Reproductive Biology of the Female Bottlenose Dolphin (Tursiops Truncatus)

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Reproductive biology of the female bottlenose dolphin (Tursiops truncatus)

Holley Stone Muraco

A Dissertation Submitted to the Faculty of Mississippi State University in Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy in Animal Physiology in the Department of Animal and Dairy Science

Mississippi State, Mississippi

December 2015

Holley Stone Muraco

2015

Reproductive biology of the female bottlenose dolphin (Tursiops truncatus)

Holley Stone Muraco

Approved:

______Scott T. Willard (Major Professor/Graduate Coordinator)

______Peter L. Ryan (Committee Member)

______Jean M. N. Feugang (Committee Member)

______Ashli Brown-Johnson (Committee Member)

______George M. Hopper Dean College of Agriculture and Life Sciences

Name: Holley Stone Muraco

Date of Degree: December 11, 2015

Institution: Mississippi State University

Major Field: Animal Physiology

Major Professor: Scott Willard

Title of Study: Reproductive biology of the female bottlenose dolphin (Tursiops truncatus)

Pages in Study: 228

Candidate for Degree of Doctor of Philosophy

The goal of this long-term study was to better understand the reproductive biology

of the female bottlenose dolphin (Tursiops truncatus) and provide a hypothesis for how

dolphins may communicate reproductive readiness to one another. Utilizing conditioned

dolphins in aquaria, this dissertation examined several previously unknown aspects of

dolphin reproduction, including ovarian follicular dynamics during the luteinizing

hormone surge, urinary prolactin levels, estrus behavior, vaginal fluid arboriform

arrangement, in-situ vaginal and cervical anatomy during estrus, reversed-phase high-

performance liquid chromatography (RP-HPLC) of urine samples to identify proteins and

peptides that may be used in chemical communication, and a review and anatomical

analysis of dolphin vibrassal crypts. The diffusely seasonal dolphin estrous cycle is not

controlled by photoperiod and has a 10-day follicular and 20-day luteal phase. A brief

ovulatory LH surge is followed by ovulation within 48 hours. An ethogram of 20

reproductive behaviors was developed, and all occurrences of reproductive behavior were

analyzed during conceptive estrous cycles. A novel form of standing heat estrus, termed

immobility, was observed, and estrus dolphins displayed genital nuzzling, active and

passive mounting with other females, and an increase of standing heat intensity as LH levels rose. Prolactin plays a role in pregnancy maintenance, mammary development, allo-mothering behavior, lactation, and lactational anestrus. Dolphins are similar to sows where weaning causes a return to estrus, and in the boar effect, where days to ovulation are shortened in the presence of a mature male. Dolphin vaginal fluid showed crystallization arrangements with large open mesh patterns, conducive to sperm transport, during the estrogenic follicular phase, and closed mesh during the luteal phase. RP-HPLC analysis revealed that urine contained large amounts of peptides and proteins with peaks that change throughout the estrous cycle and with changes in social grouping. Remnant vibrissae from dolphin follicular crypts were sectioned, and it was hypothesized that trigeminal nerve endings could act similarly to those found in the nasal mucosa of terrestrial species and respond to chemical stimuli. This study provides new data to better understand the reproductive biology of a holaquatic mammal.

DEDICATION

I dedicate this dissertation to my husband, Mike; children, Jordan and Colton; parents, Mitch and Jeanette Stone; and sister, Emily Stone. This has been a long roller coaster ride, full of extreme highs and lows, and I can’t thank you enough for the support and encouragement. I also want to make a special dedication to my late Grandfather,

“Coach” Thomas Mitchell Stone Sr. who always believed in me and encouraged me to live life to the fullest.

I love animals, and I’ve had the privilege of being around them my entire life.

From helping my father and grandfather on the farm as a child in rural Mississippi; to working as an animal trainer in zoos and aquariums; to studying wild Pacific walrus in the Arctic, I have a deep appreciation, respect and passion for animals based on firsthand experience. I also love science and have learned to regard and abide by the scientific process as I pursued my Ph.D. I’ve published in scientific journals, spoken at countless conferences and worked as the lead scientist on a wildlife nature documentary. However, most important of all, I am a mom of two boys, and I’m desperately concerned about the future generation and how they view animals, nature and the environment.

Rarely has there been such a one-sided look into a subject than with the dolphin.

Ask almost anyone about the dolphin, and the answer will most likely be one of the following: they are really smart; they have brains bigger than humans; they are sentient beings. But outside of hearsay and popular belief, what do we really know? Is science

overwhelmingly supportive that dolphin brain size is due to intelligence or that they are

indeed the “smartest” animal?

I must admit, as a student who had spent all of my time working with horses,

cows, goats, pigs, birds, and various woodland creatures, the concept that there was a

very special animal species that somehow transcended “normal” animals was a very

romantic and appealing ideal to me. Enough so, that I packed up and left the family farm

and moved to Florida for an internship with dolphins. I will never forget the first time I

saw the two highly studied dolphins of which I would get to help take care of and learn

from for the next 6 months. As I walked by their pool during our orientation tour, they both rose up and watched as we walked by. Already with a pre-conceived belief in my mind that dolphins were extremely intelligent, I was sure that I just experienced a unique connection as I was “regarded” by these magnificent animals.

Almost 20 years later, I’ve had the opportunity to work with, care for and study well over 100 dolphins, as well as other cetaceans including killer whales and beluga whales. (Killer whales and dolphins are in the same family, Delphinidae, but the beluga whale is in the Monodontidae family and are related to narwhals.)

I’ve learned a lot since that first dolphin encounter, but to this day, I never get tired of walking into an aquarium and having dolphins gazing my way. I have had the privilege of sharing dolphins with the public, and I see the same look that I had long ago as they walk by the pools and the dolphins rise up and stare. There is nothing as wonderful as connecting and inspiring people with animals.

Dolphins aside, every day we are looked at, studied and regarded by a range of creatures such as wild birds, squirrels, dogs, cats, horses, cows, pet fish, hamsters, deer

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etc. but it doesn’t evoke the same feelings as being looked at by a dolphin. Why? Is it the dolphin’s sleek exotic appearance, the foreign concept of having an aquatic creature peek into our terrestrial world? Perhaps, it’s because we believe we are looking at a being much like ourselves?

So what have I learned about dolphins after working with, training, caring for, researching and observing dolphins and whales for almost 20 years? By far the most valuable thing they have taught me is that NO animal is any more special than the next, and that EVERY animal is special, unique and intelligent in their own way. I also learned that the study of “real” dolphin behavior, anatomy and physiology is more interesting than the emotionally biased anthropomorphized versions. There is still much we don’t know about dolphins (or many other animal species for that matter), so it’s important not to jump to conclusions. Always be mindful and use scientific principles when approaching any subject.

Dolphins are easy to anthropomorphize, which gives people the tendency to feel emotional empathy towards them. Some people may even believe that dolphins can’t live fulfilling lives at a zoo or aquarium based on their own human emotions and feelings.

However, it’s a fact that dolphins can live long, enriching and healthy lives in aquaria.

Every day dedicated professionals provide a fabulous home and life for dolphins and whales in aquariums and zoos, and the dolphins work as catalysts to inspire and teach the public about caring for the environment. Even more importantly, our dolphin companions readily provide researchers with critical access to behaviors, samples and data that otherwise would be impossible.

So I also dedicate this dissertation to the dolphins that call our zoos and aquariums home (especially to my best teachers in the world, “Goofy”, “Duchess” and

“Chelsea”), and to the professional managers, trainers and veterinarians that care for them.

“The important thing is to not stop questioning. Curiosity has its own reason for

existing.”

—Albert Einstein

“The greatest enemy of knowledge is not ignorance, it is the illusion of

knowledge.”

—Daniel J. Boorstin

“You have enemies? Good. That means you've stood up for something, sometime

in your life.”

—Winston Churchill

“What is now proved was once only imagined.”

—William Blake

ACKNOWLEDGEMENTS

This work would not be possible without the talented and dedicated animal trainers and dolphins who participated in these studies. Thanks to Dr. John Fuquay, my inspiring professor who introduced me to the fascinating world of animal reproduction.

Thanks to my major professor Scott Willard Ph.D. for not being afraid of taking on a non-traditional student and allowing me time to develop my research skills. Thanks to

Peter Ryan Ph.D. for your unyielding support, encouragement and assistance. Thanks to

Jean M. N. Feugang Ph.D. for all of your assistance and having the same “you never know until you try” mentality. Thanks to Ashli Brown-Johnson Ph.D. for providing me some insight into the fascinating world of pheromones. The following people are but a few of those who helped me get to this point with their unwavering support and putting up with my often incessant questions. Thanks to Stan Kuczaj Ph.D. who first introduced me to the world of dolphin research and behavior and inspired me to think inquisitively and critically. Thanks to Paul Turek MD, FACS, FRSM, for giving me confidence to pursue new ideas. Thanks to Thomas H. Reidarson DVM, Dipl. ACZM, for always believing in me and encouraging me to keep going. Thanks to Barbara Durrant Ph.D. for your kindness and willingness to help out a small town kid with big dreams, and providing assistance in the adaptation of the canine luteinizing hormone immunochromatographic assay for use in dolphin urine. Thanks to Dennis Arn, DVM for introducing me to the canine luteinizing hormone immunochromatographic assay and

being a wonderful mentor and collaborator during our early years of trying to figure out dolphin reproduction. Thanks to Walter Threlfall DVM, MS, Ph.D. DACT for always answering my most rudimentary questions about theriogenology. Thanks to Fiona Brook

Ph.D. for patiently looking at my ultrasound images of dolphin “intestinal follicles” and

gently steering me to where the actual ovaries lie. Thanks to Jay Sweeney DVM, Rae

Stone DVM, and Michelle Campbell for entrusting me with your amazing staff and

animals. Thanks to Pat Clough for being a friend and supporter. Thanks to Jan Roser

Ph.D. for developing and validating the dolphin urinary prolactin RIA used in this study.

Thanks to Scott Willard Ph.D. and Mississippi State University for loaning me an

ultrasound machine. Thanks to facilities Six Flags Discovery Kingdom, The Mirage

Dolphin Habitat, Dolphin Research Center and Theater of the Sea. Last but certainly not

least I can’t say thank you enough to my husband and best friend Mike. Thanks for being

“a researcher’s best friend” and keeping the path clear so that progress can be made. You

make the world a better place!

