embryos into the amniotic cavity. In this study, there was only a slight improvement in chick embryo viability in PGE2 treated chicks (65% in CEC and
76% in PGE2); although, as in the first study, an obvious increase in capillary development in PGE2-treated chicks compared with non treated chicks was observed. Unfortunately, increasing blood flow to the chick embryo did not appear to improve development or quality of bovine embryos cultured in untreated chicks or in semi-defined medium (control). These findings suggest that the benefits of the chick embryo culture system may not be due to growth factors and other embryotropic properties within the amniotic fluid, but rather in the ability of the chick embryo to actively regulate the pH, osmolarity and O2 content of its own environment.
79 CHAPTER VI
DEVELOPMENT OF 8-CELL BOVINE EMBRYOS IN CULTURE MEDIUM SUPPLEMENTED WITH CHICK AMNIOTIC FLUID (CAF)
INTRODUCTION Even with continual efforts worldwide, researchers have been unable to fully replicate the conditions within the oviduct and uterus in vitro. The development of immature bovine oocytes to the blastocyst stage appear to have plateaued at 30 to 40% following in vitro maturation, fertilization and culture (Krisher and Bavister, 1998; Lonergan et al., 2003). In addition, the quality of blastocysts produced in vitro rarely, if ever, reach that of in vivo-derived blastocysts (Duby et al., 1997; Boni et al., 1999; Viuff et al., 1999) and marked differences have been reported to exist between in vivo- and in vitro-derived embryos with regard to morphology (Greve et al., 1995), timing of development (Van Soom et al., 1997), embryonic metabolism (Khurana and Niemann, 2000) and pregnancy rates (Enright et al. 2000). Current culture conditions rely heavily on the addition of fetal bovine serum (FBS) and/or bovine serum albumin (BSA) to the culture medium (Bavister, 1995), despite the undefined and variable nature of serum composition (Rizos et al., 2003). It has been suggested that serum may contain components advantageous to blastoceol development, such as growth factors or extracellular matrix components, like fibronectin (Pinyopunnintr and Bavister, 1994). However, it has also been suggested that the components of serum may contribute to fetal and calf abnormalities (Thompson et al., 1995; Walker et al., 1996; Sinclair et al., 1999). Specifically, offspring produced from embryos cultured in vitro often exhibit increased birth weight (Willadsen et al., 1991; Keefer et al, 1994; Behboodi et al., 1995), longer gestation lengths (Walker et al., 1996), increased placental anomalies, (Hasler, et al., 1995; Hill et al., 1999, 2001, 2002) and increased incidences of abortion (Walker et al., 1996; Hill et al., 1999) and perinatal (Behboodi et al., 1995; Wilson et al., 1995; Garry et al., 1996; Schmidt et al., 1996; Kruip and den Daas, 1997; Hill et al., 1999) and neonatal mortality (Schmidt et al., 1996; Kruip and den Daas, 1997; Hill et al., 1999). Attempts to circumvent potential risks associated with the addition of serum to embryo culture media have included culturing embryos in the amniotic cavity of a developing chicken embryo. Previous reports have indicated that the chick amnion is
80 capable of supporting the development of murine (Blakewood et al., 1989a), caprine (Blakewood et al., 1989a), bovine (Blakewood et al., 1989b; Murakami et al., in press) and porcine (Ocampo et al., 1993) embryos, and live transplant offspring have been produced following chick embryo co-culture of 2- to 8-cell caprine embryos (Blakewood et al., 1990). However, the inherent mortality rate of shell-less chick embryos poses a major drawback to the efficacy of this procedure. Dunn and Boone (1976) reported that chick survival rates of 90 to 95% appear to be as high as can be expected with shell-less culture, and a rapid degeneration of mammalian embryos has been reported following chick embryo death (Blakewood and Godke, 1990). Therefore, the objective of this study was to determine if amniotic fluid extracted from a developing chick embryo could be used as a supplement to standard in vitro culture media and support development of bovine embryos. MATERIALS AND METHODS Experimental Design The design for this experiment is illustrated in Figure 6.1. Upon removal from in vitro insemination, all presumptive zygotes were placed in 50-µl drops of CR1aa (10 to 15 per 50-µl drop) for in vitro culture (Rosenkrans and First, 1994). After 72 hours of culture, good quality (as indicated by equal size, well defined blastomeres) 6- to 8-cell embryos were randomly allotted to one of two treatment groups. Embryos allotted to Treatment A (control group) were cultured in CR1aa medium supplemented with 5% FBS. Embryos allotted to Treatment B (CAF group) were cultured in CR1aa medium supplemented with 20% CAF obtained from a day 7 chick embryo. All embryos were cultured in a modular incubator containing 5% CO2, 5% O2 and 90% N2 at 39°C. Media were changed on day 7 of culture and embryonic development was assessed on days 7, 8 and 9. At each evaluation time point, each embryo was assigned a quality grade score from 1 to 4 (grade 1 = excellent quality to grade 4 = poor quality or degenerate). Experimental Methods Media Preparation For the preparation of Brackett-Oliphant (BO) media, BO-A stock solutions of 150.2 mM NaCL (S-5886, Sigma-Aldrich Inc., St. Louis, MO), 2.7 mM KCl (P-5405,
Sigma-Aldrich Inc. St. Louis, MO), 3.0 mM CaCl2·2H2O (C-7902, Sigma-Aldrich Inc., St.
81
In Vitro Maturation
18 to 22 hours
In Vitro
Fertilization
5 hours
In Vitro Culture
72 hours
Control CAF
96 hours
Media Change Embryo Evaluation
24 hours
Embryo Evaluation
24 hours
Embryo Evaluation
Figure 6.1. Experimental design for culturing 8-cell bovine embryos in culture medium supplemented with chick amniotic fluid. In vitro culture was either in CR1aa supple- mented with 5% fetal bovine serum (control group) or CR1aa supplemented with 20% chick amniotic fluid (CAF group).
82 Louis, MO), 1.0 mM NaH2PO4·H2O (S-9638, Sigma-Aldrich Inc., St. Louis, MO), 0.6857 mM MgCl2·6H2O (M-2393, Sigma-Aldrich Inc., St. Louis, MO) and 0.1 ml of 0.5% phenol red (P-0290, Sigma-Aldrich Inc., St. Louis, MO) per 500 ml of sterile water were prepared. BO-B stock solutions of 154.0 mM NaHCO3 (S-8875, Sigma-Aldrich Inc., St. Louis, MO) and 0.04 ml of 0.5% phenol red per 200 ml of sterile water were also prepared (Brackett and Oliphant, 1975). For the preparation of CR1aa culture medium, a CR1aa stock solution of 114.7 mM NaCl, 1.6 mM KCl, 26.2 mM NaHCO3, 0.3999 mM pyruvic acid (P-4562, Sigma- Aldrich Inc., St. Louis, MO), 2.5 mM L(+) lactic acid (L-4388, Sigma-Aldrich Inc., St. Louis, MO), 0.5195 mM glycine (G-8790, Sigma-Aldrich Inc., St. Louis, MO), 0.5051 mM L-alanine (A-7469, Sigma-Aldrich Inc., St. Louis, MO) and 0.2 ml of 0.5% phenol red per 100 ml of sterile water was prepared (Rosenkrans and First, 1994). From the BO-A and BO-B stock solutions, fertilization stock medium (BO-AB) was prepared by adding 1.2 mM pyruvic acid, 0.05 ml of gentamicin (50 mg/ml; 15750- 060, Gibco, Grand Island, NY) and 0.018 ml of heparin sodium (10,000 IU/ml; Elkins- Sinn, Cherry Hill, NJ) per 38 ml of BO-A stock solution and 12 ml of BO-B stock solution. Fertilization medium (BO-caffeine) was prepared by adding 0.0243 g of caffeine sodium (C-4144, Sigma-Aldrich Inc., St. Louis, MO) per 25 ml of BO-AB. Sperm washing medium (0.6% BSA-BO) was prepared by adding 0.03 g of BSA (Fraction V, A-4503, Sigma-Aldrich Inc., St. Louis, MO) to 5 ml of BO-AB. Oocyte washing medium (0.3% BSA-BO) was prepared by adding 0.03 g of BSA (Fraction V) to 10 ml of BO-AB. TCM oocyte washing medium was prepared by adding 0.03 g of BSA (Fraction V) to 10 ml of Tissue Culture Medium-199 (Eagle’s salt) with 25 mM HEPES Buffer (TCM-199; Gibco Laboratories, Grand Island, NY). From the CR1aa stock solutions, CR1aa culture medium for day 0 to day 3 was prepared by adding 0.2 ml of Basal Medium Eagle (BME) amino acid solution (50X, B- 6766, Sigma-Aldrich Inc., St. Louis, MO), 0.1 ml of Modified Eagle Medium (MEM) amino acid solution (10X, 11140-050, Gibco Laboratories, Grand Island, NY), 0.01 ml of gentamicin, 1.1 mM L-glutamine (G-5763, Sigma-Aldrich Inc., St. Louis, MO) and 0.03 g of BSA (fatty acid free, A-7511, Sigma-Aldrich Inc., St. Louis, MO) to 9.5 ml of CR1aa stock solution. CR1aa culture medium for day 3 to day 7 was prepared by adding 0.2 ml of BME amino acid solution, 0.1 ml of MEM amino acid solution, 0.5 ml of FBS
83 (SH30070.02; HyClone, Logan, UT), 0.01 ml of gentamicin, 1.1 mM L-glutamine and 0.03 g of BSA (fatty acid free) to 9.5 ml of CR1aa stock solution. In Vitro Fertilization Oocytes were purchased weekly from a commercial supplier (BoMed, Inc, Madison, WI) and matured in a portable incubator (~39°C) during the overnight travel interval (20 to 22 hours). Upon arrival at the laboratory, shipping vials were removed from the portable incubator and stored in a humidified atmosphere of 5% CO2 in air at 39°C until insemination. To prepare semen for fertilization, a straw of frozen semen from a Holstein bull (CSS 7H5188, Genex Cooperative Inc., Shawano, WI) of proven fertility was thawed in a 39° to 40°C water bath for ~45 seconds. After thawing, the contents of the straw were deposited into a 15-ml plastic conical tube (Corning, New York, NY) and 9 ml of BO- caffeine was added to the tube. The suspension was then centrifuged for 6 minutes at 200 x g. After centrifugation, the washing step was repeated by resuspending the semen pellet in an additional 9 ml of BO-caffeine and centrifuging for 6 additional minutes at 200 x g. The supernatant was removed and the pellet resuspended in 4 ml of BO- caffeine and 4 ml of 0.6% BSA-BO. The semen preparation was then placed in a 39°C incubator for ~10 minutes to equilibrate. After equilibration, four 80-µl insemination drops were created in a preheated 35 x 10-mm Falcon® plastic petri dish (Becton and Dickinson, Lincoln Park, NJ) and covered with warmed mineral oil (M-8140, Sigma- Aldrich Inc., St. Louis, MO). Following 20 to 22 hours of maturation, cumulus oocyte complexes were removed from the shipping vials and washed through two 35 x 10-mm plastic petri dishes containing 2 to 3 ml of 0.3% BSA-BO. Then, 10 to 20 oocytes were placed in each of the 80-µl insemination drops previously prepared and were co-incubated for 5 hours at 39°C and 5% CO2 in air. After the 5-hour fertilization period, oocytes were washed through a single 35 x 10-mm plastic petri dish of TCM oocyte washing medium and the cumulus cells were removed by vortexing for 3 minutes in 1 ml of TCM-199 containing 0.001 g of hyaluronidase (H-3506, Sigma-Aldrich Inc., St. Louis, MO). The oocytes were then washed through two 35 x 10-mm plastic petri dishes containing TCM oocyte washing medium and one 35 x 10-mm plastic petri dish of CR1aa. Then, 10 to 15 oocytes per
84 drop were placed in 50-µl drops of CR1aa covered with warmed mineral oil and incubated in a modular incubator containing 5% CO2, 5% O2 and 90% N2 at 39°C. Collection of Chick Amniotic Fluid (CAF) The basic procedure used for the shell-less culture of chick embryos was modified from that previously described by Blakewood and Godke (1989). Briefly, fertilized domestic chicken eggs were collected on the day of oviposition and stored at 10°C until the onset of incubation. Seven days prior to the day of amniotic fluid collection the eggs were removed from cold storage and allowed to come to room temperature (25°C) before placing them in a 37°C commercial egg incubator. After 72 hours of incubation, the eggs were rinsed with a solution of 70% ethanol (Aaper Alcohol and Chemical Company, Shelbyville, KY) and then coated with 7.5% providone iodine (Betadine®, Purdue Fredrick Company, Stanford, CT) and allowed to air dry in a horizontal position to ensure proper positioning of the chick embryo. To remove the egg contents, a crack was made in the shell by striking the center of the egg against the rim of a sterile beaker. The egg contents were then released into a 100-ml plastic embryo collection dish (Veterinary Concepts, Spring Valley, WI) covered by a piece of cellophane kitchen wrap (Saran®, Dow Chemical, Midland, MI). The dish, including the egg contents, was loosely capped with a plastic lid (Veterinary Concepts, Spring Valley, WI) and maintained in an incubator at 39°C for an additional 96 hours before collection of the amniotic fluid. On day 7 of shell-less culture, fluid was aspirated from the amnion using a 22- gauge, 37.5-mm needle (Kendall, Mansfield, MA) and a 3-ml disposable syringe (Sherwood, Davis and Geck, St. Louise, MO). Amniotic fluid obtained from three chick embryos was pooled in a 15-ml plastic conical tube and stored at -20°C until use. Statistical Analysis Data were analyzed using PROC GLM of the SAS Statistical Package (v. 8e, 1999, SAS Institute, Inc., Gary, NC). Significant differences between embryo developmental rates in each group were tested using a post hoc least significant difference (LSD) test. Differences with a probability level (P) of 0.05 or less were considered significant in this study.
