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LSU Doctoral Dissertations Graduate School

2004

The chick embryo amnion as an in vitro culturesystem for IVF and NT embryosTonya Renea DavidsonLouisiana State University and Agricultural and Mechanical College

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Recommended CitationDavidson, Tonya Renea, "The chick embryo amnion as an in vitro culture system for IVF and NT embryos" (2004). LSU DoctoralDissertations. 1280.https://digitalcommons.lsu.edu/gradschool_dissertations/1280

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 E2PGF2α - 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

Onset of Egg Incubation

Preparation of Shell-less Cultures

48 h

~24 h

Injection of CEC-I

IVF IVC

72 h

Recovery of CEC-I Injection of CEC-II

96 h

Recovery of CEC-IIEvaluation

Evaluation Addition of FBS

5 h 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

Onset of Egg Incubation

48 h ~24 h

Injection of CEC-I

IVF IVC

72 h

Recovery of CEC-I Injection of CEC-II

96 h

Recovery of CEC-I Evaluation

Evaluation Addition of FBS

Chick Culture Treatment Groups

Zygotes 18 h

Preparation of Shell-less Cultures

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

0102030405060708090

100

1-cell 2-4 cell 6-8 cell cleavage

% o

f Em

bryo

s

Control Agar CEC-I CEC-IIA

a

b

a a

a

b

aa

aaaa

a

b

a a

0

10

20

30

40

50

MORL Early BLST BLST ExpandedBLST

% o

f Em

bryo

s

Control Agar CEC-I CEC-II

a

a

a

a

a

a

a

aa

a

a

a a aaa

B

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)

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

Day 3 Day 7

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

0102030405060708090

100

2-4 cell 6-8 cell >8-cell

% o

f Em

bryo

s

Control Agar CEC-I CEC-IIA

a a aa

a

aaa

a

a

a a

0

10

20

30

40

50

MORL Early BLST BLST ExpandedBLST

Hatching orHatched

BLST

% o

f Em

bryo

s

Control Ag a r CEC-I CEC-II

B

ab

b

a

b

a

a

aa

a a a b

ab ab

aa a

a a a

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 development (A) and on day 7 of development (B). Within a developmental stage, columns without a common superscript are significantly different (P<0.05).

42

0

10

20

30

40

50

Grade 1 Grade 2 Grade 3 Grade 4

% o

f Em

bryo

s

Control Agar CEC-I CEC-II

a

a

a aab

abb

a

a

a

a

aa

a

a a

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

Injection of 8-cell Embryos into Chick Amnion

Culture in CR1aa + 5% FBS

72 hours

72 hours

Evaluation Recovery 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. (%) groupa culturec recovered % MORLfh % BLSTgh % MORL & BLSTfgh

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

Chick embryo co-culture 107d 54 (51)e 42.6f 17.2f 59.9f

Control culture medium 113 -- 54.2f 19.8f 74.1f

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

by removing albumin and creating a window

Return to incubation

Apply treatments and return to incubation

Incubate eggs at 37°C and 85 to 90% relative humidity for 96 hours

Quantify 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

64

Onset of Egg Incubation

IVF IVC

Injection of Bovine Embryos

Recovery Evaluation

Injection of Bovine Embryos

Recovery Evaluation

PGE Treatment Group

CEC Treatment Group Shipment of Presumptive

Zygotes

Preparation of Shell-less Cultures Application of PGE2

Preparation of Shell-less Cultures

Onset of E g Incubation g

~24 h 96 h18 h 24 h 24 h

Shipment of Presumptive

Zygotes

IVF IVC

~24 h 96 h18 h 48 h

IVC Addition of FBS Evaluation IVF

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).

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)

PGE2 7 4 (57) 3 (43)a

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

a,b,c Values without a common superscript are significantly different (P<0.05).