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TABLE OF CONTENTS

DEDICATION ...... ii

ACKNOWLEDGEMENTS ...... vi

LIST OF TABLES ...... xiii

LIST OF FIGURES ...... xiv

CHAPTER

I. INTRODUCTION AND LITERATURE REVIEW ...... 1

1.1 Introduction ...... 1 1.2 Endocrinology ...... 2 1.2.1 Estrogen synthesis and its role during follicular development and ovulation ...... 2 1.2.2 Estrogen in the bottlenose dolphin ...... 3 1.2.3 Progesterone and corpus luteum formation ...... 3 1.2.4 Progesterone in the bottlenose dolphin ...... 5 1.2.5 Follicle-stimulating hormone and luteinizing hormone ...... 6 1.2.6 FSH and LH in the bottlenose dolphin ...... 7 1.2.7 Prostaglandin ...... 7 1.2.8 Prostaglandin in the bottlenose dolphin ...... 8 1.3 Dolphin estrous cycle ...... 8 1.4 Dolphin Reproductive Anatomy and Behavior ...... 9 1.4.1 Reproductive behavior ...... 9 1.4.2 Pregnancy and Lactation ...... 10 1.4.3 Artificial Insemination ...... 11 1.4.3.1 Introduction ...... 11 1.4.3.2 Sperm Collection and Cryopreservation ...... 13 1.4.3.3 Estrous Cycle Manipulation, Induction and Synchronization ...... 21 1.4.3.4 Artificial Insemination ...... 26 1.5 Discussion ...... 30

II. OVARIAN FOLLICULAR DYNAMICS DURING THE LUTEINIZING HORMONE SURGE IN THE BOTTLENOSE DOLPHIN (TURSIOPS TRUNCATUS) ...... 31

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2.1 Introduction ...... 31 2.2 Method ...... 33 2.2.1 Experimental ethics ...... 33 2.2.1.1 Animals ...... 33 2.2.1.2 Statistics ...... 35 2.2.1.3 Behavioral Conditioning ...... 35 2.2.1.4 Ultrasonography ...... 35 2.2.1.5 Urine Collection and Hormone Assays ...... 36 2.2.1.6 Estrous Cycle Monitoring ...... 38 2.3 Results ...... 39 2.4 Discussion ...... 40

III. URINARY PROLACTIN CONCENTRATIONS IN THE FEMALE BOTTLENOSE DOLPHIN (TURSIOPS TRUNCATUS) ...... 48

3.1 Introduction ...... 48 3.2 Method ...... 52 3.2.1 Experimental ethics ...... 52 3.2.2 Animals ...... 52 3.2.3 Statistics ...... 53 3.2.4 Behavioral conditioning ...... 53 3.2.5 Urine collection ...... 54 3.2.6 Hormone testing ...... 55 3.2.6.1 Progesterone and estradiol ...... 55 3.2.6.2 Prolactin ...... 55 3.2.6.3 Creatine (Cr) ...... 56 3.2.7 Ultrasonography ...... 56 1.1.1.1 Anestrus ...... 57 3.2.7.2 Seasonality and prolactin ...... 57 3.2.7.3 Estrous ...... 58 3.2.8 Pregnancy ...... 58 3.2.9 Lactation ...... 58 3.2.10 Cohabitation with mother and calf pairs ...... 58 3.3 Results ...... 59 1.1.1 Anestrus ...... 62 3.3.2 Seasonality and prolactin ...... 62 3.3.3 Estrous cycle ...... 62 3.3.4 Follicular phase ...... 63 3.3.5 Luteal phase ...... 66 3.3.6 Estradiol ...... 66 3.3.7 Pregnancy ...... 68 3.3.8 Lactation ...... 69 3.3.8.1 Post-partum ...... 69 3.3.8.2 Nursing durations ...... 70 3.3.8.3 Lactation in non-mothers ...... 70 3.3.9 Cohabitation ...... 71 ix

3.3.10 Mammary gland ultrasound ...... 73 3.4 Discussion ...... 76 3.4.1 Pregnancy ...... 76 3.4.2 Lactation ...... 77 3.4.3 Cohabitation ...... 79 3.4.4 Estrous Cycle ...... 80 3.4.5 Anestrus ...... 82 3.5 Conclusion ...... 83

IV. CONCEPTIVE ESTRUS BEHAVIOR IN THREE BOTTLENOSE DOLPHINS (TURSIOPS TRUNCATUS) ...... 84

4.1 Introduction ...... 84 4.2 Method ...... 86 4.2.1 Experimental ethics ...... 86 4.2.2 Animals ...... 87 1.1.1.1 Statistics ...... 87 4.2.2.2 Behavioral Conditioning ...... 88 4.2.2.3 Urine Collection ...... 88 4.2.2.4 Ultrasonography ...... 88 4.2.2.5 Hormone Testing Progesterone, Estradiol and Lutenizing Hormone ...... 89 4.2.2.6 Social Grouping ...... 90 4.2.2.7 Behavioral Recording and Analysis ...... 90 4.2.2.8 Proof of Ejaculation via the Identification of Spermatozoa in Vaginal Fluid ...... 91 4.3 Results ...... 92 4.3.1 Ultrasonography ...... 92 4.3.2 Endocrinology ...... 93 4.3.3 Reproductive behavioral ethogram ...... 94 4.3.4 Reproductive State ...... 95 4.3.5 All-occurrences of reproductive behavior ...... 96 1.1.1.1 Anestrus ...... 98 4.3.5.2 Socio-Sexual ...... 98 4.3.5.3 Estrus ...... 98 4.3.6 Passive behaviors towards animal 1 (Figure5) ...... 99 4.3.6.1 Anestrus ...... 99 4.3.6.2 Socio-sexual ...... 99 4.3.6.3 Estrus ...... 99 4.3.7 Behaviors enacted by and towards the focal nulliparous females (animals 6 and 7) during estrus ...... 100 4.3.7.1 Active reproductive behaviors by animals 6 and 7 ...... 100 4.3.7.2 Passive reproductive behaviors by animals 6 and 7 ...... 101 4.3.8 Behaviors from animal 9 towards animal 7 included push (1) ...... 101 4.3.8.1 Immobility ...... 101 4.3.9 Behavioral Correlations to Endocrinology ...... 103 x

4.3.9.1 Estradiol ...... 103 4.3.9.2 Luteinizing hormone (LH) ...... 103 4.3.10 Copulation observations and spermatozoa in vaginal fluid ...... 105 4.3.11 Mature male (animal 5) display ...... 105 4.4 Discussion ...... 106

V. CHANGES IN A BOTTLENOSE DOLPHIN (TURSIOPS TRUNCATUS) VAGINAL FLUID ARBORIFORM ARRANGEMENT DURING ANESTRUS AND THE ESTROUS CYCLE ...... 115

5.1 Introduction ...... 115 5.2 Method ...... 117 5.2.1 Experimental ethics ...... 117 5.2.2 Study animals ...... 117 1.1.1.1 Behavioral Conditioning ...... 118 5.2.2.2 Vaginal Endoscopy ...... 118 5.2.2.3 Vaginal fluid collection and analysis ...... 118 5.2.2.4 Hormone Testing Progesterone, Estradiol and LH ...... 119 5.2.2.5 Ultrasonography ...... 120 5.3 Results ...... 120 5.3.1 Vaginal fluid arboriform (ferning) arrangement ...... 120 5.3.2 Endocrinology ...... 126 5.3.3 Ultrasonography ...... 126 5.3.4 Vaginal Endoscopy ...... 127 5.4 Discussion ...... 130

VI. C18 REVERSED-PHASE HIGH-PERFORMANCE LIQUID CHROMATOGRAPHY (RP-HPLC) OF BOTTLENOSE DOLPHIN (TURSIOPS TRUNCATUS) URINE TO IDENTIFY POTENTIAL CANDIDATE PROTEIN AND/OR PEPTIDE PHEROMONES ...... 132

6.1 Introduction ...... 132 6.2 Method ...... 138 6.2.1 Experimental ethics ...... 138 6.2.2 Animals ...... 138 1.1.1.1 Behavioral Conditioning ...... 139 6.2.2.2 Urine Collection ...... 139 6.2.2.3 Ultrasonography ...... 140 6.2.2.4 RP-HPLC Analysis ...... 140 6.2.2.5 Amino Acid Sequencing ...... 141 6.2.2.6 Progesterone, Estradiol and Lutenizing Hormone ...... 141 6.2.2.7 Identification of spermatozoa in vaginal fluid ...... 142 6.2.2.8 Social Grouping ...... 142 6.3 Results ...... 143 6.3.1 Ultrasonography ...... 143 6.3.2 Endocrinology ...... 144 xi

6.3.3 Spermatozoa in vaginal fluid ...... 145 6.3.4 RP-HPLC ...... 145 6.3.5 Amino-terminal (N-terminal) sequence analysis ...... 158 6.4 Discussion ...... 160

VII. VIBRISSAL CRYPTS OF THE BOTTLENOSE DOLPHIN (TURSIOPS TRUNCATUS) ...... 166

7.1 Introduction ...... 166 7.2 Method ...... 168 7.2.1 Experimental ethics ...... 168 7.2.2 Animals ...... 169 1.1.1.1 Vibrissae ...... 169 7.2.3 Results ...... 170 7.2.3.1 Vibrissal hairs and crypts ...... 170 7.3 Discussion ...... 174

VIII. CONCLUSION ...... 176

REFERENCES ...... 181

APPENDIX

A. DOLPHIN SAMPLE COLLECTION METHODS ...... 205

A.1 Voluntary Collection Techniques ...... 206

B. ULTRASOUND CASE STUDIES ...... 208

B.1 Dolphin ovulation ...... 209 B.2 Two consecutive ovulations and conception ...... 209

C. FETAL SEXING, CERVICAL DILATION, PARTURITION AND POST-PARTUM ...... 222

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LIST OF TABLES

2.1 Dolphins in the Study ...... 34

2.2 Dolphin LH and POF Data ...... 41

3.1 Animals in Study ...... 53

3.2 Study phases, reproductive state, lactation and cohabitation ...... 60

3.3 Prolactin rises detected during the follicular phase ...... 64

3.4 Mammary gland size and state during pregnancy, lactation and post- weaning estrous ...... 74

4.1 Animals in study ...... 87

4.2 Ethogram of reproductive behavior ...... 97

4.3 Animal 1 number of reproductive behavioral occurrences and uE2 levels ...... 103

5.1 Animals in study ...... 117

5.2 Cervical mucus types ...... 121

5.3 Number of dolphin vaginal fluid arboriform types identified during anestrus, follicular and luteal phases ...... 124

6.1 Animals in study ...... 138

6.2 RP-HPLC results ...... 147

6.3 Novel peak minute range and reproductive status ...... 157

7.1 Animals in study ...... 169

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LIST OF FIGURES

2.1 LH test band color intensity scores...... 38

2.2 Dolphin ovarian follicular dynamics before during and after ovulation. Arrows indicate follicle ...... 44

2.3 Dolphin urinary progesterone levels showing increases in basal levels following ovulation ...... 45

2.4 Preovulatory follicle (POF) diameter before, during and after the luteinizing hormone (LH) surge...... 46

2.5 Dolphin corpus luteum (CL) appearance during pregnancy from artificial insemination ...... 47

3.1 Starting position of ultrasound transducer for mammary gland parenchyma visualization ...... 57

3.2 Prl means and maximum levels during studied phases ...... 61

3.3 Prl means during reproductive states and lactation...... 61

3.4 Prl, uE2 and uP during the estrous cycle of animal 2...... 63

3.5 Follicle growth and studentized residual prl rises during the follicular phase ...... 64

3.6 Identification of Prl outliers using polynomial regression during the follicular phase ...... 65

3.7 Ultrasound of follicular dynamics in animal 3 during prl rises ...... 66

3.8 uE2 and prl during anestrus, cohabitation, follicular phase and ovulation ...... 67

3.9 E2 and prl in the final days leading to parturition ...... 68

3.10 Prolactin levels during the first trimester of pregnancy in a normal and abnormal pregnancy ...... 69

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3.11 Minutes of nursing and number of latches in a 24 hour period during the first 15 days of life for two calves ...... 70