85 RESULTS Developmental rates for bovine 8-cell embryos cultured in CR1aa medium supplemented with 5% FBS (control) or in CR1aa supplemented with 20% CAF are summarized in Table 6.1. On day 7 of development a significantly greater (P<0.05) percentage of morula stage embryos resulted from bovine 8-cell embryos cultured in medium supplemented with 20% CAF than in the control culture medium (33% and 14%, respectively). In contrast, culture in FBS-supplemented medium resulted in significantly more (P<0.05) blastocysts and expanded blastocysts compared with CAF-supplemented medium (blastocysts = 31% and 17%, expanded blastocysts = 30% and 18%, respectively; Figure 6.2). On day 8 post-insemination there was no effect (P>0.05) of culture medium on the percentage of embryos that developed to the morula, early blastocyst, blastocyst, expanded blastocyst or hatched blastocyst stage. However, there were significantly more (P<0.05) hatching blastocysts in FBS-supplemented medium than in CAF-supplemented medium (11% and 3%, respectively; Figure 6.2). Although there was no difference (P>0.05) in the total number of blastocysts present in each treatment group on day 9 post-insemination, there were significantly more (P<0.05) hatched blastocysts in the FBS-supplemented medium compared with the CAF- supplemented medium (18% and 9%, respectively; Figure 6.2). Embryo quality grades are presented in Figure 6.3. Although culture in FBS- supplemented medium resulted in a significantly higher (P<0.05) blastocyst yield on day 7 post-insemination, there was no significant difference (P>0.05) in the percentage of grade 1 blastocysts between the two treatment groups (control = 16% and CAF = 15%). On day 8 post-insemination, there was no effect of culture medium on grade 1 or grade 2 embryos, but fewer grade 3 embryos were observed in the CAF-supplemented medium compared with the control medium (9% and 20%, respectively). Similar results were noted on day 9 of in vitro culture. DISCUSSION Bovine embryo culture media often require the addition of serum at the initiation of compaction to stimulate development of the blastocoel (Pinyopummintr and Bavister, 1994; Thompson et al., 1998). Addition of serum to in vitro culture systems, however, has been implicated in several fetal and/or calf abnormalities (Willadsen et al., 1991; Keefer et al., 1994; Behboodi et al., 1995; Hasler et al., 1995; Wilson et al., 1995; Garry
86
Table 6.1. Development of 8-cell bovine embryos cultured in CR1aa supplemented with 5% fetal bovine serum (control) or 20% chick amniotic fluid (CAF)
a Embryos allotted to the control group were cultured in CR1aa + 5% FBS from day 3 to day 7 post-insemination. Embryos allotted to the CAF group were cultured in CR1aa + 20% FBS from day 3 to day 7 post-insemination. b The number of 8-cell embryos allotted to each treatment. c MOR = morulae. d BLST = blastocysts. e HT/HD BLST = hatching and/or hatched blastocysts. f,g Within a column, numbers without a common superscript are significantly different (P<0.05).
87
A Control 50 CAF b 40 s
o a a y
r 30 b
m b E 20 a b a a % of 10
0 MORL Early BLST BLST Expand BLST
B Control 50 CAF
40 s o a a
bry 30
m a E 20
of a % a a 10 a a a a a b 0 MORL Early BLST BLST Expanded Hatching Hatche d BLST BLST BLST
C Control 50 CAF 40 s o y
br 30 m a E 20 a of a
% a b 10 a a a 0 BLST Expand Hatching Hatche d BLST BLST BLST
Figure 6.2. Development of 8-cell bovine embryos cultured in CR1aa supplemented with 5% fetal bovine serum (control) or 20% chick amniotic fluid (CAF) on day 7 of development (A), day 8 of development (B) and day 9 of development (C). Within a developmental stage, columns without a common superscript are significantly different (P<0.05).
88
A Control 50 CAF a
40 s o y s r a o b 30 a a a m bry
E m f 20 a o . a
No % of E 10 a
0 Grade 1 Grade 2 Grade 3 Grade 4
B Control 50 CAF
40
s a
o y 30
br a
m a a 20 a a % of E b 10 a 0 Grade 1Grade 2Grade 3Grade 4
C Control
50 CAF
40
s o
y 30
br m
E a 20 a a
% of a b 10 a a a
0 Grade 1 Grade 2 Grade 3 Grade 4
Figure 6.3. Quality grade of 8-cell bovine embryos cultured in CR1aa supplemented with 5% fetal bovine serum (control) or 20% chick amniotic fluid (CAF) on day 7 of develop- ment (A), day 8 of development (B) and day 9 of development (C). Within a quality grade, columns without a common superscript are significantly different (P<0.05).
89 et al., 1996; Schmidt et al., 1996; Walker et al., 1996; Kruip and den Daas, 1997; Hill et al., 1999, 2001, 2002). In addition, despite extensive quality control measures, the growth promoting properties of FBS and BSA have been observed to vary from batch to batch. Early studies by Sirard and Lambert (1985) found that batches of serum from different animals prepared by identical methods had very different effects on embryo development. Although the embryotropic properties of avian amniotic fluid remain largely undefined, results of the present study suggest that it may serve as a potential alternative to serum supplementation of in vitro embryo culture media, thus possibly circumventing some of the potential risks associated with the use of FBS and other materials of bovine origin. Amniotic fluid is an ultrafiltrate produced in close proximity to the developing fetus and has been reported to be less variable in chemical composition than serum or plasma (Shirley et al., 1985; Gianaroli et al., 1986) and is likely to contain growth factors, hormones and other proteins beneficial to embryo development (Mohamed and Noakes, 1985; Wintour et al., 1986). In addition, amniotic fluid, unlike serum, is not likely to contain unexpected contaminates that would adversely affect embryo development (Gianaroli et al., 1986). Amniotic fluid has been used for the maturation of porcine (Nagi et al., 1990) and bovine (Ocana Quero et al., 1995) oocytes as well as for the culture of murine (Gianaroli et al., 1986; Sueldo et al., 1988; Coetzee et al., 1989; Dorfman et al., 1989; Oettle and Wiswedel, 1989), bovine (Javed and Wright, 1990) and human (Gianaroli et al., 1986; Dorfman et al., 1989) embryos. The results of these studies are somewhat conflicting, however. For example, Gianaroli et al. (1986) and Coetzee et al. (1989) found that culturing murine embryos in undiluted human amniotic fluid did not improve blastocyst rates over controls. In contrast, Dorfman et al. (1989) and Oettle and Wiswedel (1989) reported that culturing murine embryos in undiluted human amniotic fluid significantly increased blastocyst development over controls. Similar discrepancies were noted when culturing human embryos in amniotic fluid. Both Gianaroli et al (1986) and Dorfman et al. (1989) indicated no significant difference in cleavage rates. In contrast, Dorfman et al. (1989) noted a marked decrease in pregnancy rates after culture in human amniotic fluid, whereas, Gianaroli et al. (1986) found no significant difference. One explanation for these discrepancies may be the use of amniotic fluid from women of advanced maternal
90 age by Dorfman et al. (1989). It must be assumed that these women also served as embryo recipients, as it is not stated otherwise in these publications. All in all, these results must be evaluated with caution as each experiment utilized different control media and amniotic fluid samples from varying stages of gestation. Although the use of CAF in the present experiment resulted in significantly fewer total blastocysts on day of 7 of development, there was no significant difference (P<0.05) in the number of embryos classified as quality grade 1 when compared with embryos cultured in CR1aa supplemented with 5% FBS. In addition, there were fewer grade 3 embryos in the CAF treatment group on both day 8 and day 9 of development. These results suggest that the significant increase in blastocyst yield in the control group was composed mostly of embryos of quality grade 3 or grade 4. Results of the present study do indicate, however, that embryos cultured in the presence of CAF undergo a delay in development of ~24 hours. It has been previously reported that development of in vitro-produced bovine embryos is delayed by ~30 to 40 hours when compared with embryos produced in vivo (McGaugh et al, 1974; Prather and First, 1988; Funahashi et al, 1994; Grisart et al., 1994). Van Soom et al. (1992) suggested that this delay indicates that development past the 8- to 16-cell stage is extremely sensitive to culture environment. The fact that embryos cultured in the presence of CAF develop slower than the controls is somewhat concerning. There are reports that indicate that morula and blastocyst stage embryos are able to increase their rate of development to match that of the uterus after transplantation, however (Wilmut and Sales, 1981). This finding would theoretically allow for the normal development of a transferred embryo from the CAF treatment group even though it may be developmentally slower than embryos in the control group. In conclusion, results of the present study indicate that CAF, when added as a supplement to bovine culture medium, provides proteins and growth factors which are able to support in vitro development of bovine embryos. Thus, supplementation of bovine embryo culture medium with chick amniotic fluid may be a feasible alternative to serum supplemented medium for the production of quality bovine embryos, particularly in situations where FBS is not readily available. The fact that CAF-supplemented medium resulted in more grade 1 embryos from fewer total blastocysts is promising and warrants further investigation. More thorough examination of such factors as CAF
91 concentration in the medium, and more importantly, evaluation of pregnancy and calving rates following transfer of CAF-cultured embryos to recipient females, is warranted and must be conducted to determine the potential benefits of this supplement to bovine in vitro embryo production.