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

PGE2 28 24.1c 1.26

PGF2α 16 19.1bc 1.22

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

Control Cytochalasin EtOH PGE PGF0

5

10

15

20

25

30

c

bc

b

Num

ber o

f Nod

es

a a

n = 12 n = 4 n = 20 n = 28 n = 16

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

0

10

20

30

40

MORL EarlyBLST

BLST ExpandBLST

HatchingBLST

% o

f Em

bryo

s

ControlCECPGE

A

aa

a

bb

a

a

aa

a a

a

b

ab

a

0

10

20

30

40

MORL Early BLST Expanded Hatching Hatched

% o

f Em

bryo

BLST BLST BLST BLST

s

ControlCECPGE

a

a a a

a

aa

a

a

b

ab

aaa

aa

B

a

a

0

5

10

15

20

BLST ExpandBLST

HatchingBLST

HatchedBLST

% o

f Em

bryo

s

ControlCECPGE

C

a

aa

aa

a

a

a

a

bb

a

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

0

10

20

30

40

50

Grade 1 Grade 2 Grade 3 Grade 4

% o

f Em

bryo

s

ControlCECPGE

A

aa

aa

aa

aa

a

a

a

a

0

10

20

30

40

50

Grade 1 Grade 2 Grade 3 Grade 4

% o

f Em

bryo

s

ControlCECPGE

B

aa

aa

aa

aa

a

aa

a

0

10

20

30

40

50

Grade 1 Grade 2 Grade 3 Grade 4

% o

f Em

bryo

s

ControlCECPGE

C

aa

a

a

a

aa

a

a

a

b

a

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

CAF

Control

In Vitro Culture

In Vitro Fertilization

96 hours

72 hours

5 hours

18 to 22 hours

Media Change Embryo Evaluation

Embryo Evaluation

24 hours

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

0

10

20

30

40

50

MORL Early BLST BLST Expand BLST

% o

f Em

bryo

s

ControlCAF

a

b

a a

a

b

a

b

A

0

10

20

30

40

50

MORL Early BLST BLST ExpandedBLST

HatchingBLST

HatchedBLST

% o

f Em

bryo

s

B ControlCAF

a a a

aaa a a

a baa

0

10

20

30

40

50

BLST ExpandBLST

HatchingBLST

HatchedBLST

% o

f Em

bryo

s

ControlCAF

b

a

aa

aa

a a

C

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

0

10

20

30

40

50

Grade 1 Grade 2 Grade 3 Grade 4

No.

of E

mbr

yos

ControlCAF

% o

f Em

bryo

s

aa

a

a

a

a

aa

A

0

10

20

30

40

50

Grade 1 Grade 2 Grade 3 Grade 4

% o

f Em

bryo

s

ControlCAF

a

aa

a a

b aa

B

0

10

20

30

40

50

Grade 1 Grade 2 Grade 3 Grade 4

% o

f Em

bryo

s

ControlCAF

a a

a a ab a a

C

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 mTCM = modified Tissue Culture Medium. b mDMEM = modified Dulbecco’s Modified Eagle’s Medium.

Treatment A Bovine Fibroblasts mTCMa

Treatment B Bovine Fibroblasts mDMEMb

Treatment C Fetal Chick Cells mTCMa

Treatment D Fetal Chick Cells mDMEMb

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 mTCM = modified Tissue Culture Medium. b mDMEM = modified Dulbecco’s Modified Eagle’s Medium.

Treatment A Bovine Fibroblasts mTCMa

Treatment B Bovine Fibroblasts mDMEMb

Treatment C Chick Amnion Cells mTCMa

Treatment D Chick Amnion Cells mDMEMb

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

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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.

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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).

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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).

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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).

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0

1

2

3

4

5

1 2 3 4 5

Days of Growth

Popu

latio

n D

oubl

ings

Passage 1 Passage 2 Post-Thaw

Figure 7.1. Growth of chick amnion cells in vitro in mTCM culture medium.

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0

1

2

3

4

5

1 2 3 4 5

Days of Growth

Popu

latio

n D

oubl

ings

Passage 1 Passage 2 Post-Thaw

Figure 7.2. Growth of chick amnion cells in vitro in mDMEM culture medium.

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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.

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0

0.2

0.4

0.6

0.8

1

1 2 3 4 5Days of Growth

Cell

Dens

ity (x

106 )

Fibroblast Cells Epithelial Cells Total Cells

Figure 7.3. Cell populations of chick amnion cells cultured in vitro in mTCM during the first passage.