3.12 Prl levels during cohabitation with a mother/calf pair ...... 71

3.13 Non-mother cohabitation with mother-calf dyads ...... 72

3.14 Ultrasound of the mammary parenchyma ...... 75

4.1 Animal 1 ultrasound ...... 92

4.2 Animal 6 ultrasound ...... 93

4.3 Animal 7 ultrasound ...... 93

4.4 Animal 1 urinary estradiol, progesterone and luteinizing hormone ...... 95

4.5 Percentage of reproductive behaviors during each reproductive phase for animal 1 ...... 100

4.6 Immobility (IM) behavior...... 102

4.7 Animal 1 immobility bouts, total duration in seconds and maximum bout duration in seconds ...... 102

4.8 Animal 1 immobility, uE2, uLH and regression ...... 104

4.9 Animals 6 and 7 immobility duration and uLH correlation ...... 104

4.10 A dolphin mother gooses her 3 month old female calf...... 109

4.11 Genital tracking (GT) of animal 1 by animal 4 ...... 111

4.12 Goose (GOS) of animal 1 by animal 4 ...... 112

4.13 Dorsal Mount (DM) of animal 1 by animal 2 ...... 112

4.14 Copulation (COP) between animal 8 and animal 6 ...... 113

4.15 Line Up (LU) animal 1 at the front with animals 4 and 5 behind ...... 113

4.16 Mount (MNT) by animal 8 to animal 6...... 114

4.17 Avoid (A) by animal 6 of animal 8...... 114

5.1 Dolphin vaginal fluid type G ...... 122

5.2 Dolphin vaginal fluid type L ...... 122

5.3 Dolphin vaginal fluid type S ...... 123

5.4 Dolphin vaginal fluid type P ...... 123

5.5 Dolphin vaginal fluid type P4 ...... 124

5.6 Vaginal fluid arboriform types and spermatozoa during anestrus, follicular and luteal phases ...... 125

5.7 Spermatozoa on Vaginal Fluid Slide With Type P Mucus...... 125

5.8 uE2 and uP levels of animal 1 ...... 126

5.9 Animal 1 ultrasound ...... 127

5.10 Animal 2 ultrasound ...... 127

5.11 Vaginal walls ...... 128

5.12 Pseudo-cervix/vaginal flap ...... 129

5.13 Spermathecal Recess ...... 129

5.14 True cervix ...... 130

6.1 Animal 1 POF ...... 143

6.2 Animal 1 Ovulation ...... 144

6.3 Animal 1 uterine fluid and embryo ...... 144

6.4 Animal 1 uE2, uP and uLH ...... 145

6.5 Animal 1 baseline RP-HPLC peaks ...... 146

6.6 Novel peaks per minute range per day of study ...... 155

6.7 Number of novel peaks per day ...... 156

6.8 RP-HPLC of samples taken during lacational anestrus and static social grouping of animal 1 ...... 159

6.9 Milk protein fragment in dolphin urine ...... 160

7.1 Natal vibrassal hairs ...... 170

7.2 Mature dolphin vibrissal crypts ...... 171

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7.3 Vibrissal crypts ...... 172

7.4 In-situ vibrissae inside the crypts ...... 172

7.5 Capillary bed at the crypt terminus ...... 173

7.6 Formalin preserved vibrissae ...... 173

7.7 Sectioned formalin preserved vibrissae ...... 174

A.1 Sample collection photos ...... 206

A.2 Ultrasound exams ...... 207

B.1 Dolphin ovulation sequence ...... 209

B.2 Preliminary ovarian ultrasound upon arrival showed no follicular activity on either ovary...... 210

B.3 Follicular activity ...... 210

B.4 Ovulation and CL development during the non-mated cycle ...... 211

B.5 CL regression and new follicular development ...... 212

B.6 Pre-ovulatory follicle during the LH surge ...... 213

B.7 Ovulation and CL development during the conceptive cycle ...... 214

B.8 Early pregnancy ...... 215

B.9 Fetal development 4–24 weeks ...... 216

B.10 Dolphin ovarian cysts ...... 217

B.11 Serum progesterone levels in a dolphin with a reoccurring chronic luteal cyst...... 218

B.12 Follicular and luteal cyst ...... 220

B.13 Luteal cyst during treatment period ...... 221

C.1 Fetal sexing ...... 223

C.2 Dilated cervix ...... 223

C.3 Emerging flukes...... 224

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C.4 Expulsion of calf ...... 225

C.5 Delivery ...... 226

C.6 Calf’s first breath ...... 227

C.7 Mammary gland appearance before and after nursing ...... 228

C.8 Behavior ...... 228

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CHAPTER I

INTRODUCTION AND LITERATURE REVIEW

1.1 Introduction

The Atlantic bottlenose dolphin (Tursiops truncatus) is a medium sized species of

the order Cetacea which includes whales, dolphins and porpoises. Evidence suggests that

modern Cetaceans are close relatives of Hippopotamid artiodactyla (Geisler & Uhen,

2003; Gatesy, Hayashi, Cronin & Arctander, 1996). Slightly sexually dimorphic towards

males, Tursiops range in size from 135-635 kg and lengths of 2-4 meters, with a light gray to black coloration; they are found in temperate and tropical waters, costal and offshore populations around the world. They feed upon a wide variety of invertebrates and fish, and dolphins often utilize echolocation to catch prey (Mann, 1999; Ridgway &

Harrison, 1999).

Reproductive knowledge of this aquatic species is improving with recent advances in artificial insemination and sperm sexing technology (O'Brien & Robeck,

2006; Robeck et al., 2005; Muraco et al., 2010). However, research has primarily focused

on treatment induced estrous cycles specifically looking at how to maximize artificial

insemination success.

This long-term study utilizing conditioned dolphins in aquaria provides data

previously missing from the literature to better understand female dolphin reproductive

biology and provide a hypothesis of how dolphins may communicate reproductive readiness to one another.

1.2 Endocrinology

1.2.1 Estrogen synthesis and its role during follicular development and ovulation

Estradiol-17β (E2) is the most potent form of mammalian estrogenic steroids. It is produced by the theca interna and granulosa cells of the ovarian follicle under the synergistic, positive control of follicle stimulating hormone (FSH) and lutenizing hormone (LH) and under the negative influence of inhibin. As the follicle matures, LH receptors begin to develop in the granulosa cells and reach their highest concentration in the mature follicle, prior to ovulation. The appearance of receptors of LH in both thecal and granulosa cells marks a major event which enables the follicle to respond to the

rising levels of LH during the preovulatory surge. The interaction between the thecal and

granulosa cells is required for follicular steroidogenesis, and LH stimulates the

biosynthesis of androgens from cholesterol by thecal cells. The androgens then diffuse

across the basement membrane of the follicular wall, and a portion of these androgens reach the antrum. The granulosa cells uptake the androgens diffusing across the basement membrane and aromatize them to estogens under the influence of FSH. A portion of these

estrogens accumulate in the follicular fluid filling the antrum, but the majority diffuse

into the rich capillary bed of the thecal layer to be distributed into the systemic

circulation. Countercurrent transfer of androgens and other steroids from the venous to

the arterial blood occurs at the ovarian pedicle in some species providing additional

androgenic substrate for follicular steroidogenesis and estrogen biosynthesis (Austin &

Short, 1972; Holstein, 1994; Pineda & Dooley, 2003; Griffith, 2004). 2

The main physiologic role of E2 and estrogens are the development and

maintenance of the functional structure of the female sex organs by stimulating protein

synthesis and mitosis of estrogen-dependent organs. Secondary sex characteristics are

under estrogenic control and are responsible for the induction of sexual receptivity to

mating of the female. Estrogens secreted during the cycle by the developing follicles

induce pronounced vascularization increased blood flow and hyperemia water and salt

retention and edema of the uterus vaginal and vulva resulting in increased uterine tone.

Estrogens increase antibody levels in the uterus, enhance local protection against

infection and is responsible for the cornification of the vaginal epithelium in some

species (Austin & Short, 1972; Holstein, 1994; Pineda & Dooley, 2003; Griffith, 2004;

Greenspan, Gardner & Shoback, 1997).

1.2.2 Estrogen in the bottlenose dolphin

Sawyer-Steffan (1983) noted fluctuating immunoreactive serum estrogen levels in

non-pregnant and pregnant dolphins but did not identify cyclic patterns. Robeck et al.

(2005) found that peak urinary estrogen conjugates (EC; 5.4 +/- 3.8 ng/mg creatinine (Cr)

in bottlenose dolphins occurred 8 h prior to the LH surge.

1.2.3 Progesterone and corpus luteum formation

Progesterone is the primary hormone produced by the corpus luteum of the non-

pregnant, cycling animal and by the corpus luteum and placenta (of some species) during

pregnancy. It is needed for pregnancy maintenance supplied by the corpus luteum, the

placenta or both. Progesterone has been isolated from the adrenal cortices of several

species, however the main source are the luteal cells of the corpus luteum. The corpus

luteum of domestic species is formed by cells with distinct morphologic characteristics.

The CL of the cow, ewe, and sow contain two major cell types, large and small lutein cells which differ in their steroidogenic capabilities even though both types contain the same enzymatic machinery 3β hydroxysteroid dehydrogenase and capacity to synthetize and secrete progesterone. One model suggests that under the influence of LH, the small lutein cells, which probably originate from the thecal cells, have the capability to uptake and store cholesterol. Upon further stimulation by LH, the small lutein cells respond by secreting progesterone in short pulses. The large lutein cells probably originate from the granulosa cells and are more attenuated than the small lutein cells in their response to LH stimulation and in the uptake cholesterol from the small lutein cells. These cells would be able to secrete more progesterone for prolonged periods (Austin & Short, 1972; Holstein,

1994; Pineda & Dooley, 2003; Griffith, 2004; Greenspan, Gardner & Shoback, 1997).

The effects of progesterone are seen after the target tissue has been subjected to a period of estrogen stimulation. The estrogen priming leads to a synergistic response.

Progesterone acts on the uterus to cause quietening of the myometrium (Austin & Short,

1972; Holstein, 1994; Pineda & Dooley, 2003; Griffith, 2004; Greenspan, Gardner &

Shoback, 1997).

Large doses of progesterone inhibit gonadotropin output of the pituitary gland. In some species such as the cow, ewe, mare and sow this may regulate the length of diestrus because when the CL regresses and fails to secrete progesterone, a burst of FSH follows which in synergy with LH causes the development and maturation of the ovarian follicles

(Austin & Short, 1972; Holstein, 1994; Pineda & Dooley, 2003; Griffith, 2004;

Greenspan, Gardner & Shoback, 1997).

Declining blood levels of progesterone to less than 1.0 ng/ml in synergistic action with rising levels of estrogens brings about behavioral estrus in most domestic species. In seasonal breeders, such as the goat and ewe, progesterone is required at the beginning of the season before a full response to estrogen can be seen. This may explain the silent heat of the ewe during the first cycle of the breeding season. The first surge of estrogen is unable, after a prolonged period of seasonal anestrus, to elicit heat in the absence of

exposure to progesterone from a previous CL. The influence of progesterone on lactation

is important as it acts on the mammary glandular tissue as it acts on the uterine glands

(Austin & Short, 1972; Holstein, 1994; Pineda & Dooley, 2003; Griffith, 2004;

Greenspan, Gardner & Shoback, 1997).

Progesterone is essential during pregnancy. The early rise in progesterone from

the CL prepares the uterus for pregnancy. Progesterone acts on the endometrium to

inhibit myometrial activity and cause preparation for nidation regardless of whether an

embryo is present. Progesterone is known as nature’s immunosuppressant of the mother’s

rejection mechanism of the fetus (Austin & Short, 1972; Holstein, 1994; Pineda &

Dooley, 2003; Griffith, 2004; Greenspan, Gardner & Shoback, 1997).