92 CHAPTER VII
ESTABLISHMENT AND CULTURE OF CELLS DERIVED FROM DAY-6 CHICK EMBRYOS
INTRODUCTION In vitro embryo culture has been an active area of research for many years; however, few researchers have completely identified the developmental needs of the resulting embryos (Liebfried-Rutledge, 1999). With current embryo culture methods, only 30 to 40% of in vitro-matured and fertilized ova routinely reach the blastocyst stage (Krisher and Bavister, 1998; Lonergan et al., 2003). Moreover, in vitro-produced (IVP) embryos have marked differences in morphology (Greve et al., 1995), timing of development (Van Soom et al., 1997), embryonic metabolism (Khurana and Niemann, 2000) and subsequent pregnancy rates (Enright et al., 2000) when compared with in vivo-derived embryos. In vitro embryo production has also been associated with abnormal fetal and neonatal development and growth, known collectively as large calf or large offspring syndrome (reviewed by Kane, 2003). Specifically, offspring produced from embryos cultured in vitro often exhibit increased birth weights (Willadsen et al., 1991; Keefer et al, 1994; Behboodi et al., 1995), longer gestation lengths (Walker et al, 1996), increased placental anomalies, (Hasler, et al., 1995; Hill et al., 1999, 2001, 2002) and increased incidences of abortion (Walker et al., 1996; Hill et al., 1999) and perinatal (Behboodi et al., 1995; Wilson et al., 1995; Garry et al., 1996; Schmidt et al., 1996; Kruip and den Daas, 1997; Hill et al., 1999) and neonatal mortality (Schmidt et al., 1996; Kruip and den Daas, 1997; Hill et al., 1999). Breathing difficulties (Young et al., 1998) and perturbations in the growth and development of body organs are also common following culture of IVP embryos (Kane, 2003). It has been hypothesized that exposure of pre-elongation embryos to unusual environmental conditions may increase the incidence of large offspring syndrome (reviewed by Kane, 2003); however, the exact cause of this condition still remains unclear. The inadequacies associated with in vitro embryo production have led to the development of several alternative culture systems. One such system involves culturing mammalian embryos in the amniotic cavity of a live chick embryo. Previous reports
93 indicate that factors within the amniotic cavity of developing chick embryos are capable of supporting the maturation of porcine oocytes (Ocampo et al., 1994) and development of embryos in various mammalian species (Blakewood et al., 1989a, 1989b, 1990). In early experiments, chick embryo co-culture consistently resulted in significantly more expanded and hatched blastocysts compared with controls in mice (Blakewood et al., 1989a), pigs (Ocampo et al., 1993), goats (Blakewood et al., 1989a, 1990) and cattle (Blakewood et al., 1989b). In addition, live offspring were produced following transfer of caprine embryos that had been cultured in the chick amnion (Blakewood et al., 1990). Experiments presented in this dissertation indicate that the chick embryo co-culture system is capable of supporting development of bovine embryos at rates similar to more modern embryo culture systems. It is unclear, however, exactly where the benefits of this system lie. One theory is that the chick embryo is a “live” system compared with the static embryo culture systems currently in use. Because the living chick embryo actively regulates the pH, osmolarity and O2 content of its own environment, it may serve as a more stable physiological system for mammalian embryo development than current in vitro culture systems (Blakewood et al., 1989a). An alternative hypothesis is that the chick embryo and/or amnion actively synthesize growth factors and other factors that may stimulate growth and development of mammalian embryos, a benefit unavailable in current embryo culture systems without supplementation of fetal bovine serum (FBS) or bovine serum albumin (BSA) (Blakewood et al., 1989a). It is also possible that the embryotropic properties of the chick embryo amnion are a combination of both aforementioned hypotheses. Therefore the objective of the current research is to develop a protocol for the in vitro culture of fetal chick cells and chick amnion cells as a preliminary experiment to study the contributions of these cell types to bovine embryo development. More specifically, we wanted to: 1) determine which of two media support cell growth and proliferation most efficiently, 2) evaluate the growth characteristics of these cells in vitro and 3) evaluate the post-thaw viability of these cells.
94 MATERIALS AND METHODS Experimental Design Experimental designs for Experiment 7.1 and 7.2 are given in Table 7.1 and Table 7.2, respectively. Treatment groups for Experiment 7.1 consisted of bovine fibroblasts cultured in modified Tissue Culture Medium-199 (mTCM; Treatment A; control), bovine fibroblasts cultured in modified Dulbecco’s Modified Eagle’s Medium (mDMEM; Treatment B; control), fetal chick cells cultured in mTCM (Treatment C) and fetal chick cells cultured in mDMEM. Treatment groups for Experiment 7.2 consisted of bovine fibroblasts cultured in mTCM (Treatment A; control), bovine fibroblasts cultured in mDMEM (Treatment B, control), chick amnion cells cultured in mTCM (Treatment C) and chick amnion cells cultured in mDMEM (Treatment D). In both experiments the parameters evaluated for each treatment included number of days to 50% confluence, number of days before each of two passages, evaluation of morphology and cell type, population doublings, plating efficiency and post-thaw viability. Primary cultures of bovine fibroblasts, fetal chick cells or chick amnion cells were taken and processed. All cell types were maintained in both mTCM and mDMEM to determine the effect of culture media on cell growth and proliferation. Media were changed every 48 hours and cells were passaged at ~ 95% confluence. Cells were passaged two times before being frozen and stored in liquid nitrogen. Plating efficiency was determined 24 hours after passage and growth curves were performed every 24 hours for 5 days or until cells reached confluence. Cell viability was determined at each passage. To determine post-thaw viability, cells were maintained in liquid nitrogen for a minimum of 48 hours before being thawed and replated. Plating efficiency was determined 24 hours after thawing, and viability was determined when cells reached ~ 95% confluence. Experimental Methods Media Preparation Modified TCM was prepared by adding 5 ml of FBS (SH30070.02 HyClone, Logan, UT) and 0.05 ml of gentamicin (50 mg/ml; 15750-060, Gibco, Grand Island, NY) to 45 ml of TCM-199 (Gibco Laboratories, Grand Island, NY). Modified DMEM was
95
Table 7.1. Experimental design for Experiment 7.1 for growth and proliferation of fetal chick cells
Treatment group Cell type Medium type
a Treatment A Bovine Fibroblasts mTCM
b Treatment B Bovine Fibroblasts mDMEM
Treatment C Fetal Chick Cells mTCMa
Treatment D Fetal Chick Cells mDMEMb a mTCM = modified Tissue Culture Medium. b mDMEM = modified Dulbecco’s Modified Eagle’s Medium.
96
Table 7.2. Experimental design for Experiment 7.2 for growth and proliferation of chick amnion cells
Treatment group Cell type Medium type
a Treatment A Bovine Fibroblasts mTCM
b Treatment B Bovine Fibroblasts mDMEM
Treatment C Chick Amnion Cells mTCMa
Treatment D Chick Amnion Cells mDMEMb a mTCM = modified Tissue Culture Medium. b mDMEM = modified Dulbecco’s Modified Eagle’s Medium.
97 prepared by adding 7.5 ml of FBS, 0.5 ml of Modified Eagle Medium (MEM) amino acid solution (10X, 11140-050, Gibco Laboratories, Grand Island, NY) and 0.05 ml of gentamicin to 41.5 ml of DMEM (10313-021, Gibco, Grand Island, NY). All media was filtered through 0.2 µm Acrodisc filters (Pall Corporation, East Hills, NY) and allowed to equilibrate in a 39°C and 5% CO2 incubator for a minimum of 30 minutes. Sterile, pre- warmed TCM-199 was used to wash the serum residue from cells prior to trypsinization. Shell-less Chick Culture for Chick Embryo Retrieval The basis for the shell-less chick embryo culture procedure was modified from that previously described by Blakewood and Godke (1989). Briefly, fertilized domestic chicken eggs were obtained on the day of oviposition and stored at 10°C until the onset of incubation. Six days prior to the day of embryo retrieval, eggs were removed from cold storage and allowed to come to room temperature (25°C) before placing them in a 37°C commercial egg incubator. After 72 hours of incubation, the eggs were rinsed with a 70% ethanol solution (Aaper Alcohol and Chemical, CO, Shelbyville, KY) and then coated with 7.5% providone iodine solution (Betadine®, Purdue Fredrick Company, Stanford, CT) and allowed to air dry in a horizontal position to ensure proper positioning of the chick embryo. Egg contents were removed by making a crack in the shell by striking the center of the egg against the rim of a sterile beaker. The egg contents were then released into a 100-ml plastic embryo collection dish (Veterinary Concepts, Spring Valley, WI) that had been covered with a piece of cellophane kitchen wrap (Saran®, Dow Chemical, Midland, MI). The dish, including the egg contents, was loosely capped with a plastic lid (Veterinary Concepts, Spring Valley, WI) and maintained in an incubator at 38°C for an additional 72 hours before retrieval of the chick embryo and surrounding amnion. On day 6 of shell-less culture the chick embryo and surrounding amnion were dissected away from the rest of the egg components and then placed in a 35 x 10-mm Falcon® plastic Petri dish (Becton and Dickerson, Lincoln Park, NJ).They were then rinsed three times with sterile Dulbecco’s Phosphate Buffered Saline (DPBS; D1408, Sigma-Aldrich Inc., St. Louis, MO) containing 1% FBS. The amniotic membrane was gently dissected away from the chick embryo and placed in a 35 x 10-mm plastic petri dish of mDPBS (mDPBS: 10 ml DPBS containing 0.01 ml of gentamicin). Any remaining extraembryonic membranes were trimmed away from the chick embryo and the embryo
98 was placed in a 15-ml plastic conical tube (Corning, New York, NY) of mDPBS until processing. Establishment of Bovine Fibroblast Cell Lines. Skin biopsies were taken from three adult cows of similar age (3 to 5 years of age), body weights (408 to 498 kg) and body condition scores (4.5 to 5.5) using a 6-mm diameter biopsy punch (Miltex Instrument Co, Bethphage, NY). Biopsies were obtained from the dorsal aspect of the shoulder directly over the supraspinatus muscle. The tissue was maintained in mDPBS until processing. Using heat sterilized iris scissors, the dermal portion of the biopsy was finely minced into pieces measuring ~1 mm2 in mDPBS. The tissue suspension was transferred to a 15-ml plastic conical tube and washed in DPBS by swirling. The tissue pieces were allowed to settle to the bottom of the tube, the supernatant discarded and 3 ml of fresh mDPBS was added. The process was repeated two additional times. After the last wash the tissue was resuspended in 3 ml of culture medium (either mTCM or mDMEM) and transferred to a 35 x 10-mm plastic petri dish. All cultures were maintained at 39°C and 5% CO2 in a humidified atmosphere. Fibroblasts began to grow from the tissue explants by about the fifth day at which time the tissue explants were removed from the culture. Once a fibroblast cell layer was established, the cells were maintained in the 35 x 10-mm plastic petri dish until they reached ~95% confluency. Cells were then trypsinized by cold trypsinization. Briefly, cells were rinsed two times with pre-warmed TCM and then maintained in cold (4°C) trypsin (Trypsin EDTA, 1X, T3924, Sigma-Aldrich Inc., St. Louis, MO) at room temperature for 5 minutes and then incubated at 39°C for an additional 5 to 10 minutes. The resulting cell suspension was transferred to a 15-ml plastic conical tube and the dish rinsed twice with culture medium. The cells were then washed by centrifugation at 500 x g for 10 minutes and the resulting pellet resupsended in 1 ml of culture medium for counting. Small tissue culture flasks (25 cm2; Costar, Cambridge, MA) were seeded with 1x106 cells/ml and filled with 2 to 3 ml of culture medium before being placed in incubation. The cells were passaged twice prior to cryopreservation in TCM or DMEM supplemented with 20% FBS and 5% dimethylsulfoxide (DMSO; D128-500, Fischer Scientific, Fairlawn, NJ) and then frozen in aliquots of 1x106 cells per cryovial.