108

0

1

2

3

4

5

6

1 2 3 4 5Days of Growth

Cell

Dens

ity (x

106 )

Fibroblast Cells Epithelial Cells Total Cells

Figure 7.4. Cell populations of chick amnion cells cultured in vitro in mTCM during the second passage.

109

0

0.5

1

1.5

2

2.5

3

3.5

1 2 3 4 5Days of Growth

Cell

Dens

ity (x

106 )

Fibroblast Cells Epithelial Cells Total Cells

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.

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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

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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.

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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.

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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

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(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-

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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

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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

In vivo embryos Frozen-thawed in vivo embryos In vitro-cultured embryos Chick-cultured embryos Positive control

32

32

64c

21

21

10.1d

8.9d

10.3d

13.3de

20.0e

No. of Early Treatment group embryos Morulae blastocysts Blastocysts Degenerate

1.0d

na

8.9de

0.0de

15.0e

0.9d

8.2e

3.8de

6.7de

18.0f

9.5d

na

13.6de

19.0d

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

Mean no. of apoptotic cellsab

No. of Early Treatment group embryos Morulae blastocysts Blastocysts Degenerate

In vivo embryos Specific Profuse Frozen-thawed in vivo embryos Specific Profuse In vitro-cultured embryos Specific Profuse Chick-cultured embryos Specific Profuse Positive control Specific Profuse

23 9

2111

43c

21

516

219

1.0d

na

na

na

0.6d

20.0

0.0d

na

0.0d

20.0

1.7d

20.0

na

na

3.1de

20.0

5.7e

20.0

na na

1.5d

20.0

3.4e

20.0

0.5d

20.0

0.0de

20.0

na 20.0

0.9d

na

1.5d

20.0

1.2d

20.0

0.0d

20.0

0.0d

20.0

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

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).

In vivo embryos Frozen-thawed in vivo embryos In vitro-cultured embryos Chick-cultured embryos Positive control

25

32

56f

7

21

7.9g

7.0g

9.1g

11.8gh

17.7h

0.8g

na

7.8gh

0.0gh

13.3h

0.5g

5.0g

3.8g

5.8g

15.8h

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

a Includes both specific and profuse patterned embryos.

Embryo quality gradesab

Treatment group No. Grade 1 Grade 2 Grade 3 Grade 4 Degenerate

In vivo embryos Frozen-thawed in vivo embryos In vitro-cultured embryos Chick-cultured embryos

32

32

64c

21

6.5dx

6.8dx

6.2dx

0.0dx

9.5dx

na

13.6dey

19.0ey

0.3dx

18.2ey

20.0dexy

na

10.0dex

10.0dex

6.7dxy

20.0ey

2.3dx

4.7dx

6.0dx

8.9dxy

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 Specific Profuse Frozen-thawed in vivo embryos Specific Profuse In vitro-cultured embryos Specific Profuse Chick-cultured embryos Specific Profuse

239

2111

43c

21

5 16

0.8dx

20.0

1.8dx

20.0

1.2dx

20.0

0.0dx

na

2.3dfxy

na

3.3dy

20.0

0.7efx

20.0

0.0fx

20.0

0.0dxy

na

0.0dx

20.0

1.0dx

20.0

na 20.0

0.3dx

na

7.3ez

20.0

na 20.0

na na

1.7dy

20.0

na na

3.1dey

20.0

5.7ey

20.0

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

a Apoptotic index = mean number of apoptotic cells/total number of cells in the embryo. Total number of cells for in vivo-

Treatment group No. Grade 1 Grade 2 Grade 3 Grade 4

Apoptotic indexabcde

In vivo embryos Frozen-thawed in vivo embryos In vitro-cultured embryos Chick-cultured embryos

25

29

28f

7

5.0gx

4.3gx

5.4gx

0.0gx

0.3gx

13.2gy

17.7gx

na

7.8fgx

6.0fgxy

6.7fx

17.6gy

1.7gx

3.5gx

5.2gx

8.9xxy

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

B

A

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.

139

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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.

173