1.2.4 Progesterone in the bottlenose dolphin

Progesterone has been identified in serum, urine, feces, blubber and milk to

identify pregnancy in bottlenose dolphins (West et al., 2000; Robeck et al., 2005; Robeck

& O'Brien, 2004; Robeck, Curry, McBain & Kraemer, 1994; Robeck et al., 2009;

Sawyer-Steffan, Kirby & Gilmartin, 1983; Sawyer-Steffan & Kirby, 1980; Kirby &

Ridgway, 1984; Schroeder & Keller, 1990; Schroeder, 1990; Cornell et al., 1987;

Schneyer et al., 1985; Wells, 1987; Biancani et. al, 2009; Pérez et al., 2011; Muraco et 5

al., 2009; Muraco et al., 2010).O’Brien and Robeck (2012) reported that progesterone concentrations peaked by week 7 post ovulation and remained elevated for the remainder of pregnancy (Weeks 8 up to 54). Additionally there is evidence that during the dolphin pregnancy progesterone is solely produced by the corpus luteum (Robeck et al., 2012).

Muraco et al., (2010) found that urinary progesterone levels reached above basal levels

(>1.0 ng/ml) between days 11-17 days post ovulation indicating an active corpus luteum.

1.2.5 Follicle-stimulating hormone and luteinizing hormone

Follicle-stimulating hormone (FSH) and luteinizing hormone (LH) are gonadotropins that are secreted from the anterior pituitary. FSH promotes follicular growth in the ovary, and LH acts on an ovary previously stimulated by FSH and is essential for estrogen synthesis, ovulation and initial development of the corpus luteum in most species. LH and FSH share biological similarities and close functional relationships.

Secretion of FSH and LH is regulated by gonadal steroids such as estrogens and progesterones and one known peptide, inhibin. The steroids and inhibin interact with hypothalamic GnRH secretion. Inhibin causes a negative feedback on the hypothalamus and pituitary gland to reduce secretion of FSH. The gonadotropes are bihormonal as they synthesize both LH and FSH. The frequency and amplitude of GnRH secretory pulses determine the relative proportions of LH and FSH secreted by the gonadotropes (Austin

& Short, 1972; Holstein, 1994; Pineda & Dooley, 2003; Griffith, 2004; Greenspan,

Gardner & Shoback, 1997). FSH and LH are synthesized continuously and stored in the

pituitary gland and are released throughout the estrous cycle. Proportions and levels

change depending on the cycle phase. There are three major forms of these hormones: the

basal level, where low and constant levels are found in the blood; pulses, sharp and 6

increased concentrations in the blood lasting for short periods; and a surge, a large, statistically significant increase in the concentration of a hormone in the blood above basal level that lasts for more than an hour. The ovarian follicles in mammals are dependent upon FSH and LH for follicular growth and maturation. Rising blood levels of estrogen suppress the pituitary release of FSH and facilitate release of LH (Austin &

Short, 1972; Holstein, 1994; Pineda & Dooley, 2003; Griffith, 2004; Greenspan, Gardner

& Shoback, 1997).

1.2.6 FSH and LH in the bottlenose dolphin

Schneyer et. al. (1985) was able to use a radioimmunoassay in dolphin serum to

identify FSH and LH levels. He found mean levels of FSH and LH from all dolphins to

be 0.22 ± 0.08 and 0.37 ± 0.18 ng/ml, respectively. He did not identify what phase of the

estrous cycle the dolphins were in other than pregnant or not. He found a slight increase

in both FSH and LH during pregnancy. Robeck et. al (2005) measured dolphin urinary

LH by a single antibody, direct enzyme immunoassay. He found the preovulatory LH

surge to last 20.3 +/- 5.1 h with time to peak LH being 9.4 +/- 3.1 h, and peak LH

concentrations to be 70.9 +/- 115.7 ng/mg Cr. Muraco et. al (2009 and 2010) utilized a

rapid immunochromatographic assay with dolphin urine to identify the LH surge and

peak effective for artificial insemination timing.

1.2.7 Prostaglandin

PGF2α is a luteolysin for domestic species that is responsible for induction of

labor, abortion and luteolysin or destruction of the corpus luteum. Several mechanisms

have been described to explain the luteolytic activity of PGF2a such as a prostaglandin-

induced constriction of the utero-ovarian vessels causing ischemia and starvation of the luteal cells; interference with progesterone synthesis; competition with LH for the receptor site; or destruction of LH receptor sites (Austin & Short, 1972; Holstein, 1994;

Pineda & Dooley, 2003; Griffith, 2004; Greenspan, Gardner & Shoback, 1997).

1.2.8 Prostaglandin in the bottlenose dolphin

Prostaglandin has been used to as a treatment to attempt to induce abortion in the

bottlenose dolphin (Robeck et al., 2012).

1.3 Dolphin estrous cycle

Female dolphins reach maturity at 5-12 years of age with a length of 220-235 cm

but most primaparous females are 8-10 years. There is no indication of reproductive

senescence, with a female lifespan of up to 50 years, females over 45 years old have

given birth (Wells et al., 1991; Ridgway and Harrison, 1999).

Bottlenose dolphins are sporadic, spontaneous ovulators and display a diffuse and

flexible reproductive seasonality (Urian et al, 1996; Muraco et al, 2010). They may give

birth in any season, but there is a slight increase in births in spring and fall (Urian et al.,

1996; Wells, Scott, Irvine, 1987; Thayer et al., 2003). Dolphins may also experience long

term anestrus lasting months or years (Yoshioka et al., 1986).

The estrous cycles of dolphins have been monitored by serum hormone levels,

urinary steroid metabolites, and urinary gonadotropins (Robeck et al., 2005; Robeck &

O'Brien, 2004; Robeck, Curry, McBain & Kraemer, 1994; Robeck et al., 2009; Robeck et

al., 1993; Sawyer-Steffan, Kirby & Gilmartin, 1983; Sawyer-Steffan & Kirby, 1980;

Kirby & Ridgway, 1984; Schroeder & Keller, 1990; Schroeder, 1990; Cornell et al.,

1987; Schneyer et al., 1985; Wells, 1984; Muraco et. al. 2009; Muraco et. al 2010).

The average length of the dolphin estrous cycle is 30 days with an average 10 and

20 day follicular and luteal phases respectively (Muraco et. al. 2009; Muraco et. al 2010;

Robeck et al., 2005). In addition, follicle growth and ovulation have been documented with real time ultrasonography indicating an average follicular size at ovulation to be 2.0 cm (Muraco et al., 2010; Robeck et al., 1998; Brook, 2001; Robeck et al., 2005).

1.4 Dolphin Reproductive Anatomy and Behavior

The female dolphin has a mid-ventral genital slit which houses the urethral

orifice, the clitoris and vagina. On either side of the genial slit, lie the oval mammary

glands with a small slit opening for each of the two nipples. The thick vaginal walls

contain numerous folds just caudal to the true cervix, forming a spermathecal recess, and

the true cervix delineates the uterine body from the vagina. The short uterine body joins

two horns forming a bicornuate uterus, where small fallopian tubes join the anterior ends

of the horns. The ovaries are located posterior to the kidneys and are held in place by the

broad ligament. Each ovary is covered by a large fallopian funnel at the outer end of the

fallopian tube (Ridgway, 1972).

1.4.1 Reproductive behavior

It is unknown how dolphins communicate reproductive readiness to one another

(Leatherwood & Reeves, 1989; Connor, Smolker & Richards, 1992), and until recently

estrus specific behaviors were unknown due to the socio-sexual nature of dolphins. In a

paper published by the author of this dissertation several estrus specific behaviors were

identified including genital tracking and immobility, a novel form of standing heat estrus

(Muraco & Kucjaz, 2015).

1.4.2 Pregnancy and Lactation

Gestation length in the dolphin is approximately 1 year (Ridgway & Harrison,

1999; O'Brien & Robeck, 2012). Real-time B-mode ultrasound can be used to identify

pregnancy in as early as 4 weeks gestation (Lacave et al., 2004), obtain skull and thoracic

fetal measurements (Williamson, Gales & Lister, 1990; Lacave et al., 2004) and predict

births (Lacave et al., 2004).

Active labor is typically less than 24 hours. The highly precocious calves are born

flukes first under normal circumstances with the umbilical cord intact until the final expulsion of the calf from the uterus into the water. The flukes may be seen emerging from the female for several minutes to hours before the actual birth. Calves born headfirst

are not uncommon, but there is a slightly greater risk of premature umbilical rupture

leading to drowning. The dolphin placenta is epitheliochorial similar to horses (Watanabe

et al., 2007) and is passed 8-18 hrs after birth. At birth, calves range in length from 84-

140 cm (Ridgway & Harrison, 1999) and weights of 11-18 kg.

Lactation is the primary source of nutrition for at least 1 year, but lactation may continue for several years (Ridgway & Harrison, 1999). Solid food has been found in the stomachs of 4-11 m.o. calves (Ridgway & Harrison, 1999). Weaning typically occurs at

18-20 months, but material care may continue for 3-6 years (Wells, Scott MD & Irvine

AF, 1987). At separation, calves are typically 150 kg. Calving interval is 3 years, and

dolphins can be pregnant and lactating at the same time (Ridgway & Harrison, 1999).

1.4.3 Artificial Insemination

1.4.3.1 Introduction

Artificial insemination (AI) has had an enormous impact on improving reproduction and genetics of domestic animals. The first successful insemination was performed by Spallanzani (1784) in a dog, then 100 years later Heape (1897) reported that AI had been used in rabbits, dogs, and horses (Review by Foote, 2002). In a review by Durrant (1990), the first pregnancies from AI in an exotic species was the fox (Vulpes vulpes) in 1972, which was followed by other exotic animals in the 1970’s such as reindeer (Rangifer tarandus), wolf (Canis lupus), and red deer (C. elaphus). Modern advancements have enabled several high profile exotic animals to benefit from AI including but limited to: the giant panda (Ailuropoda melanoleuca)(Masui et al., 1989), the clouded leopard (Neofelis nebulosa) and cheetah (Acinonyx jubatus) ( Howard et al.,

1997), the Asian elephant (Elephas maximus) (Brown et al., 2004) and African elephant

(Loxodonta Africana) (Hildebrandt et al., 1999), and the Siberian tiger (Panthera tigris altaica)(Donoghue et al., 1993). The biggest challenge when trying to understand exotic animal reproduction is the limited availability of samples for analysis and funding shortages (Durrant, 1990). In many instances, these challenges can be quickly resolved by utilizing information and techniques developed for closely related domestic animals

(Durrant, 1990). However, taxonomically unique species may require extensive research to elucidate appropriate protocols, so cooperation among zoological institutions and universities is essential to enhance research efforts for exotic and endangered wildlife

(Durrant, 1990).

Cetaceans posed an especially unique challenge for early reproductive research pioneers. They had to work with extremely limited sample sizes, an inability to obtain serial samples for endocrinology, fragile animals who did not tolerate anesthetization or stress, and no directly related domestic animal to compare to. It was only after years of dedicated work in the fields of dolphin veterinary care, husbandry and training (Ridgway,

2008), that researchers were able to start testing the various hypothesis regarding cetacean reproduction.

The early challenges of dolphin artificial insemination were the inability to identify or predict estrous cycles, collect semen and properly time inseminations. Success with dolphin AI came by pinpointing ovulation using ultrasonography of ovarian follicles

(Brook, 1999; Brook, 2001), identification of the luteinizing hormone surge (Muraco et al., 2003; Robeck et al., 2005; Muraco et al., 2009; Muraco et al., 2010), the use of altrenogest for estrous synchronization (Young & Huff, 1996; Dougherty, Bossart &

Renner, 1999; Asa, 1999; Robeck et al., 2005; van Elk, Muraco & Durrant, 2005) and voluntary collection of semen samples (Schroeder, et al., 1983; Keller, 1986; Schroeder

& Keller, 1990; Hudson, 2002; Yuen et al., 2009).