99 Establishment of Fetal Chick Cell Lines Primary cell cultures were prepared from day 6 chicken embryos. After removal of the head and limbs, the chick embryos were finely minced into pieces measuring ~1 mm2 in mDBPS using heat-sterilized iris scissors. Tissue pieces were washed three times in mDPBS by allowing the tissue to settle to the bottom of the petri dish and removing the supernatant. To further disassociate the cells, the tissue pieces were subjected to cold trypsinization as described above. Any undigested tissue pieces were allowed to settle to the bottom of the dish and the supernatant was removed and placed in a 15-ml plastic conical tube. The cells were then washed by centrifugation at 500 x g for 10 minutes in 4 ml of mTCM. The resulting pellet was subjected to cold trypsinization one additional time. Prior to centrifugation, 0.5 ml of the resulting cell suspension was diluted in 3 ml of mTCM and 0.5 ml in 3 ml of mDMEM. Each cell suspension was then centrifuged at 500 x g for 10 minutes and counted using a hemocytometer. Small tissue culture flasks (25 cm2) were seeded at a density of 1x106 cells per ml and cultured at 39°C and 5%
CO2 in a humidified atmosphere. The cells were passaged twice prior to cryopreservation in either TCM or DMEM supplemented with 20% FBS and 5% DMSO and then frozen in aliquots of 1x106 cells per cryovial. Establishment of Chick Amnion Cell Lines The basic procedure for the establishment of chick amnion cell lines was adapted from that previously reported for use in humans (Neeper et al., 1990). Using heat sterilized forceps, pieces of the amnion were washed through three 35 x 10-mm plastic petri dishes of mDPBS before being subjected to cold trypsinization as described above. Any tissue pieces remaining after cold trypsinization were manually disassociated by pipetting. The resulting cell suspension was aspirated from the dish and placed in a 15- ml plastic conical tube and the dish washed twice with 2 ml of mTCM. The rinse medium was added to the conical tube and the suspension mixed well. A 2.0 ml aliquot of cell suspension was added to each of two 15-ml plastic conical tubes and the cells in each tube were counted using a hemocytometer. Each cell suspension was then centrifuged at 500 x g for 6 minutes and resuspended in the appropriate culture media (mTCM or mDMEM). The cells were subpassaged twice prior to cryopreservation in either TCM or
100 DMEM supplemented with 20% FBS and 5% DMSO and then frozen in aliquots of 1x106 cells per cryovial. Determination of Cell Viability Cell viability was determined by a dye exclusion test with trypan blue (0.05%; Sigma-Aldrich Inc., St. Louis, MO) as previously described by Phillips (1973). Briefly, cells were trypsinized by cold trypsinization. After centrifugation, the pellet was resupsended in 1 ml of culture medium. The cell suspension was diluted with trypan blue at a 1:1 (v/v) ratio and cells were counted using a hemocytometer. Exclusion of trypan blue from the interior of the cell was considered an indicator of viability. Viability was determined by the formula [(number of live cells/total number of cells) x 100] and reported as a percentage. Determination of Plating Efficiency To determine plating efficiency, each well of a Nunc® 4-well plate (Apogent Technologies, Rochester, NY) was seeded with ~15,000 cells and incubated at 39°C and 5% CO2. After 24 hours, medium was removed from each well and pooled for counting. The pooled cell suspension was centrifuged at 500 x g for 6 minutes and the pellet resuspended in 1 ml of culture medium. The cell suspension was diluted with culture medium at a 1:1 (v/v) ratio and cells were counted using a hemocytometer. Plating efficiency was determined by the formula [1 – (number of unplated cells/total number of cells seeded)] x 100 and was reported as a percentage. Determination of Population Doublings For determination of amnion cell population doublings, five 35 x 10-mm plastic petri dishes were seeded at 300,000 cells/ml and incubated at 39°C and 5% CO2 in a humidified atmosphere. Every 24 hours, one petri dish from each treatment was removed from incubation, trypsinized and counted to determine total cell number. Population doublings were determined by the formula [log (final concentration/initial concentration) x 3.33]. Statistical Analysis Data were analyzed using PROC GLM of the SAS Statistical Package (v. 8e, 1999, SAS Institute, Inc., Gary, NC). To determine statistical differences for time to first, second and post-thaw passages, cell viability, plating efficiency and population doublings means were compared using a post hoc least significant difference (LSD) test.
101 Differences with a probability (P) value of 0.05 or less were considered significant in this study. RESULTS The growth characteristics of in vitro-cultured fetal chick cells are summarized in Table 7.3. There was no effect of culture media (P>0.05) on the time required for fetal chick cells to become confluent. With the exception of time required for primary culture, these cells displayed a similar growth pattern to bovine fibroblasts (P>0.05). Plating efficiency was not affected (P>0.05) by culture media in either the first (84.2% and 70.4% for mTCM and mDMEM, respectively) or the second (84.7% and 93.2% for mTCM and mDMEM, respectively) passage (data not shown). Similar results were observed for viability [first passage: mTCM = 98.3% and mDMEM = 90.4%; second passage: mTCM = 85.0% and mDMEM = 91.9% (data not shown)]. Plating efficiency and viability were not evalutated for primary culture or post-thaw passages. The growth characteristics of in vitro-cultured chick amnion cells are summarized in Table 7.4. There was no effect of culture media (P>0.05) on the time required for chick amnion cells to become confluent (Table 7.4). Plating efficiency was not affected (P>0.05) by culture media in primary culture (93.4% and 94.0% for mTCM and mDMEM, respectively), first passage (73.9% and 59.0% for mTCM and mDMEM, respectively), second passage (72.7% and 69.4% for mTCM and mDMEM, respectively) or after freezing and thawing (47.0% and 39.5% for mTCM and mDMEM, respectively; data not shown). Similar results were observed for viability (first passage: mTCM = 74.0% and mDMEM = 72.1%; second passage: mTCM = 66.0% and mDMEM = 58.0%; post-thaw: mTCM = 77.0% and mDMEM = 63.4%; data not shown). Population doublings were evaluated for each passage to determine if chick amnion cells were actively proliferating in vitro. When chick amnion cells were cultured in mTCM, population doubling rate increased over the five day evaluation period for the first passage, but remained relatively constant for the second passage and after freezing and thawing (Figure 7.1). There was no significant difference (P>0.05) between passages, however. In contrast, when chick amnion cells were cultured in mDMEM, the population doubling rate remained fairly constant for all three passages evaluated (Figure 7.2). As with the cells grown in mTCM, there was no significant difference between passages (P>0.05).
102
Table 7.3. Growth characteristics of fetal chick cells during in vitro culture in mTCM and mDMEM culture media
a BF = bovine fibroblasts. b FC = fetal chick cells. c Data from four replicates have been combined. d,e Within a column, values without a common superscript are significantly different (P<0.05).
103
Table 7.4. Growth characteristics of chick amnion cells during in vitro culture in mTCM and mDMEM culture media
a BF = bovine fibroblasts. b FC = fetal chick cells. c Data from four replicates have been combined. d,e Within a column, values without a common superscript are significantly different (P<0.05).
104
Passage 1 Passage 2 Post-Thaw
5
4
oublings 3
ion D t 2
opula
P 1
0 12345 Days of Growth Figure 7.1. Growth of chick amnion cells in vitro in mTCM culture medium.
105
Passage 1 Passage 2 Post-Thaw
5
4 gs
ublin
o 3
D ion t 2
pula o
P 1
0
12345
Days of Growth
Figure 7.2. Growth of chick amnion cells in vitro in mDMEM culture medium.
106 Both epithelial cells and fibroblast cells were present in in vitro chick amnion cell cultures. During the first passage, epithelial cell populations increased through day 3 but then declined. In contrast, fibroblast cell populations increased progressively throughout the five day evaluation period (Figure 7.3). Similar results were observed during the second passage and after freezing and thawing (Figures 7.4 and 7.5, respectively). DISCUSSION Results of the present study indicate that cultures of fetal chick cells and chick amnion cells can be maintained in vitro. Furthermore, these culture systems appear to be able to promote cell growth and proliferation through multiple passages. In general, cultures of both cell types maintained under the conditions described herein survive well in vitro and do not appear to be adversely affected by freezing and thawing. The two media (mTCM and mDMEM) were selected for this experiment because one (mTCM-199) is a traditional medium developed for chick embryonic cells and the other (mDMEM) was developed for the culture of amniocytes. TCM-199, the major component of mTCM, was originally reported for the culture of chick embryo fibroblasts (Morgan et al., 1950) and is now commonly used for the culture of many mammalian cell types. It is the traditional medium used in our laboratory for the cultivation and maintenance of bovine and caprine fibroblast cell lines. DMEM was chosen as an alternative medium as it has been successfully used for the propagation of human amnion cells (Neeper et al., 1990; Wang et al., 1992) and is the base medium for several commercial human amnniocyte cell culture media. Plating efficiency has been suggested to be a highly sensitive test for the detection of noxious substances in the culture environment (Seemayer et al., 1987). Since plating efficiency was not significantly different between the two media, it can be concluded that there were no immediate adverse affects of either culture medium on fetal chick cells. It must be pointed out, however, that although plating efficiency can be used as a measure of the cell’s response to its surroundings, it does not indicate if those surroundings are capable of supporting sustained cell growth. The results of the trypan blue exclusion test indicate that there was no significant difference in cell viability between the two media. Viability was determined at the end of each passage and when taken together with the outcome of the plating efficiency test, it can be concluded that both media are capable of sustaining fetal chick cells in vitro.
107
Fibroblast Cells Epithelial Cells Total Cells 1
0.8 ) 6
0 1 x
( 0.6
y it s
n e 0.4
ll D e C 0.2
0
12345 Days of Growth
Figure 7.3. Cell populations of chick amnion cells cultured in vitro in mTCM during the first passage.
108
Fibroblast Cells Epithelial Cells Total Cells
6
5 ) 6
0
1 4 x (
y it
s 3
n e
ll D 2 e
C 1
0
12345 Days of Growth
Figure 7.4. Cell populations of chick amnion cells cultured in vitro in mTCM during the second passage.
109
Fibroblast Cells Epithelial Cells Total Cells
3.5
3
) 6
0 2.5 1 x
( 2 y it s
n e 1.5
ll D e 1 C 0.5
0
12345 Days of Growth
Figure 7.5. Cell populations of chick amnion cells cultured in vitro in mTCM after freezing and thawing. .