The first successful pregnancy of a marine mammal from artificial insemination

(AI) was a killer whale (Orcinus orca) in 2000 at SeaWorld in San Diego through a collaborative effort with SeaWorld, The National Zoological Park, and The Zoological

Society of San Diego (Robeck et al., 2004a). The first successful artificial insemination of bottlenose dolphins resulted from a project led by Dr. Fiona Brook at Ocean Park in

Hong Kong where two dolphins (Tursiops truncatus aduncas) gave birth to a healthy calf

each in 2001 following inseminations in 2000 (Fiona Brook, personal communication;

Young, 2001).

Today artificial insemination in the bottlenose dolphin has become an achievable

population management tool in both the large (Robeck et al., 2013) and small aquarium

facility (Brook & Kinoshita, 2004; van Elk, Muraco & Durrant, 2005; Muraco et al.,

2006; Muraco et al., 2007; Muraco et al., 2010). Artificial insemination pregnancies have

been achieved using liquid stored semen (Brook & Kinoshita, 2004; Robeck et al., 2005;

van Elk, Muraco & Durrant, 2005; Muraco et al., 2007), frozen-thawed spermatozoa

(Robeck et al., 2005), and fresh and frozen-thawed sex-selected spermatozoa (Robeck et

al., 2013).

1.4.3.2 Sperm Collection and Cryopreservation

Application of dolphin sperm collection was briefly reported in 1977 when Hill

and Gilmartin manually collected a large volume ejaculate; then in 1981, a detailed

analysis was conducted on a semen sample collected via electro-ejaculation (Hill &

Gilmartin, 1977; Seager et al., 1981). The electro-ejaculated sample had a concentration

of 189 x 107 with 85% motility, and the sample was cryopreserved using a canine

cryodiluent (20% egg yolk, 11% lactose and 4% glycerol) in pellets on dry ice (Seager et

al., 1981). Upon thawing, motility levels remained high at 80% (Seager et al., 1981).

Fleming, Yanagimachi, and Yanagimachi (1981) collected an 18 ml semen sample by

electro-ejaculation and cryopreserved the sample using the same methods described by

Seager et al., 1981 and Seager, Platz & Fletcher, 1975. Samples showed 85% and 80%

motility pre and post-freeze and penetrated hamster oocytes (Fleming, Yanagimachi, and

Yanagimachi, 1981). 13

Later in the 1980’s and early 1990’s, dolphins were trained for consistent hand- service semen collection (Schroeder, et al 1983; Keller, 1986; Schroeder and Keller,

1990) to better facilitate artificial insemination. Several ejaculates were characterized and frozen using the canine pellet method described by Seager, Platz & Fletcher, 1975. Post- thaw motility was 60% by freezing pellets in liquid nitrogen vapor (Keller, 1986;

Schroeder and Keller, 1989; Schroeder and Keller, 1990).

Durrant et al., 2000, using one ejaculate from two male dolphins, described the effects of freezing rates, incubation times and the addition of glycerol before or after cooling to 4oC using pellets and cryovials. The findings showed that the most effective protocol used a moderate freezing rate -12.8oC min with 4% glycerol added after cooling of diluted 1:1 semen in a N-tris(hydroxymethyl)methyl-2-aminoethane sulfonic acid

(TES) TRIS yolk buffer (TYB) to 4oC (Durrant et al., 2000).

In 2002, Hudson presented a new method for training semen collection in the

bottlenose dolphin. The method included using positive reinforcement to increase the

reflexive muscle response of the penis retractor muscle. Once the muscle response was

strong, the penis naturally extended from the genital slit and became erect. Once fully

extended, a brief grasp of the trainers hand to the penis tip resulted in a full ejaculate into

a collection device (Hudson, 2002).

In 2004, Robeck and Obrien reported data on three different cryopreservation

methods of dolphin sperm. Four ejaculates were collected from three males using

methods described by Schroeder & Keller (1989), and subjectively reviewed for total

motility, percentage of progressive motility and kinetic rating (Robeck & Obrien, 2004).

For viability assessments, a live-dead exclusion stain (eosin-ingrosin) was used, and for

acrosome and morphology, fixed and stained sperm were analyzed using bright-field optics (Robeck & Obrien, 2004). All samples were standardized to approximately 400 x

106 spermatozoa per ml (Robeck & Obrien, 2004). Final pre-freeze concentration for all

methods was 100 x 106 spermatozoa per ml, and the sperm suspension was transferred to

0.25-ml or 0.5 ml straws, sealed with ball bearings, and cryopreserved using a

programmable freezer (Robeck & Obrien, 2004). For method 1, semen was slowly

diluted over 5 min with Platz Diluent Variant (PDV) without glycerol (Robeck & Obrien,

2004). The PDV (350 mOsm kg21, pH 7.2) contained 11% lactose, 20% egg yolk, and 50

mg/ml of gentamicin The sperm suspension was cooled from 21oC to 5oC over 1 h, and

once at 5oC, the sperm suspension was placed into an ice-water bath (2oC) for 1 h, then

diluted 1:1 slowly with PDV containing 6% glycerol (3% final glycerol concentration)

(Robeck & Obrien, 2004). Dilutions with glycerolated cryodiluent for all methods (CM1,

CM2, and CM3) were made in a stepwise fashion (25%, 25%, and 50% of volume) at 10-

min intervals, and straws were frozen with a programmable freezer (Robeck & Obrien,

2004). For method 2, semen was diluted 1:3 (semen: diluent) with AE (Androhep

Enduraguard) and cooled to 15oC over 1 h, and after reaching 15oC, the sperm suspension

was centrifuged (15oC, 800, 3 g, 20 min) (Robeck & Obrien, 2004). The supernatant was

removed and the pellet was re-suspended 1:1 (semen:diluent) to a concentration of 150 x

106 spermatozoa ml with PDV (prepared as in method 1) at 15oC by diluting drop wise

over 5 min (Robeck & Obrien, 2004). After gentle mixing, the sperm suspension was

placed at 5oC and cooled over 1 h (Robeck & Obrien, 2004). Once at 5oC, the extended

semen was diluted 2:1 (semen:diluent) with cryodiluent (89.5% PDV, 9% glycerol, 1.5%

Equex STM for a final glycerol concentration of 3%), and straws were frozen using a

programmable freezer (Robeck & Obrien, 2004). For method 3, semen was diluted 1:1 with a 320 mOsm kg21, pH 7.2 TYB (Refrigeration Media; Irvine Scientific) without glycerol slowly over 5 min (Robeck & Obrien, 2004). The sperm suspension was cooled from 21oC to 5oC over 1 h (Robeck & Obrien, 2004). Once at 5oC, the sperm suspension

was diluted 1:1 with TYB containing 6% glycerol (Freezing Media; Irvine Scientific;

modified from 12% by dilution with Refrigeration Media) (Robeck & Obrien, 2004).

Straws were frozen in a programmable freezer (Robeck & Obrien, 2004). Semen was

thawed using three different methods: slow, medium and fast. Straws were shaken during

thawing, combined in a 5-ml polystyrene tube, and then diluted (1: 1, over 5 min) with

AE pre-warmed to 35oC. After dilution, sperm suspensions were held at 21oC (Robeck &

Obrien, 2004). The optimal method based on this study showed that a protocol

comprising a TYB/glycerol cryodiluent, slow cooling rate, rapid freezing rate, and

medium or fast thawing rate maintained high levels of initial and 6-h post-thaw sample

sperm characteristics (Robeck & Obrien, 2004). Results also showed that the

concentration of spermatozoa at freezing had a significant effect on post-thaw motility

with decreased motility in samples cryopreserved using a low sperm concentration (20 x

106 spermatozoa per ml) compared with the standard concentration (100 x 106

spermatozoa per ml) (Robeck & Obrien, 2004).

In 2005, Robeck et al. collected and cryopreserved nine ejaculates from 3 males

for the purposes of artificial insemination. Ejaculate concentration, volume, sperm

motility, and viability (plasma membrane integrity) were determined using standardized

techniques (Robeck & O’Brien 2004). Three different methods were used to cryopreserve

spermatozoa that were used during the AI attempts (Robeck et al., 2005). For method 1,

semen was pelleted using a sperm egg-yolk-citrate cryodiluent (EYC; 2.9% Na citrate,

20% egg yolk and 8% glycerol) and cooled from 21oC to 5oC over 1 h (Robeck et al.,

2005). At 5oC the sample was further diluted (1:1) over 5 min with EYC for a final glycerol concentration of 6%, and after incubation at 5oC for 2 h, the sperm suspension was frozen as approximately 0.2 ml pellets on dry ice for 5 min prior to plunging in liquid nitrogen (Robeck et al., 2005). For method 2, sperm samples were diluted with Biladyl

Fraction A (Minitube of America) (2:1, semen: diluent) slowly over 5 min (Robeck et al.,

2005). The sperm suspension was cooled from 21oC to 5oC over 1h (Robeck et al., 2005).

Once at 5oC, the sperm suspension was placed into an ice water bath (2oC) for 1 h, then diluted 1:1 slowly with Biladyl Fraction B (Fraction A with 14% glycerol (7% final glycerol concentration)) (Robeck et al., 2005). The sperm suspension was transferred to

0.5 ml straws, sealed and frozen in liquid nitrogen vapor at a distance of 4.5 cm above the vapor for 10 min, then plunged into liquid nitrogen (Robeck et al., 2005). For method 3, semen was diluted 1:1 with a TES-TRIS yolk buffer (TYB) (Refrigeration Media, Irvine

Scientific) without glycerol slowly over 5 min (Robeck et al., 2005). The sperm suspension was cooled from 21 to 5oC over 1 h (Robeck et al., 2005). Once at 5oC, the sperm suspension was diluted 1:1 with TYB containing 6% glycerol (Freezing Media,

Irvine Scientific, modified from 12% by dilution with Refrigeration Media; 3% final concentration) and loaded into 0.5 ml straws (Robeck et al., 2005). Straws were frozen using a programmable freezer (Minidigicool, IMV International) (Robeck et al., 2005).

These experiments showed that post-thaw sperm quality was highest using the programmable freezer in TYB extender (Robeck et al., 2005).

Muraco et al., 2005, presented information regarding experimental sex-sorting of dolphin spermatozoa at the 2005 Dolphin Neonate and Reproduction Symposium. Five male dolphins had ejaculates sex-sorted into 97% pure x spermatozoa with sort rates of

2,500–3,500 sperm per second at XY Inc. (Fort Collins, CO) using a high speed cell sorter (MoFlo SX; DakoCytomation) modified for sperm sorting (Johnson and Welch,

1999). Sperm was collected using voluntary methods described by Hudson, 2002.

Following the sort, 17 x 106 sorted sperm were cryopreserved into 0.25 ml straws using a

dolphin extender PDV (Platz Diluent Variant) with 3.5% glycerol, 11% lactose, and 20%

egg yolk. The sorted sperm had a post-thaw motility of 40%.