110 As with the fetal cells, both media appear to support the growth of chick amnion cells in vitro. Neither plating efficiency nor viability were significantly different between the two media. These findings were somewhat unexpected since DMEM has been used quite successfully for the culture of human amnion cells (Neeper et al., 1990; Wang et al., 1992). However, although both media are capable of sustaining growth of chick amnion cells in vitro, neither appear to promote adequate cell proliferation. With the exception of the primary culture passage of mTCM, evaluation of population doublings indicate that the rate of cell growth was approximately equal to the rate of cell death. These findings suggest that while capable of sustaining cell growth, neither medium is suitable for the proliferation of chick amnion cells in vitro at an optimal level. General morphometic features of both the fetal chick cells and chick amnion cells indicate a mixed culture of epithelial cells and fibroblast cells. Epithelial cells appeared generally round and tended to form dense colonies. In contrast, fibroblasts were spindle shaped with long, branching pseudopodia and tended to form monolayers rather than colonies. It is generally believed that trypsinization of cell cultures, as performed in this experiment, selects against epithelial cells, resulting in cell populations predominated by fibroblast cells (Kenneth Bondioli, personal communication). This does not appear to be the case when culturing bovine amnion cells as epithelial cells were present in all three passages evaluated. Further passages may be required in order to observe this phenomenon. When population doublings were evaluated within a single passage, epithelial cell numbers increased progressively through day 3 of culture before declining. This suggests that neither media type are conducive to epithelial cell proliferation. It may be necessary to use a medium capable of supporting growth and proliferation of both epithelial and fibroblast cells if the integrity of the amnion is to be maintained. In conclusion, we have successfully developed a repeatable protocol for culturing both fetal chick cells and chick amnion cells. However, both systems may require some optimization. This is believed to be the first report of culture of cells derived from the chick embryo amnion. The current research provides a tool for in vitro studies of the amnion and the chick embryo and provides an in vitro model for the study of the effects of chick embryo co-culture on mammalian embryo development.
111 CHAPTER VIII
THE INCIDENCE OF APOPTOSIS IN BOVINE EMBRYOS AFTER IN VITRO CULTURE AND CHICK EMBRYO CO-CULTURE
INTRODUCTION Although much research has been devoted to the development and improvement of in vitro embryo production techniques, differences still remain between in vitro- and in vivo-derived embryos. The development of immature bovine oocytes to the blastocyst stage appear to have plateaued at about 30 to 40% following in vitro maturation, fertilization and culture (Krisher and Bavister, 1998; Lonergan et al., 2003). In addition, the quality of blastocysts produced in vitro rarely, if ever, reach that of in vivo-derived blastocysts (Duby et al., 1997; Boni et al., 1999; Viuff et al., 1999), and marked differences have been reported to exist between in vivo- and in vitro-derived embryos with regard to morphology (Greve et al., 1995), timing of development (Van Soom et al., 1997), embryonic cell numbers (Van Soom et al., 1997), embryonic metabolism (Khurana and Niemann, 2000) and gene expression (Wrenzycki et al., 1996; Knijn et al., 2002) and incidences of chromosomal abnormalities (Viuff et al., 2001). With many of the current embryo culture systems, the addition of fetal bovine serum (FBS) is necessary for successful development of the embryonic blastoceol and continued embryo development (Pinyopummintr and Bavister, 1994; Thompson et al., 1998). However, serum and other compounds of bovine origin have been implicated in delayed embryo development (Van Soom et al., 1997), reduced embryo quality (Duby et al., 1997; Boni et al., 1999; Viuff et al., 1999), reduced pregnancy rates (Enright et al., 2000) and numerous fetal and/or calf abnormalities (Thompson et al., 1995; Walker et al., 1996; Sinclair et al., 1999). Apoptosis is a critical process during pre-implantation development for the disposal of excess cells, cells that are in the way, abnormal or potentially dangerous (Gjorret et al., 2003). However, elevated levels of apoptotic cells have been correlated with abnormal embryo morphology (Hardy et al., 1989), and it has been hypothesized that when apoptosis surpasses a certain “threshold level” further embryonic development is compromised (Tam, 1988; Juriscova et al., 1998; Hardy, 1999) and nonviable offspring are eliminated (Byrne et al., 1999). In addition, apoptosis may be a cellular response to stress and suboptimal culture conditions (Gjorret et al., 2003) and
112 evidence suggests that the fetal anomalies and increased incidences of abortion associated with in vitro culture may be the result of distortions to normal apoptotic processes (Sadler and Hanter, 1994). As an alternative to traditional in vitro culture systems utilizing serum supplementation, Blakewood and Godke (1989) developed an alternative culture system in which pre-implantation mammalian embryos were cultured in the amniotic cavity of live chicken embryos. This culture system has been successful in supporting the development of pre-implantation embryos from several mammalian species including mice (Blakewood et al., 1989a), goats (Blakewood et al., 1989a), cattle (Blakewood et al., 1989b; Murakami et al., in press) and pigs (Ocampo et al., 1993). Moreover, live transplant offspring have been produced following chick embryo co-culture of 2- to 8-cell caprine embryos (Blakewood et al., 1990). In this study, we propose to compare the occurrence of apoptosis in fresh and frozen-thawed in vivo-derived embryos and in embryos cultured in vitro and in the chick embryo co-culture system in order to evaluate the efficacy of the use of the chick embryo co-culture system as a potential embryo culture environment. Traditional analysis of apoptosis in pre-implantation mammalian embryos have utilized the TUNEL reaction to label fragmented DNA in apoptotic cells. However, in this study we evaluated the incidence of apoptosis using a fluorescent labeled inhibitor to caspases-3 and -7. MATERIALS AND METHODS Experimental Design The synchronization and superovulation scheme for in vivo embryo production is illustrated in Figure 8.1. In vivo embryos were collected from beef females which were synchronized and superstimulated. Briefly, females were synchronized with CIDRs for 8 days. On day 11 of the synchronized cycle, females were superstimulated with follicle stimulating hormone (FSH) in a decreasing dose schedule and inseminated 12 and 24 hours after visual observation of estrus. Embryos were nonsurgically recovered either 6.5 or 7 days after the onset of estrus (estrus = day 0). Embryos were fluorescently labeled and evaluated for caspase activity immediately upon recovery. The process for production of in vitro embryos is illustrated in Figure 8.2. In vitro embryos were produced using the standard IVF protocol for our laboratory.
113
EXPERIMENTAL DESIGN
Figure 8.1. Experimental design for the production and fluorescent labeling of in vivo-derived embryos.
114 Briefly, oocytes were matured for 20 to 22 hours before being exposed to motile spermatozoa from a bull of proven fertility for IVF. After 5 hours of co-incubation with spermatazoa, oocytes were removed from fertilization and placed into culture in CR1aa. On day 3 post-insemination medium was removed and replaced with CR1aa containing 5% FBS. Embryos were fluorescently labeled and evaluated for caspase activity on day 7 post-insemination. Because it was not possible to perform all treatments concurrently, two replications of IVF were performed (IVF-I and IVF-II). The process for production of chick co-cultured embryos is illustrated in Figure 8.3. Chick co-cultured embryos were produced as described for in vitro embryos; however on day 3 post-insemination, bovine embryos were embedded in agar and injected into the amniotic cavity of 96-hour chick embryos. After 96 hours of culture, embryos were recovered from the chick amnion and fluorescently labeled for caspase activity. Experimental Methods Production of In Vivo Embryos To obtain in vivo embryos, clinically healthy, non-lactating, beef females (age = 4 to 7 years of age; body weights = 386 to 612 kg; BCS = 4 to 6) were first synchronized with a progesterone device (CIDR; Agtech, Inc, Manhattan, KS) for 8 days. Two days ®; prior to CIDR withdrawl, prostaglandin F2α (25 mg Lutalyse Pharmacia and Upjohn Company, Kalamazoo, MI) was administered to ensure complete regression of the corpus luteum (CL). At the time of CIDR withdrawl, GnRH (100 µg; Factrel®; Fort Dodge Animal Health, Fort Dodge, IA) was administered to induce an LH surge. Beginning on day 11 of the synchronized cycle (CIDR removal = Day 0), females received porcine FSH (Sioux Biochemical, Inc., Sioux City, IA) twice daily, in decreasing doses, for 4 consecutive days: 5 units on the first day; 4 units on the second day; 4 units on the third day; and 3 units on the fourth day. Prostaglandin-F2α (25 mg) was administered concomitant with the sixth and seventh doses of FSH. Females were inseminated 12 and 24 hours after visual observation of standing heat with two straws of semen from a bull of proven fertility. Embryos were collected nonsurgically from donors 6.5 or 7 days after the first insemination. All embryos were evaluated for morphological stage and quality grade by three technicians and a mean quality grade was determined.
115
EXPERIMENTAL DESIGN
Figure 8.2. Experimental design for the production and fluorescent labeling of embryos cultured in vitro.
116
EXPERIMENTAL DESIGN
Figure 8.3. Experimental design for the production and fluorescent labeling of embryos cultured in the chick embryo amnion.
117 Frozen in vivo-derived embryos were graciously provided by Dr. Neil Schrick (University of Tennessee, Knoxville, TN). All embryos were obtained from beef females (Charolais, Angus or Senepol) and were frozen in glycerol. All embryos were thawed for 2 minutes in air and cryoprotectant removed using a 3-step protocol. Production of In Vitro Embryos Media Preparation Brackett-Oliphant (BO) media were prepared as previously described by Brackett and Oliphant (1975). Briefly, BO-A stock solutions contain 150.2 mM NaCL (S-5886, Sigma-Aldrich Inc., St. Louis, MO), 2.7 mM KCl (P-5405, Sigma-Aldrich Inc. St. Louis,
MO), 3.0 mM CaCl2·2H2O (C-7902, Sigma-Aldrich Inc., St. Louis, MO), 1.0 mM
NaH2PO4·H2O (S-9638, Sigma-Aldrich Inc., St. Louis, MO), 0.6857 mM MgCl2·6H2O (M- 2393, Sigma-Aldrich Inc., St. Louis, MO) and 0.1 ml of 0.5% phenol red (P-0290, Sigma- Aldrich Inc., St. Louis, MO) per 500 ml of sterile water. BO-B stock solutions contain
154.0 mM NaHCO3 (S-8875, Sigma-Aldrich Inc., St. Louis, MO) and 0.04 ml of 0.5% phenol red per 200 ml of sterile water. CR1aa medium was prepared as previously described by Rosenkrans and First (1994). Briefly, CR1aa stock solution was prepared by adding 114.7 mM NaCl, 1.6 mM
KCl, 26.2 mM NaHCO3, 0.3999 mM pyruvic acid (P-4562, Sigma-Aldrich Inc., St. Louis, MO), 2.5 mM L(+) lactic acid (L-4388, Sigma-Aldrich Inc., St. Louis, MO), 0.5195 mM glycine (G-8790, Sigma-Aldrich Inc., St. Louis, MO), 0.5051 mM L-alanine (A-7469, Sigma-Aldrich Incorporated, St. Louis, MO) and 0.2 ml of 0.5% phenol red per 100 ml of sterile water. Fertilization stock medium (BO-AB) was prepared by adding 1.2 mM pyruvic acid, 0.05 ml of gentamicin (50 mg/ml; 15750-060, Gibco, Grand Island, NY) and 0.018 ml of heparin sodium (10,000 IU/ml; Elkins-Sinn, Cherry Hill, NJ) per 38 ml of BO-A stock solution and 12 ml of BO-B stock solution. Fertilization medium (BO-caffeine) was prepared by adding 0.0243 g of caffeine sodium (C-4144, Sigma-Aldrich Inc., St. Louis, MO) per 25 ml of BO-AB. Sperm washing medium (0.6% BSA-BO) was prepared by adding 0.03 g of bovine serum albumin (BSA; Fraction V, A-4503, Sigma-Aldrich Inc., St. Louis, MO) to 5 ml of BO-AB. Oocyte washing medium (0.3% BSA-BO) was prepared by adding 0.03 g of BSA (Fraction V) to 10 ml of BO-AB. TCM oocyte washing medium was prepared by adding 0.03 g of BSA (Fraction V) to 10 ml of Tissue Culture Medium-199
118 (Eagle’s salt) with 25 mM HEPES Buffer (TCM-199; Gibco Laboratories, Grand Island, NY). CR1aa culture medium for day 0 to day 3 was prepared by adding 0.2 ml of Basal Medium Eagle (BME) amino acid solution (50X, B-6766, Sigma-Aldrich Inc., St. Louis, MO), 0.1 ml of Modified Eagle Medium (MEM) amino acid solution (10X, 11140- 050, Gibco Laboratories, Grand Island, NY), 0.01 ml of gentamicin, 1.1 mM L-glutamine (G-5763, Sigma-Aldrich Inc., St. Louis, MO) and 0.03 g of BSA (fatty acid free, A-7511, Sigma-Aldrich Inc., St. Louis, MO) to 9.5 ml of CR1aa stock solution. CR1aa culture medium for day 3 to day 7 was prepared by adding 0.2 ml of BME amino acid solution, 0.1 ml of MEM amino acid solution, 0.5 ml of FBS (SH30070.02; HyClone, Logan, UT), 0.01 ml of gentamicin, 1.1 mM L-glutamine and 0.03 g of BSA (fatty acid free) to 9.5 ml of CR1aa stock solution. In Vitro Fertilization Oocytes were purchased weekly from BoMed, Inc. (Madison, WI). The oocytes were shipped in a portable incubator (~39°C) and allowed to mature during travel (20 to 22 hours). Upon arrival at the laboratory, shipping vials were removed from the portable incubator and stored in a humidified atmosphere of 5% CO2 in air at 39°C until insemination. Semen was prepared by thawing a straw of frozen semen from a Holstein bull (CSS 7H5188, Genex Cooperative Inc., Shawano, WI) of proven fertility in a 39° to 40°C water bath for ~45 seconds. The contents of the straw were then deposited into a 15-ml plastic conical tube (Corning, New York, NY) and 9 ml of BO-caffeine was added to the tube. The suspension was then centrifuged for 6 minutes at 200 x g. After centrifugation the supernatant was removed and the remaining pellet was resuspended in an additional 9 ml of BO-caffeine. The resulting suspension was centrifuged for 6 additional minutes at 200 x g. The supernatant was removed again and the pellet resuspended in 4 ml of BO- caffeine and 4 ml of 0.6% BSA-BO and placed in a 39°C incubator for ~10 minutes to equilibrate. After equilibration, four 80-µl insemination drops were created in a preheated 35 x 10-mm Falcon® plastic Petri dish (Becton and Dickinson, Lincoln Park, NJ) and covered with warmed mineral oil (M-8140, Sigma-Aldrich Inc., St. Louis, MO). After the 20 to 22 hour maturation period, cumulus oocyte complexes were washed through two 35 x 10-mm plastic petri dishes containing 2 to 3 ml of 0.3% BSA-
119 BO. Then, 10 to 20 oocytes were placed in each of the 80-µl insemination drops previously prepared and were co-incubated for 5 hours at 39°C and 5% CO2 in air. After fertilization, oocytes were washed through a single 35 x 10-mm plastic petri dish of TCM oocyte wash media. Cumulus cells were removed by vortexing for 3 minutes in 1 ml of TCM-199 containing 0.001 g of hyaluronidase (H-3506, Sigma-Aldrich Inc., St. Louis, MO). After vortexing, oocytes were washed through two 35 x 10 mm plastic petri dishes containing TCM oocyte wash and one 35 x 10-mm plastic petri dish of CR1aa. Then, 10 to 15 oocytes per drop were placed in 50-µl drops of CR1aa covered with warmed mineral oil and incubated in a modular incubator containing 5%
CO2, 5% O2, and 90% N2 at 39°C. Chick Embryo Co-Culture The basic procedure for chick embryo co-culture was modified from that previously described by Blakewood and Godke (1989). Briefly, fertilized domestic chicken eggs were collected on the day of oviposition and stored at 10°C until the onset of incubation. Four days prior to the introduction of bovine embryos, the eggs were removed from cold storage and allowed to come to room temperature (25°C). They were then placed in a commercial egg incubator maintained at 37°C and 80 to 90% relative humidity. After 72 hours of incubation, the eggs were rinsed with a solution of 70% ethanol (Aaper Alcohol and Chemical Company, Shelbyville, KY) and then coated with 7.5% providone iodine solution (Betadine®, Purdue Fredrick Company, Stanford, CT) and allowed to air dry in a horizontal position to ensure proper positioning of the chick embryo. To remove the egg contents, a crack was made in the shell by striking the center of the egg against the rim of a sterile beaker. The egg contents were then released into a 100-ml plastic embryo collection dish (Veterinary Concepts, Spring Valley, WI) covered by a piece of cellophane kitchen wrap (Saran®, Dow Chemical, Midland, MI). The dish, including the egg contents, was loosely capped with a plastic lid (Veterinary Concepts, Spring Valley, WI) and maintained in an incubator at 38°C for an additional 24 hours before introducing the reconstructed bovine embryos into the amniotic cavity of the chick embryo. The procedure for embedding embryos in agarose was modified from that of Willadsen (1979) for the culture of ovine embryos in the ligated oviducts of sheep. On
120 day 3 of culture, two or three 8-cell bovine IVF embryos were placed in a 50-µl drop of low-melting agarose solution [1.2% agarose (A-9799, Sigma-Aldrich Inc., St. Louis, MO) dissolved in Dulbecco’s Phosphate-Buffered Saline (DPBS; D-1408, Sigma-Aldrich Inc., St. Louis, MO)] at ~37oC and then aspirated into a beveled glass pipette (200 µm outer diameter). After holding in air for ~30 seconds the agar (containing the embryos) within the pipette was expelled as a gel cylinder into room temperature TL-HEPES (BioWittaker, Walkersville, MD). The agarose cylinder, including the bovine embryos, was then aspirated back into the beveled pipette, and carefully expelled into the amniotic cavity of a 96-hour chick embryo. After introducing the embedded embryos into the amniotic cavity the shell-less culture systems were returned to incubation for an additional 72 hours at 39oC. Thus, a single agarose cylinder containing two or three 8-cell bovine IVF embryos was co- cultured in the amnion of each developing chick embryo for 4 days. On day 7 of in vitro culture, the amnion containing the agarose cylinder was carefully dissected away from the rest of the egg components and placed in a 35 x 10- mm plastic petri dish. The amnion was then rinsed several times with DPBS containing 1% FBS. After rinsing, the petri dish was placed under a stereomicroscope (Nikon SMZ- 2B; Tokyo, Japan) equipped with an overhead light source and the agarose cylinder was removed from the amniotic cavity by using two 22-guage, 37.5-mm needles (Kendall, Mansfield, MA) to make a small opening in the amniotic membrane. The agarose cylinder was carried from the amnion with the escaping amniotic fluid, settling on the bottom of the dish. The cylinder was then washed twice in CR1aa medium and dissected using two 22-gauge needles to isolate and remove the developing embryos. Caspase Assay Embryos were evaluated for the presence of cells undergoing apoptosis using a fluorescent based assay for the detection of active caspases-3 and -7 (Caspatag® Caspase 3,7 In Situ Assay Kit, Chemicon International, Inc.). Briefly, a 150X stock solution of Caspatag® reagent [a carboxyflouroscein-labeled flouromethyl ketone peptide inhibitor of caspases-3 and -7 (SR-DEVD-FMK)] was prepared by reconstituting 1 vial of Caspatag® reagent in 0.05 ml of dimethyl sulfoxide (DMSO, D128-500, Fischer Scientific, Fair Lawn, NJ). Embryos were then incubated in a 1500x Caspatag® working solution (1 µl 150x Caspatag® stock solution diluted in 1.5 ml of phosphate buffered
121 saline, protected from light, for 1 hour at 39°C under 5% CO2 in air). After the 1 hour incubation period, embryos were washed in 1x Caspatag® wash buffer (1ml of Caspatag® wash buffer diluted in 10 ml of deionized water) for 5 minutes and then stained in Hoerscht 33342 (1 mg/ml; bisBenzimide, B-2261, Sigma-Aldrich Inc., St. Louis, MO). After Hoerscht staining, embyos were placed on slides and the total number of apoptotic cells were counted using an inverted microscope (Nikon Diphot; Tokyo, Japan) equipped with a ultraviolet light and a rhodamine filter. Each embryo was counted three times and the mean number of apoptotic cells was determined. Embryos in which individual fluorescent points were not apparent were categorized as having profuse staining patterns and were assigned an arbitrary apoptotic cell value of 20. Apoptotic indices were calculated by dividing the mean number of apoptotic cells by the total number of cells. The total number of cells for in vivo- and in vitro-derived embryos were previously reported by Knijn et al. (2003; in vivo morulae and early blastocysts = 128 cells; in vivo blastocysts = 166; in vitro morulae and early blastocysts = 113; in vitro blastocysts = 114). These number of cells per embryo were in agreement with those reported by Van De Velde et al. (1999) and Koo et al. (2002). Positive controls were prepared by incubating in vitro-produced embryos in cyclohexamide (1 mg/ml; C-7698; Sigma-Aldrich Inc., St. Louis, MO) for 3 to 4 hours prior to staining. Statistical Analysis Data are expressed as means and were analyzed using PROC GLM of the SAS Statistical Package (v. 8e, 1999 SAS Institute, Inc., Gary, NC). Significant differences among means for number of apoptotic cells and apoptotic index were tested using a post hoc least significant difference test (LSD test). Differences with a probability (P) value of 0.05 or less were considered significant in this study. RESULTS Estrous synchronization and superovulation response is summarized in Table 8.1. Of the 18 females receiving CIDRs, 17 (94%) were administered GnRH at the time of CIDR removal. One female was removed from the study at this time due to a lost CIDR. Of the females remaining, 17 (100%) were determined to have a CL, as
122
Table 8.1. Response of cows to synchronization/superovulation protocol
No. cows receiving CIDRs 18
No. cows receiving GnRH 17
No. cows with a CL at initiation of FSH 17
No. cows observed in heat after FSH and PGF2α 13
No. cows inseminated 17
No. cows nonsurgically collected 13
123 determined by ultrasound (Aloka, SSD 500V, Japan) evaluation, at the initiation of FSH treatment. Following PGF2α administration, 13 of these females were detected in estrus and were inseminated 12 and 24 hours following observation of standing estrus. Four females did not display any sign of behavioral estrus and were inseminated at 60 and 72 hours after the last PGF2α injection. Following insemination, five females were collected nonsurgically 6.5 days post-insemination and 8 females were collected nonsurgically 7 days post-insemination. These time points were selected to ensure both morula and blastocyst stage embryos were harvested. Of the four females not collected, two were found to have uterine infections, one was diagnosed with a cystic follicle without a CL present, and one did not have a CL present on either ovary. There were 62 embryos recovered from 146 detectable CL (42% recovery rate) in 13 females (Table 8.2). Of the 62 embryos recovered, 28 were morulae, 13 were blastocysts and 21 were degenerate. Of the 62 total embryos recovered, 38 (61%) were classified as quality grade 1 or grade 2 embryos. The mean number of apoptotic cells in morula, early blastocyst and blastocyst stage embryos are presented in Table 8.3. Overall, there was no effect (P>0.05) of treatment on the mean number of apoptotic cells in morulae and early blastocysts. When blastocyst stage embryos were evaluated there was a significant increase in the number of apoptotic cells in frozen-thawed in vivo embryos when compared with fresh in vivo embryos. There was no difference (P>0.05) detected between fresh in vivo embryos, in vitro-cultured embryos and chick-cultured embryos. When only those embryos exhibiting specific staining patterns were evaluated (Table 8.4), there were fewer apoptotic cells in fresh in vivo morulae than in frozen-thawed in vivo morulae (P<0.05); morulae resulting from in vitro culture did not differ from fresh in vivo morulae. There was no effect of treatment on mean number of apoptotic cells in early blastocyst and blastocyst stage embryos (P>0.05). The apoptotic indices of embryos are presented in Table 8.5. There was no effect (P>0.05) of treatment on apoptotic index in morula and blastocyst stage embryos. The apoptotic indices of in vitro-cultured and chick-cultured early blastocysts were similar to those of fresh in vivo embryos. No differences were detected (P>0.05), however, between in vitro culture, chick culture and positive controls.