In 2006, O’Brien and Robeck reported on cryopreservation of dolphin

spermatozoa using the directional freezing technique (Arav et al., 2002) as well as sex-

sorting spermatozoa using a high speed cell sorter (MoFlo SX; DakoCytomation)

modified for sperm sorting (Johnson and Welch, 1999). A penis rinse was used prior to

unrestrained semen collection using a N-(Tris(hydroxymethyl)methyl)-

2aminoethanesulfonic acid (TES)-Tris buffered solution supplemented with 11mM

fructose and 50 ug mL-1 gentamycin (330 +/- 5 mOSm kg-1, pH 7.3 +/-0.1) (O’Brien &

Robeck, 2006). For transport to the sorting facility, within 10 minutes of collection, fresh

semen was diluted 3:1 (semen to dilutent) with TYB slowly over 5 minutes at 21oC

(O’Brien & Robeck, 2006). Semen was transported from San Diego to sorting facilities in

Texas within 15 hours of collection using an equitainer at 7-8oC (O’Brien & Robeck,

2006). After transport semen was held at 7oC until processing for flow cytometric sorting

(O’Brien & Robeck, 2006). In vitro sperm evaluations were made following semen

collection, transport to sort facility, staining with H33342, sorting (3500 sperm sex per

sec) and centrifugation (O’Brien & Robeck, 2006). Both sorted and non-sorted sperm were processed with addition of seminal plasma, cooled to 5oC and resuspended in the cryodiluent with final sperm concentration of 20 x 106 sperm per ml (O’Brien & Robeck,

2006). Sperm samples were transferred to 2 ml hollow tubes (IMT International) for

directional solidification. Machine (MTG-516 Isreal) (O’Brien & Robeck, 2006). The

hollow tube was moved through the first block (5oC) for 45s at a constant velocity before

reaching a distance of 2 mm into the opening of the second block (-50oC) and held for

15s for initiation of seeding, a rapid induction of ice nucleation throughout the length of the glass tube (O’Brien & Robeck, 2006). The tube was then moved at the same velocity across the second block for 3 min before entering the collection chamber (-100 to -110oC) followed by transfer to liquid nitrogen 50s later (O’Brien & Robeck, 2006). The hollow tubes were thawed in air for 45 s then transferred to a 35oC water bath equipped with modifications to enable uniform sample thawing over 45 s (O’Brien & Robeck, 2006).

The study found that directional freezing worked well for both sorted and non-sorted dolphin sperm (O’Brien & Robeck, 2006). However, acrosomal integrity and quality was decreased in sorted vs non-sorted thawed samples (O’Brien & Robeck, 2006). It was concluded that dolphin sperm can be transported to facilities within 20 h for sorting and freezing, and that the minimal AI dose for sexed frozen-thawed sperm was 150 million

(O’Brien & Robeck, 2006).

In 2009, Yuen et al. provided a detailed analysis of semen collection and ejaculate characteristics for the bottlenose dolphin (Tursiops aduncas). The findings showed that the first ejaculate collected was typically higher in volume but lower in sperm concentration, motility, and viability. The second and subsequent ejaculates showed a

decrease in volume but an increase in sperm concentration, motility and viability which provided a better analysis when determining potential fertility (Yuen et al., 2009).

In 2013, Robeck et al. discussed the development of deep intra-uterine artificial insemination with cryopreserved sex-sorted spermatozoa in the dolphin. For sperm processing, sex-sorting and cryopreservation, ejaculates from each male were diluted to

200–400 × 106 spermatozoa/ml with HEPES-TALP medium, without glucose and

calcium supplemented with 0.3% BSA , 96.7 IU mL−1 penicillin-G and 38.9 IU mL−1

streptomycin sulfate (Robeck et al., 2013). The sperm solution was incubated with

H33342 as previously described (O’Brien and Robeck, 2006). A 40 psi high-speed flow

cytometer (SX MoFlo, Dako Colorado, Inc.) modified for sperm sorting (Rens et al.,

1998) was used to analyze and sort spermatozoa (Robeck et al., 2013). X chromosome-

bearing spermatozoa were sorted for two hours into the following solution: 3 ml of Part A

(TYB); 176 mM TES, 80 mM Trizma base (Tris), 9 mM fructose, 50 mg/ml gentamicin

sulfate, 20% egg yolk, ~330 mOsm/kg, pH ~7.3; with 1% seminal plasma, prepared as

previously described (O’Brien and Robeck, 2006). Following the sort, it was centrifuged

at 850 × g for 25 min at room temperature; the supernatant was removed and the pellet

was resuspended with the appropriate volume of Part A (containing 1% of seminal

plasma) to a final concentration of 40 × 106 spermatozoa/ml (Robeck et al., 2013). An

aliquot of sorted spermatozoa was re-analyzed by the flow cytometer to determine the

proportions of X- and Y-bearing spermatozoa (Welch and Johnson, 1999). Sperm

suspensions were cooled to 5oC over a 1.5–2 h period and diluted with Part B extender

(Part A with 3% glycerol) over 30 min (three steps of 25 vol%, 25 vol% and 50 vol%) to

a final concentration of 1.5% glycerol and ∼30 × 106 spermatozoa/ml (Robeck et al.,

2013). The samples were equilibrated for 1 h after the addition of Part B, and the sperm suspension was transferred to 2-ml hollow glass tubes (at 5oC, IMT International) for

cryopreservation using a directional solidification machine (MTG-516) as described

previously for dolphins (Montano et al., 2012; O’Brien and Robeck, 2006). The study

concluded that using a mix of TYB, TES, Tris, fructose, gentamicin sulfate, egg yolk,

seminal plasma at 330 mOsm/kg, pH ~7.3 with 1.5% glycerol and directional freezing

provided the best methodology for cryopreservation of 30 x 106 sorted dolphin sperm

(Robeck et al., 2013).

1.4.3.3 Estrous Cycle Manipulation, Induction and Synchronization

In 1975, Richkind and Ridgway conducted experiments to investigate plasma

levels of estrogens, corticosteroids and progestagens measured at different times of the

year in the dolphin. Gravid females were given 10 mg of NIH-FSH-OVINE-S9 I.M. in

mid-gestation and late-gestation, and in non-gravid females in December and February.

Females in early gestation did not receive FSH, and plasma levels of previously

mentioned hormones were obtained. The conclusion was that plasma levels of estrogens,

corticosteroids and progestagens in the dolphin are influenced more strongly by

pregnancy than by administration of FSH (Richkind & Ridgway, 1975).

In 1979, Sawyer-Steffan and Kirby attempted to define bottlenose dolphin

reproduction and induce ovulation. Under the assumption that dolphins were induced

ovulators, the authors used mechanical stimulation of the dolphin cervix and vagina using

an endotracheal tube in an attempt to induce ovulation. In addition to manipulation, an

injection of PMSG (Pregnant Mare Serum Gonadotropin) was given to stimulate

follicular growth. Some animals got just manipulation, some got both PMSG and 21

manipulation, and some just got PMSG. Hormonal induced ovulation experiments also were attempted, and ovulation was assumed by elevated progesterone levels of greater than 3 ng/ml (Sawyer-Steffan & Kirby, 1979).

The following list summarizes the induction treatments given (Sawyer-Steffan &

Kirby, 1979):

Injection: 800 IU PMSG followed by LRF (Luteinizing releasing factor)

Injection: 1200 IU PMSG followed by LRF

Injection: 800 IU PMSG

Injection: 1200 IU PMSG

Injection: 675 IU Pergonal (Human menopausal gonadotropin)

Injection: 2800 IU PMSG followed by 1300 IU HCG (Human chorionic

gonadotropin)

Injection: 2800 IU PMSG followed by 1800 IU PMSG followed by 1000 IU HCG

The authors didn’t prove or disprove reflex ovulation due to no progesterone increase during the experiments, but there was evidence of spontaneous ovulation through elevated progesterone results of a non-experimental animal who was not with a male. PMSG doses of 1800 or less did not stimulate follicular growth, but ovulation was induced twice in one female using high doses of PMSG (>2800) (Sawyer-Steffan &

Kirby, 1979). PMSG greatly increased estrogen levels (7 pg/ml to 700 pg/ml) and caused strong sexual behavior in the females. Overstimulation of the ovary occurred on several occasions (Sawyer-Steffan & Kirby, 1979).

Sawyer-Steffan, Kirby and Gilmartin (1983) reported that bottlenose dolphins are capable of spontaneous ovulation without gonadotropin therapy. It was suggested that

dolphins can be induced to ovulate using high doses of IM injected PMSG followed by

HCG, and that spontaneous ovulation is likely to followed an induced one (Sawyer-

Steffan, Kirby & Gilmartin, 1983). Kirby and Ridway (1984) concluded that bottlenose dolphins exhibit spontaneous ovulation although they may experience anestrus lasting for one year. Schroeder and Keller (1990) reportedly induced ovulation in 14 out of 20 bottlenose dolphins using two injections of PMSG (1,600 IU followed by 800 or 1,000

IU) 2 d apart followed 5 d later by one injection of hCG (1,000 or 3,000 IU). However, artificial insemination performed 8 and 24 hours after induced ovulation did not result in any pregnancies (Schroeder and Keller 1990).

In 1996, Young and Huff reported on the safe and effective use of ReguMate

(altrenogest) as a fertility management treatment in a female killer whale (Orcinus orca).

In 1998, Robeck et al. evaluated an ovulation induction protocol for the bottlenose dolphin using eCG (equine chorionic gonadotropin) for follicular recruitment and hCG to induce ovulation. Ovulation was assumed by a post eCG treatment rise in serum estradiol and post hCG treatment rise in serum progesterone. No pregnancies resulted when the treated animals had free access to a breeding male nor did the 5 females who were inseminated, although 2 of the 5 females experienced a brief rise in progesterone (Robeck et al., 1998). Exogenous gonadotropins were further studied as ovulatory induction agents using PG600 (each 5-ml dose of PG600 contains 400 I.U. of pregnant mare serum gonadotropin (PMSG) and 200 I.U. of human chorionic gonadotropin (hCG)) and hCG.

Two female dolphins were determined to be anestrus through weekly serum P4 and E2 for one month, and both animals were stimulated with PG600 for follicular recruitment and 6 days later attempted ovulation induction with hCG (Robeck et al., 1998). The study

found that exogenous doses of PG600 and hCG in dolphins can result in abnormally high numbers of follicles and follicular cysts as observed on ultrasound (Robeck et al., 1998).

This suggests that dolphins are sensitive to exogenous gonadotropins causing the ovaries to become hyper-stimulated. There was a P4 rise 48 hours post hCG administration, however no evidence that ovulation occurred was visualized by ultrasound (Robeck et al.,

1998). Ultimately it was found that PG600 had a better response than eCG and that hCG may play a role in follicular recruitment and growth (Robeck et al., 1998).

In 1999 there was a Bottlenose Dolphin Reproduction Workshop in Silver

Springs, MD as part of the AZA Marine Mammals Taxon Advisory Group. This workshop resulted in two major advancements for dolphin reproduction: ultrasound and altrenogest treatment. Brook (1999) was the first to report on the successful methodology of using ultrasound to observe follicular development and ovulation in dolphins that had not been treated with exogenous gonadotropins and without hyper-stimulated ovaries.

This technique enabled clinicians a precise method for monitoring the reproductive cycle in the dolphin. Two clinical experiments were presented that showed that the progestin

ReguMate (altrenogest) was a safe and effective contraception and estrus synchronization tool in Pacific white-sided dolphins and bottlenose dolphins (Asa, 1999; Dougherty,

Bossart & Renner, 1999).