124
Table 8.2. Number, stage and quality grade of embryos recovered from superovulated cows
a MORL = morulae. b BLST = blastocysts. c G1 = quality grade 1. d G2 = quality grade 2. e G3-G4 = quality grade 3 and quality grade 4.
125
Table 8.3. Effect of culture environment on mean number of apoptotic cells in morula, early blastocyst and blastocyst stage embryos Mean no. of apoptotic cellsab
No. of Early Treatment group embryos Morulae blastocysts Blastocysts Degenerate d In vivo embryos 32 10.1d 1.0d 0.9d 9.5
Frozen-thawed in vivo embryos 32 8.9d na 8.2e na
In vitro-cultured embryos 64c 10.3d 8.9de 3.8de 13.6de
de de de d Chick-cultured embryos 21 13.3 0.0 6.7 19.0
Positive control 21 20.0e 15.0e 18.0f na
a Includes both specific and profuse patterned embryos. b na, not applicable. No embryos were available for evaluation in these categories. c Both in vitro culture groups were combined for statistical evaluation. d,e, f Within a column, values with different superscripts are significantly different (P<0.05).
126 Table 8.4. Effect of culture environment on mean number of apoptotic cells in morula, early blastocyst and blastocyst stage embryos ab Mean no. of apoptotic cells No. of Early Treatment group embryos Morulae blastocysts Blastocysts Degenerate In vivo embryos d Specific 23 1.5d 1.0d 0.9d 1.7 Profuse 9 20.0 na na 20.0
Frozen-thawed in vivo embryos Specific e d 21 3.4 na 1.5 na Profuse 11 20.0 na 20.0 na
In vitro-cultured embryos Specific 43c 0.5d 0.6d 1.2d 3.1de
Profuse 21 20.0 20.0 20.0 20.0
Chick-cultured embryos Specific de d d e 5 0.0 0.0 0.0 5.7 Profuse 16 20.0 na 20.0 20.0
Positive control Specific 2 na 0.0d 0.0d na
Profuse 19 20.0 20.0 20.0 na
a Embryos were categorized as having either specific or profuse staining patterns. In embryos exhibiting specific staining patterns, individual fluorescent points were apparent and could be quantified. Individual fluorescent points were not apparent in embryos exhibiting profuse staining patterns. These embryos were assigned an arbitrary apoptotic cell value of 20. b na, not applicable. No embryos were available for evaluation in these categories c Both in vitro culture groups were combined for statistical evaluation. d,e Within a column, values with different superscripts are significantly different (P<0.05).
127
Table 8.5. Effect of culture environment on apoptotic index in morula, early blastocyst, and blastocyst stage embryos Apoptotic indexabcde Treatment group No. Morulae Early blastocysts Blastocysts
In vivo embryos 25 7.9g 0.8g 0.5g
Frozen-thawed in vivo embryos 32 7.0g na 5.0g
f g gh g In vitro-cultured embryos 56 9.1 7.8 3.8
Chick-cultured embryos 7 11.8gh 0.0gh 5.8g
Positive control 21 17.7h 13.3h 15.8h
a Apoptotic index = mean number of apoptotic cells/total number of cells in the embryo. Total number of cells for in vivo- derived morulae and early blastocysts = 128; for in vivo-derived blastocysts = 166; for in vitro-produced morulae and early blastocysts = 113; for in vitro-produced blastocysts = 114 (Knijn et al., 2003). b Embryos were categorized as having either specific or profuse staining patterns. In embryos exhibiting specific staining patterns, individual fluorescent points were apparent and could be quantified. Individual fluorescent points were not apparent in embryos exhibiting profuse staining patterns. These embryos were assigned an arbitrary apoptotic cell value of 20. c Includes both specific and profuse patterned embryos. d Apoptotic index could not be calculated for degenerate embryos because embryo stage could not always be determined. e na, not applicable. No embryos were available for evaluation in these categories f Both in vitro culture groups were combined for statistical evaluation. g,h Within a column, values with different superscripts are significantly different (P<0.05).
128 The mean number of apoptotic cells per embryos in relation to quality grade are presented in Table 8.6. There was no effect (P>0.05) of treatment on the mean number of apoptotic cells in grade 1 or grade 2 embryos (morulae, early blastocysts and blastocysts). For grade 3 embryos, there were significantly more (P<0.05) apoptotic cells in chick-cultured embryos than in in vitro-cultured embryos; however, when only those embryos exhibiting specific staining patterns were evaluated there was no treatment effect (P>0.05) (Figure 8.7). For grade 4 embryos, frozen-thawed in vivo embryos had significantly more (P<0.05) apoptotic cells than fresh in vivo embryos; in vitro-cultured grade 4 embryos did not differ (P>0.05) from either treatment group. As with the mean number of apoptotic cells, there was no effect (P>0.05) of treatment on apoptotic indices of grade 1 or grade 2 embryos; similar results were observed for grade 4 embryos (Table 8.8). For grade 3 embryos, chick cultured-embryos had a greater apoptotic index than in vitro-cultured embryos; however, neither treatment was significantly different (P>0.05) from either fresh or frozen-thawed in vivo-derived embryos. DISCUSSION Dead cells were first observed in mammalian embryos in 1967 by Potts and Wilson. Soon thereafter it was determined that cells die by apoptosis (Kerr et al., 1972), a self directed form of cell suicide (Hardy, 1997; Betts and King, 2001). Since that time, apoptosis has been determined to be a critical process during pre-implantation development for the disposal of excess cells, cells that are in the way, abnormal or potentially dangerous (Gjorret et al., 2003). A normal pattern of apoptosis is critical for continued embryonic development (Brill et al., 1999), and any deviation in this pattern can compromise future development, often resulting in early embryonic death or fetal abnormalities (Knijn et al., 2003). There are several mechanisms by which apoptosis can be induced in cells, but regardless of the type of induction, receptor binding initiates a cascade of events that results in activation of an effector caspase (generally caspase-3 or -7) (Lawen, 2003). Effector caspases cleave cellular proteins and are responsible for the typical morphological changes observed in cells undergoing apoptosis, such as DNA fragmentation (Lawen, 2003). To our knowledge, all studies to date evaluating apoptosis in pre-implantation mammalian embryos utilize the TUNEL (terminal deoxynucleotidyl transferase mediated
129
Table 8.6. Effect of culture environment on mean number of apoptotic cells in grade 1, grade 2, grade 3, grade 4 and degenerate embryos
ab Embryo quality grades
Treatment group No. Grade 1 Grade 2 Grade 3 Grade 4 Degenerate
In vivo embryos 32 6.5dx 2.3dx 10.0dex 0.3dx 9.5dx
dx dx dex ey Frozen-thawed in vivo embryos 32 6.8 4.7 10.0 18.2 na
In vitro-cultured embryos 64c 6.2dx 6.0dx 6.7dxy 20.0dexy 13.6dey
Chick-cultured embryos 21 0.0dx 8.9dxy 20.0ey na 19.0ey
a Includes both specific and profuse patterned embryos. b na, not applicable. No embryos were available for evaluation in these categories (P<0.05). c Both in vitro culture groups were combined for statistical evaluation. d,e Within a column, values with different superscripts are significantly different (P<0.05). x,y Within a row, values with different superscripts are significantly different (P<0.05).
130
Table 8.7. Effect of culture environment on mean number of apoptotic cells in grade 1, grade 2, grade 3, grade 4 and degenerate embryos Embryo quality gradesab Treatment group No. Grade 1 Grade 2 Grade 3 Grade 4 Degenerate In vivo embryos dx dfxy dxy dx dy Specific 23 0.8 2.3 0.0 0.3 1.7 Profuse 9 20.0 na na na 20.0
Frozen-thawed in vivo embryos Specific 21 1.8dx 3.3dy 0.0dx 7.3ez na
Profuse 11 20.0 20.0 20.0 20.0 na
In vitro-cultured embryos Specific 43c 1.2dx 0.7efx 1.0dx na 3.1dey Profuse 21 20.0 20.0 20.0 20.0 20.0
Chick-cultured embryos Specific 5 0.0dx 0.0fx na na 5.7ey Profuse na 20.0 20.0 na 20.0 16 a Embryos were categorized as having either specific or profuse staining patterns. In embryos exhibiting specific staining patterns, individual fluorescent points were apparent and could be quantified. Individual fluorescent points were not apparent in embryos exhibiting profuse staining patterns. These embryos were assigned an arbitrary apoptotic cell value of 20. b na, not applicable. No embryos were available for evaluation in these categories. c Both in vitro culture groups were combined for statistical evaluation. d,e,f Within a column, values with different superscripts are significantly different (P<0.05). x,y,z Within a row, values with different superscripts are significantly different (P<0.05).
131
Table 8.8. Effect of culture environment on apoptotic index in grade 1, grade 2, grade 3 and grade 4 embryos abcde Apoptotic index Treatment group No. Grade 1 Grade 2 Grade 3 Grade 4
In vivo embryos 25 5.0gx 1.7gx 7.8fgx 0.3gx
Frozen-thawed in vivo embryos 29 4.3gx 3.5gx 6.0fgxy 13.2gy
In vitro-cultured embryos 28f 5.4gx 5.2gx 6.7fx 17.7gx
xxy gy Chick-cultured embryos 7 0.0gx 8.9 17.6 na a Apoptotic index = mean number of apoptotic cells/total number of cells in the embryo. Total number of cells for in vivo- derived morulae and early blastocysts = 128; for in vivo-derived blastocysts = 166; for in vitro-produced morulae and early blastocysts = 113; for in vitro-produced blastocysts = 114 Knijn et al., 2003). b Embryos were categorized as having either specific or profuse staining patterns. In embryos exhibiting specific staining patterns, individual fluorescent points were apparent and could be quantified. Individual fluorescent points were not apparent in embryos exhibiting profuse staining patterns. These embryos were assigned an arbitrary apoptotic cell value of 20. c Includes both specific and profuse patterned embryos. d Apoptotic index could not be calculated for degenerate embryos because embryo stage could not always be determined. e na, not applicable. No embryos were available for evaluation in these categories. f Both in vitro culture groups were combined for statistical evaluation. g,h Within a column, values with different superscripts are significantly different (P<0.05). x,y Within a row, values with different superscripts are significantly different (P<0.05).
132 dUTP nick end labeling) technique to label fragmented DNA (mice - Wuu et al., 1999; Brison and Schultz, 1997; rats - Pampfer et al., 1997; Hinck et al., 2001; pigs – Long et al., 1998; cattle - Byrne et al., 1999; Matwee et al., 2000; Makarevich and Markkula, 2002; Kolle et al., 2002; Neuber et al., 2002; Paula-Lopes and Hansen, 2002; humans – Yang et al., 1998; Hardy, 1999). However, DNA fragmentation is not an event unique to apoptosis as cells undergoing necrosis (a second form of cell death) also exhibit substantial DNA fragmentation (Grasl-Kraupp et al., 1995). In addition, it has been reported that improper tissue handling is sufficient to induce enough DNA fragmentation for TUNEL positive staining of nuclei (Negoescu et al., 1996).Thus there is a critical need for evaluation of different markers of apoptotic cell death for adequate determination of apoptosis in pre-implantation embryos. The objective of the present study was to evaluate the presence of caspases-3 and -7 as indicators of apoptosis in bovine embryos fertilized and cultured under different conditions. Caspase-3 mRNA and protein have been detected in mouse and rat blastocysts (Hinck et al., 2001). However, this is believed to be the first report of caspase activity in bovine embryos. In the present study, more than 65% of all embryos displayed at least one fluorescent cell, regardless of treatment (data not shown). These findings are consistent with those of previous studies (Byrne et al., 1999; Matwee et al., 2000; Neuber et al., 2002; Gjorret et al., 2003) and suggest that apoptosis is a normal process during early embryo development in cattle. Overall, culture conditions did not dramatically affect the incidence of apoptosis in bovine embryos in the current research. Frozen-thawed in vivo embryos did have significantly more apoptotic cells than did their fresh counterparts; however, when only those embryos exhibiting specific staining patterns were evaluated, this statistical difference was no longer present. It is somewhat unclear why some embryos exhibit specific staining patterns while others exhibit profuse staining patterns, but we hypothesize that profuse staining embryos are likely degenerate or degenerating. There is conflicting information in the literature regarding the effect of in vitro culture on apoptosis. Byrne et al. (1999) reported that the extent of apoptosis in embryos was affected by in vitro production systems; however, they did not evaluate in vivo- derived embryos in this study. Based on TUNEL analysis, Knijn et al. (2003) reported
133 that apoptotic index tended to be higher in in vitro-produced embryos than in embryos produced in vivo. Similarly, Gjorret et al. (2003) reported that the incidence of apoptotic cells was higher in blastocysts cultured in vitro than their in vivo-derived counterparts when analyzed with TUNEL. They contributed this difference specifically to higher levels of apoptosis in the inner cell mass (ICM) of in vitro-produced embryos. Although we were not able to specifically localize apoptotic cells to the ICM or trophoectoderm, there were consistently more caspase-positive (flourescent) cells in the region of the ICM, regardless of culture system (Figure 8.4). These findings are in agreement with previous reports indicating that apoptosis predominates in the ICM (El- Shershaby and Hinchliffe, 1974; Copp, 1978; Mohr and Trouson, 1982; Handyside and Hunter, 1986; Hardy and Handyside, 1996), with much lower levels detected in the trophoectoderm (Copp, 1978; Hardy and Handyside, 1996; Brison and Schultz, 1997). It has been hypothesized that apoptosis occurs naturally in vivo as a means to help eliminate ICM cells that still have the potential to form trophoectoderm, thus decreasing the chance of inappropriate trophoectoderm expression during germ layer differentiation (Handyside and Hunter, 1986; Pierce et al., 1989). In addition, Gjorret et al. (2003) suggested that the aberrations in apoptosis in the ICM could result in morphologically normal appearing blastocysts carrying subcellular deviations that could affect the developmental competence of the embryo. One interesting observation was the presence of different degrees of fluorescence within an embryo. Often, small, pinpoint areas of fluorescence were detected while other times the fluorescence appeared to consume the entire cytoplasm of a cell (Figure 8.5). The capase protein has been localized to the cytoplasm of the cell, therefore we hypothesize that the different degrees of fluorescence represent different degrees of apoptosis. Those cells with small pinpoint areas of fluorescence are likely in the very early stages of apoptosis with only small amounts of caspase present in the cytoplasm. In contrast, the instances in which the entire cell appears to fluoresce likely represent a more advanced stage of apoptosis with greater amounts of caspase present. In the present study, there was no major effect of in vitro culture environment on the incidence of apoptosis when embryos were categorized by quality grade. Moreover, within a treatment, there was no significant difference in apoptosis across quality grades. This is believed to be the first report of apoptosis evaluated in relation to embryo quality
134
Figure 8.4. Localization of apoptosis staining to the ICM of a grade 2 frozen in vivo- derived bovine blastocyst.
135
A
B
Figure 8.5. The presence of different degrees of fluorescence within an embryo. Pinpoint areas of fluorescence are thought to represent cells in the early stages of apoptosis with only small amounts of caspase present in the cytoplasm (A), whereas extensive fluorescence are thought to represent cells in more advanced stages of apoptosis with much more caspase present in the cell (B).
136 grade. Based on observations in the present study, it is likely that apoptosis is a naturally occurring event in embryo development and is not necessarily a reliable indicator of embryo viability or quality grade. The value of morphological evaluation as an indicator of embryo viability and quality has been contested for many years. In the early 1980s, Shea (1981) reported that embryos showing obvious signs of degeneration resulted in lower pregnancy rates than embryos with a normal morphology; however, these embryos were still able to produce pregnancies in many instances. It was further speculated that the extent of embryo development was a better indicator of embryo quality than morphological assessments. In the present study, some morphologically high quality embryos were observed to have multiple apoptotic cells while some presumptive degenerate embryos were observed to have no apoptotic cells. These observations add support to Shea’s hypothesis that morphological evaluation is not an optimal indicator of embryo viability and quality. In summary, the majority of embryos evaluated had more than one apoptotic cell, suggesting that apoptosis is a normal event in bovine embryo development both in vivo and in vitro. There was no apparent effect of culture system on the incidence of apoptosis when evaluated by embryo stage or quality grade. Based on the findings presented in this report, apoptosis does not appear to be heavily involved in the reduced developmental potential of in vitro-derived bovine embryos.
137 CHAPTER IX
SUMMARY AND CONCLUSIONS The primary purpose of this series of experiments was to evaluate the use of the chick embryo co-culture system in bovine embryo culture. In the early 1990s, this system was shown to improve development of pre-implantation embryos from various mammalian species when compared with the medium used at that time. Since that time, in vitro embryo culture media have progressed, but many problems still exist (e.g., poor developmental rates, poor embryo quality, fetal and neonatal abnormalities). These problems led us to revisit the use of the chick embryo co-culture system as an alternative to in vitro embryo culture. In the initial experiment, development of bovine 1-cell and 2-cell embryos produced by IVF was assessed after culture in the chick embryo amnion. Results indicated that bovine embryos could develop in the embryotropic environment of the chick embryo amnion; however, development was not significantly different when compared with the semi-defined culture medium (CR1aa). It was concluded that although not significantly better, the chick embryo co-culture system could serve as a viable alternative to more traditional methods of in vitro culture. In the second experiment of this series, development of bovine 8-cell embryos derived from somatic cell nuclear transfer was assessed after culture in the chick embryo amnion. Results indicated that these embryos are capable of developing into transferable embryos and implanting at a rate comparable with embryos culture in vitro in a semi-defined medium (CR1aa). The third experiment was conducted in an effort to improve the chick embryo co- culture system. To accomplish this, angiogenesis was examined in chick chorioallantoic membranes treated with a vasodilator (PGE2) and a vasoconstrictor (PGF2α). Based on the findings of the first part of this experiment, the development of bovine embryos after co-culture in PGE2-treated chicks was evaluated. It was concluded that increasing blood flow to the chick choriallantoic membrane did not appear to improve development or quality of bovine embryos cultured in untreated chicks over embryos cultured in semi- defined medium (control). These findings suggest that the benefits of the chick co- culture system lie in the ability of the chick to actively regulate its own environment rather than in embryotropic properties within the amniotic environment.
138 Because of the inherent mortality rate of shell-less chick embryos a fourth experiment was conducted to determine if amniotic fluid extracted from a developing chick embryo could be used as a supplement to in vitro culture medium and support development of bovine embryos. It was concluded that chick amniotic fluid could provide the proteins and growth factors needed to support in vitro development of bovine embryos, thus providing a feasible alternative to the use of fetal bovine serum in bovine embryo culture. In the fifth experiment of this series, protocols for culturing fetal chick cells and chick amnion cells were developed. This research provides an in vitro model for future studies into the effects of chick embryo co-culture on mammalian embryo development. In the last experiment, apoptosis was evaluated in in vivo-derived embryos, in vitro-produced embryos and embryos cultured in the chick amnion. This experiment utilized a novel approach to evaluate the incidence of apoptosis in embryos. Findings of this study indicate that apoptosis is a normal event in bovine embryo development both in vivo and in vitro and does not appear to be affected by the in vitro culture environment. Based on these findings, it was concluded that apoptosis is not extensively involved in the reduced developmental potential of in vitro derived bovine embryos. In conclusion, it was demonstrated that bovine embryos could develop in the amniotic cavity of a developing chick embryo at rates similar to those of embryos cultured in current culture media. Although the chick embryo co-culture system does not surpass current in vitro culture systems, it should not be completely dismissed. Pregnancy and calving rates resulting from current in vitro embryo production systems are significantly lower than those achieved with artificial insemination and embryo transfer. Moreover, numerous fetal and calf abnormalities have been associated with in vitro embryo production, particularly with the use of serum. If the chick embryo culture system could produce even one more viable offspring, free from developmental abnormalities, then its use may be warranted and significant research gains could be obtained.
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169
APPENDIX A
BO-A STOCK SOLUTION1
Component Product number Company Amount
NaCl S-5886 Sigma 4.3092 g
KCl P-5405 Sigma 0.1974 g
CaCl2•2H2O C-7902 Sigma 0.2171 g
NaH2PO4•H2O S-9638 Sigma 0.0743 g
MgCl2•6H2O M-2393 Sigma 0.0697 g
Milli Q water – Milli Q 500 ml
Phenol red 0.5% P-0290 Sigma 0.1 ml
1 Prepare solution in a 500-ml bottle and store in a refrigerator for up to 3 months.
170
APPENDIX B
BO-B STOCK SOLUTION1
Component Product number Company Amount
NaHCO3 S-8875 Sigma 2.5873 g
Milli Q water – Milli Q 200 ml
Phenol red 0.5% P-0290 Sigma 0.04 ml
1 Prepare solution in a 250-ml bottle. Inject CO2 gas into the bottle for 1 to 2 minutes until a color change occurs. Seal the bottle with parafilm to maintain proper pH and store in a refrigerator for up to 3 months.
171
APPENDIX C
CR1aa STOCK SOLUTION1
Component Product number Company Amount
NaCl S-5886 Sigma 0.6703 g
KCl P-5405 Sigma 0.0231 g
NaHCO3 S-8875 Sigma 0.2201 g
Pyruvic acid P-4562 Sigma 0.0044 g
L (+) Lactic acid L-4388 Sigma 0.0546 g
Glycine G-8790 Sigma 0.0039 g
L-Alanine A-7469 Sigma 0.0045 g
Milli Q water – Milli Q 500 ml
Phenol red 0.5% P-0290 Sigma 0.2 ml
1 Prepare solution in a 100-ml bottle and stored in a refrigerator for up to 3 months.
172 VITA
Tonya Renea Davidson was born on September 24, 1972 in Lufkin, Texas to James and Linda Davidson. Tonya has one younger brother, Casey Millspaugh, who resides in Fayetteville, Arkansas. Tonya grew up in Fort Smith, Arkansas where she attended Woods Elementary School and later attended Southside High School. Following graduation from high school in 1991, Tonya enrolled in Carl Albert State University, in Poteau, Oklahoma, completing an Associate of Science degree in May of 1993. She then transferred to Oklahoma State University in Stillwater, Oklahoma to pursue a Bachelor of Science degree in Animal Science. After receiving her Bachelor’s degree in December of 1997, Tonya entered the graduate program at Oklahoma State University under Dr. Leon J. Spicer. After completing her class work and research, Tonya received a Master’s of Science in December of 2000. She was then accepted to begin a doctoral degree at Louisiana State University under the guidance of Dr. Robert Godke, Boyd Professor of Reproductive Physiology. She is a candidate for the Doctor of Philosophy degree in reproductive physiology in the Department of Animal Sciences at Louisiana State University, Baton Rouge, Louisiana.
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