In 2000, Robeck et al., using Brook’s (1999) ultrasound techniques and Young and Huff (1996), Asa, (1999) and Dougherty, Bossart & Renner’s (1999) application of

ReguMate (altrenogest) in cetaceans, presented information on an ovulation induction trial with three dolphins. Ultrasound was used to classify the dolphins as anestrus or cycling with cycling animals showing follicles > 5 mm (Robeck et al., 2000). All three

animals were placed on altrenogest (Regu-Mate) for 16 days. After a single dose of 1500

IU of PG600 IM and 17.6 mg FSH IM on day 14, animal 2 and 3 responded with increased cortical activity or antral follicle activity (Robeck et al., 2000). After a second dose of 1800 IU PG600 IM on day 22, animal 2 exhibited no ovarian change and animal

3 had grown three follicles > 25 mm (Robeck et al., 2000). Animal 3 was administered two doses of 100 ug of GnRH 10 days apart, but GnRH administration did not stimulate ovulation or luteinization despite the presence of follicles similar in size to preovulatory follicles (Robeck et al., 2000).

In 2005, Robeck et al., evaluated the effects of orally administered altrenogest as an estrous synchronization tool in dolphins. Altrenogest had already been used to synchronize estrous in killer whales, Pacific white-sided dolphins and dolphins (Young &

Huff, 1996; Asa, 1999; Dougherty, Bossart & Renner, 1999; Robeck et al., 2004b). A total of 27 treatments were administered to 12 female dolphins, and animals were placed on 0.044mgkg21 p.o. altrenogest (Regu-Mate) for 27–77 days (Robeck et al., 2005). This study, based on urinary endocrine monitoring of two natural and nine post-altrenogest cycles, suggested that the dolphin estrous cycle was 36 days long and comprised of an 8 day and 19 day follicular and luteal phase, respectively (Robeck et al., 2005). Thirteen of the 27 estrous synchronization attempts resulted in an ovulatory cycle with ovulation occurring 21 days post-altrenogest treatment (Robeck et al., 2005). Artificial insemination was attempted in 8 of the estrous cycles resulting in 5 pregnancies (Robeck et al., 2005).

In 2005, van Elk, Muraco and Durrant reported on the use of altrenogest to synchronize dolphins for artificial insemination. In 2004, at the Dolfinarium Harderwijk

in Holland, 4 female dolphins were treated with altrenogest (12 mg per animal SID) for

35 days. Three out of the four treated animals ovulated on day 19, 20 and 26 post- altrenogest based on urinary LH testing (Muraco et al., 2003), ultrasound (Brook, 2001) and serum progesterone testing. Artificial insemination was performed on the three cycling females using the uLH surge for timing the procedures. One pregnancy resulted and was the first dolphin pregnancy by artificial insemination in Europe. Additionally, one of the non-pregnant females ovulated again 35 days following the altrenogest cycle, was inseminated using the uLH for timing and became pregnant (van Elk, Muraco &

Durrant, 2005).

In 2006, O’Brien and Robeck, and 2013, Robeck et al., continued to successfully synchronize dolphin estrus for artificial insemination using altrenogest treatment (0.044

mg kg p.o.) for 20 days.

1.4.3.4 Artificial Insemination

In 1990, Schroeder and Keller performed dolphin artificial insemination with

fresh and frozen-thawed semen collected using Keller’s (1986) hand-service techniques.

For the inseminations, a flexible fiber-optic laryngoscope (50 cm long and 6.5 mm in

diameter) with remote end-tip control was used. The inseminations were timed for 8 and

24 h after induced ovulation using two injections of PMSG and one injection of hCG

(Schroeder & Keller, 1990). Semen was placed into the spermathecal recess between the

external opening of the pseudo-cervix and the external opening of the true cervix

(Schroeder & Keller, 1990). Unfortunately, the elevated progesterone levels from two of

the females post-insemination spontaneously terminated after 10 and 14 weeks, so

pregnancy could not be confirmed (Schroeder & Keller, 1990). 26

In 1994, Robeck et al. described the internal reproductive anatomy of the dolphin including the spermathecal fold and cervical opening.

The first successful dolphin artificial insemination with subsequent births resulted from a project led by Dr. Fiona Brook at Ocean Park in Hong Kong where two dolphins

(Tursiops truncatus aduncas) gave birth to a healthy calf each in 2001 following inseminations using fresh liquid stored semen in 2000 (Fiona Brook, personal communication; www.newscientist.com). The inseminations were timed using ultrasound techniques described previously (Brook, 1999; Brook, 2001).

In 2003, Muraco et al, reported on the qualitative analysis of luteinizing hormone in the dolphin using a canine Rapid Immuno-Migration (RIM™) benchtop assay. This assay would quickly pinpoint the pre-ovulatory surge of LH facilitating artificial insemination timing. Then to further AI advancement efforts, Muraco et al, (2004), discussed the training of dolphins for artificial insemination using positive reinforcement.

These behaviors included semen collection, female blood collection, female urine collection, vaginal swabs, and ultrasound layout.

In 2005, Robeck et al, performed intrauterine and intracornual inseminations in eight dolphin estrous cycles resulting in five pregnancies. One pregnancy resulted from liquid stored semen and four using cryopreserved semen (Robeck et al 2005). To time the inseminations, females were given an altrenogest treatment for synchronization; urinary estradiol and luteinizing hormone were tested using a mobile EIA assay; and ultrasound was used to visualize follicular development and ovulation (Robeck et al, 2005). For the procedures, the females were sedated with diazepam (0.1–0.2 mg/kg), removed from the water, and placed in lateral recumbency. Inseminations were performed with flexible

endoscopes (9–11mm in diameter and 190–250cm long) advanced into the cranial vagina

(Robeck et al, 2005). The vagina was insufflated with air and a modified bullet tipped

catheter (400 cm, 2.2mm external diameter [6 French]) was used as a stylet to direct the

endoscope into the cervix (Robeck et al, 2005). Once the endoscope was in the uterus, the

inseminations were performed at the uterine body, partially into each horn or placed high

in the ipsilateral horn to the pre-ovulatory follicle (Robeck et al., 2005).

The first successful pregnancy in Europe from artificial insemination was

achieved in 2005 using liquid stored sperm (van Elk, Muraco, Durrant, 2005). Three

females were inseminated with timing based on altrenogest synchronization, a canine

rapid immuno-migration (RIM™) benchtop assay for urinary LH surge identification and

ultrasound for follicular development. For the procedures, the females were removed

from the water and placed in lateral recumbency. The inseminations were performed

using a 5.9 mm external diameter and 35 cm long flexible endoscope guided to the

vaginal fold by a 9.5 trachea tube. The vagina was not insufflated. A catheter was placed

through the working channel and into the cervix, and sperm was deposited into the

uterine body. One female became pregnant on the first cycle post-altrenogest and a

second female became pregnant from artificial insemination on the second (natural) cycle

post-altrenogest, 35 days from the first ovulation (Van Elk, Muraco, Durrant, 2005).

Additionally in 2005, Muraco et al., reported on sex-sorting dolphin sperm using

a using a high speed cell sorter (MoFlo SX; DakoCytomation) modified for sperm sorting

(Johnson and Welch, 1999). Four female dolphins were inseminated with methods

described previously (van Elk, Muraco, Durrant, 2005) using 250 million sex-sorted and

thawed sperm per female but no pregnancies resulted.

In 2006, O’Brien and Robeck inseminated 3 dolphins with sorted, frozen-thawed x bearing sperm resulting in 1 pregnancy. Estrus was synchronized using altrenogest

(0.044 mg kg p.o. for 20 days), and ovarian activity and uLH was monitored by ultrasound and an EIA. Sperm was sorted using a high speed cell sorter and frozen using a directional freeze technique (O’Brien & Robeck, 2006). Female dolphins were given diazepam for sedation, removed from the water and placed in lateral recumbency. The

20–30 minute long inseminations were performed 20 h after peak uLH concentration and

8 h before ovulation (O’Brien & Robeck, 2006). For the procedures, a flexible endoscope

(11.2 mm external diameter and 133 cm long) with a catheter in the working channel was

used. The females were inseminated with 360, 150 and 105 x 106 thawed sorted sperm,

placed in the uterine horn ipsilateral to the ovary containing the pre-ovualtory follicle

(O’Brien and Robeck, 2006). The minimal insemination dose recommended for sexed

frozen thawed sperm was 150 million (O’Brien and Robeck, 2006).

In 2006, Muraco, Arn and Muraco and in 2007, Muraco, Arn and Clough reported

on the artificial insemination, pregnancy and birth of a dolphin calf using fresh, chilled,

transported semen. The success was the result of small aquarium facilities working

together in order to obtain genetic diversity without moving animals.

In 2007, XY Inc. (Fort Collins, CO) sold the patent licenses for sorting non-

human sperm using a flow cytometer to Inguran LLC dba Sexing Technologies. In turn,

Sea World Parks and Entertainment purchased the exclusive rights to the sorting of all

exotic animal sperm using the proven flow cytometer methodologies.

In 2009, six dolphin calves had been born using sex-sorted fresh or frozen-thawed

sperm (O’Brien, Steinman and Robeck, 2009), and by 2012, Montano et al, reported that

13 dolphin calves had been born using sex-sorted cryopreserved spermatozoa with a sex predetermination success rate of 92%.

In 2013, Robeck et al., provided a methodology review of previously published data (Robeck et al., 1994; Robeck et al., 2005; O’Brien and Robeck, 2006; O’Brien et al.,

2009) as well as a description of deep-uterine inseminations in the dolphin. The findings showed that acceptable conception and calving rates using sexed frozen-thawed spermatozoa are achieved after mid-horn deposition of 200 × 106 spermatozoa with

females aged less than 25 y.

1.5 Discussion

Methodologies exist for the successful application of assisted reproductive

technologies to bottlenose dolphin population management, however numerous

challenges still remain. Despite the research on ovulation induction, semen

cryopreservation and sex-sorting spermatozoa, there is a lack of understanding of basic

bottlenose dolphin reproductive biology. The mechanisms of what initiates a natural

dolphin estrous cycle, why dolphins occasionally experience long-term anestrus,

nutritional effects on fertility, male infertility, genetic traits, and how dolphins

communicate reproductive readiness to one another are a few of the unknown aspects of

their biology. Many of the successful ART techniques that have been described in the

literature (Robeck et al., 2013) are not economically feasible or even obtainable to the

small zoo and aquarium facility. More research needs to be conducted on affordable ART

methodologies as well as general biology.

CHAPTER II

OVARIAN FOLLICULAR DYNAMICS DURING THE LUTEINIZING HORMONE

SURGE IN THE BOTTLENOSE DOLPHIN (TURSIOPS TRUNCATUS)

Muraco, H., Clough, P., Teets, V P. Clough, V. Teets, D. Arn, & M. Muraco. 2010. Ovarian Follicular Dynamics during the Luteinizing Hormone Surge in the Bottlenose Dolphin (Tursiops truncatus). International Journal of Comparative Psychology–Special Marine Mammal Research Issue. 23, 723–733

2.1 Introduction

The study of reproduction often requires identification and assessment of fleeting

but key physiological processes. The relationship between the ovulatory luteinizing

hormone (LH) surge and ovulation is one such example. Often such processes take place

within a few hours. Time from the ovulatory LH surge to ovulation in domestic species,

with the exception of the mare, occurs within 24-40 hrs (Pineda & Dooley, 2003). In the

Asian Elephant (Elephas maximus), the ovulatory LH surge lasts 24-48 hrs (Brown,

Schmitt, Bellen, Graham, Lehnhardt, 1999). The LH surge in the Giant Panda

(Ailuropoda melanoleuca )is less than 12 hrs (Durrant, Ravida, Spady, Cheng, 2006). In

the Pacific white-sided dolphin (Lagenorhynchus obliquidens) and killer whale (Orcinus orca), ovulation occurs within 31 and 38 hrs respectively following the ovulatory LH surge (Robeck et al., 2009; Robeck et al., 2004).

To accurately study the ovulatory LH surge, ovarian follicle dynamics, and ovulation in bottlenose dolphins (Tursiops truncatus), daily sampling is required. One of 31

the benefits of studying reproduction in zoo and aquarium species is the ability to obtain numerous samples from conditioned animals. In particular, with dolphins, non-invasive techniques such as urine collection and trans-abdominal ultrasound exams can be conducted multiple times daily. In this study, the dolphins were being monitored for artificial insemination and controlled natural breeding; therefore, samples were opportunistically collected based on clinical needs. Of the 10 dolphin ovulatory estrous cycles characterized, 9 were natural cycles and one was altrenogest induced. Altrenogest treatment is a method used to synchronize bottlenose dolphin estrous cycles (Robeck et al., 2005). The LH surge was identified by a commercially available rapid immunochromatographic assay (ICG) designed to detect serum LH in the domestic dog

(Canis familiaris). This is a previously discussed method for identification of bottlenose dolphin urinary LH (Muraco et al., 2009). Additionally, the same canine ICG LH assay has been used to identify LH in the Giant Panda (Durrant et al., 2006). The structure of

LH is well conserved among mammal species allowing for cross-reactivity of various LH antibodies (Liao et al., 2003). Real-time B-mode trans-abdominal ultrasound imaging was used to characterize ovarian follicular dynamics before, during and after the LH surge. Ultrasound has previously been proven to be an accurate method for identification of the pre-ovulatory follicle (POF) and ovulation in dolphins (Brook, 2001). Urinary progesterone assays were performed to ensure ovulation had occurred. Urinary steroid metabolites have been monitored in bottlenose dolphins by electrochemiluminescence immunoassay (ECLIA) and enzyme-linked immunosorbant assay (ELISA) (Muraco et al., 2009; Robeck et al., 2005).

The goal of the study was to better understand the relationship between ovarian follicular dynamics and the LH surge in the bottlenose dolphin. This knowledge will further understanding of dolphin reproduction, assist in the accurate timing for artificial insemination, and maximize controlled natural breeding programs.

2.2 Method

2.2.1 Experimental ethics

All animals, materials and methods used in the study were individually evaluated

by each facility’s Internal Animal Care and Use Committee (IACUC) to ensure safety,

health and well-being of the animals. Samples were taken during routine behaviors in

which the animals were conditioned to participate.

2.2.1.1 Animals

Bottlenose dolphins used in the study consisted of 8 females from 4 different

facilities (Table 1). Animals 1 and 2, located at Six Flags Discovery Kingdom (SFDK;

Vallejo, California USA), were housed in a 2717.93-L manufactured seawater pool

maintained at 21-22 oC year round. Animals 1 and 2 were fed a diet of 10.4 and 5.9 kg,

respectively, of whole frozen thawed herring (Clupea harengus) and capelin (Mallotus

villosus) daily. Animals 3 and 4, located at Dolphin Research Center (DRC; Grassy Key,

FL USA), were housed in 1093.98 and 552.67-L, respectively, of natural seawater with

an average temperature of 21.5 oC December-March. Animal 3 received a diet of 5.9 kg

of whole frozen thawed herring (C. harengus), capelin (M. villosus), and silversides

(Menidia menidia). Animal 4 was fed a diet of 5.4 kg of whole frozen thawed herring (C.

harengus), capelin (M. villosus), and marine smelt (Hypomesus pretiosus). Animal 5,

located at Theater of the Sea (TOTS; Marathon, FL USA), was housed in 548,884-L of natural seawater with an average temperature of 31.3 oC in July and August. Animal 5

was fed 6.22 kg of herring (C. harengus), capelin (M. villosus), squid (Loligo

opalescens), and lake smelt (Osmerus eperlanus mordax). Animals 6-8, located at The

Mirage Hotel (Mirage; Las Vegas, Nevada USA), were housed in a 3,141,892-L manufactured seawater pool, maintained at 21 to 23° C year-round. Animals 6-8 received a diet of frozen thawed herring (C. harengus), capelin (M. villosus), sardine (Sardina pilchardus), and squid (L. opalescens) for daily totals of 8.62, 7.26 and 8.16 kg, respectively.

Table 2.1 Dolphins in the Study

Reproductive Animal Facilitya Sex Birth Date Weight (kg) Historyb

1 SFDK F 7/1979c 184.09 Parous 2 SFDK F 7/1986c 189.09 Parous 3 DRC F 11/1984d 189.55 Parous 4 DRC F 2/2001d 168.64 Nulliparous 5 TOTS F 1/1983c 172.36 Parous 6 Mirage F 3/2000d 159.66 Nulliparous 7 Mirage F 5/1997d 176.44 Nulliparous 8 Mirage F 11/1975c 191.86 Parous

Note. aSFDK, Six Flags Discovery Kingdom; DRC, Dolphin Research Center; TOTS, Theater of the Sea; Mirage, The Mirage Dolphin Habitat. bReproductive history prior to urine collection used in the study. cEstimated birth date for wild caught animals. dAquarium born

2.2.1.2 Statistics

Arithmetic means and standard deviations (SD), presented as mean +/- SD, were

calculated using SYSTAT software (Systat Software, Inc. 225 W Washington St., Suite

425, Chicago, IL 60606).

2.2.1.3 Behavioral Conditioning

Operant conditioning techniques were used to train the dolphins for ultrasound

exams and urine sampling. Behaviors were positively reinforced and slow

approximations were taken until the dolphin was fully conditioned for daily sampling and

exams.

2.2.1.4 Ultrasonography

Real-Time B-Mode trans-abdominal ultrasound imaging was utilized. The dolphins were trained to float laterally and stationary at the water’s surface. The blowhole was submerged, but the animal could lift her head and take a breath as needed. To visualize the ovaries and POF, the ultrasound transducer was placed between the junction of the rectus abdominus muscle and the hypaxialis lumborum muscle (Brook, 2001). For animals 1-2 and 4, POF’s were visualized with a Sonosite Titan portable ultrasound machine with a 5-2 MHz curvilinear transducer (Sonosite Inc. 21919 30th Drive SE,

Bothell, WA 98021-3904, USA). Images were digitally captured using the internal digital

memory of the machine or by the use of an Archos 605 Audio/Visual Player (Archos Inc.

7951 E.Maplewood Avenue #260 Greenwood Village, CO 80111 USA). For animals 3,

7, and 8, an SSD 900 Aloka ultrasound machine with a 3.5 MHz transducer (Aloka

America, 10 Fairfield Boulevard Wallingford, CT 06492 USA) was used and images

recorded with a Sony digital video cassette recorder (Sony Corporation of America, 550

Madison Avenue, 27th Floor, New York, NY 10022-3211). For animals 5 and 6, a

Sonosite 180 Plus ultrasound machine with a 5-2 MHz transducer (Sonosite Inc., 21919

30th Drive SE, Bothell, WA 98021-3904, USA) was used and images recorded using

either the internal digital memory of the machine or by the use of an Archos 605

Audio/Visual Player (Archos Inc., 7951 E.Maplewood Avenue #260 Greenwood Village,

CO 80111 USA). For all animals, ultrasound exams were conducted 1-3 times daily

(Table 2) and exams typically lasted 3-5 minutes for both the left and right lateral

positions.

2.2.1.5 Urine Collection and Hormone Assays

The dolphins were trained to urinate on cue. Initially the trainer placed firm hand

pressure to the dolphin’s bladder, which often resulted in reflexive urination. This

reflexive behavior was paired with primary reinforcement to increase the frequency. Over

time, each dolphin was conditioned to urinate by the application of gentle pressure to the

bladder. For animals 1-5, urine collection was achieved by placing the dolphin in a

ventral position and the genital slit was wiped clean with a dry cloth. A sterile plastic 25-

ml syringe was used to draw up the urine once it free flowed from the urethra. For

animals 6-8, the dolphins were partially pulled out of the water so that the genital slit was

dry. First, the trainer stood on a dry ledge and grasped the tail flukes of the dolphin that

was stationary in a dorsal position floating at the pool surface. The dry ledge was 7 to 10

cm higher than the water surface. Next, the trainer gently stepped backward pulling the

dolphin’s flukes backward and laterally over the ledge until the dolphin’s genital slit was

laterally out of the water. The dolphin’s head and pectoral flippers remained in the water. 36

The genital slit was wiped clean with gauze. A 50-ml sterile plastic specimen cup was placed under the urethra and gentle pressure was applied to the bladder. The urine was caught in the cup as it freely flowed from the urethra. An average of 5-10 ml of urine was collected from all study animals. Urine was immediately frozen at -70 oC (animals 1-2), -

17 oC (animals 3-5), and -33.9oC (animals 6-8) in 3-5 ml increments in plastic storage vials.

Urinary progesterone (uP) was measured using an ECLIA tested with the Elecsys

2010 instrument (Roche Diagnostics, Mannheim, Germany) at Clinical Pathology

Laboratory in Las Vegas (4275 S. Burnham Ave, Suite 325, Las Vegas, NV 89119,

USA). Urine was shipped frozen to the laboratory from each facility.

Urinary luteinizing hormone (uLH) was measured using the ICG Canine Witness®

Luteinizing Hormone Assay (Witness Synbiotics Corp., Kansas City, MO, USA). The assay provides a rapid, semi-quantitative visible color band in the presence of dolphin LH

(Muraco et al., 2009). Dolphin urine was applied to the test strip either directly following collection, or, if frozen, after thawing. Urine was applied to LH assays either neat

(nUr)(Animals 1-6) or after concentrating (cUr) (Animals 7-8). For the nUr samples, a

100-µl volume was placed onto the test strip and results were obtained in less than 1 h after application. For the cUr samples, urine was centrifuged and the supernatant was concentrated on a Microcon-10 filter device (Millipore Corp., Bedford, MA, USA) at

13,900 g for 35 min before application of 100 µl cUr to the LH assay. Results were obtained in less than 1 h following application.

All LH test band results were scored from 0-4 based on color band intensity. Test

bands were scored as follows (Figure 1):

0=no visible test band

1=faint test band

2=test band less intense than control band

3=test band and control band equally intense

4=test band more intense than control band

Figure 2.1 LH test band color intensity scores.

C = Control. T = Test.

2.2.1.6 Estrous Cycle Monitoring

Animal 3 was given 110 mg/kg of altrenogest (Regu-Mate, Intervet Inc.,

Millsboro, DE, USA) once a day for 113 d for the purposes of estrous synchrony. She was monitored for ovulation following the altrenogest treatment, and the subsequent

natural estrous cycle, by ultrasonography of the POF, uLH, and uP. Animals 1-2 and 4-8,

experienced natural estrous cycles without any altrenogest pre-treatment. Monitoring for

ovulation included ultrasonography of the POF, uLH, and uP.

2.3 Results

Ultrasound was used to measure the POF vertical diameter before, during, and

after the LH surge as well as to identify the disappearance of the POF indicating that

ovulation had occurred (Table 2) (Figure 2a-d) (Figure 4). Mean POF diameters 18-29 h

prior to the LH surge were 1.767 +/0.189 cm (n=6). Mean POF diameters during the LH

surge were 1.942 +/- 0.098 cm (n=9). Mean POF diameters 14-24 h post the last recorded

LH surge were 1.835 +/- 0.289 cm (n=4). Animals 1 and 2 ovulated 24 and 24 h 30 min,

respectively, from the maximum recorded POF diameter during the LH surge. Mean time

to disappearance of the POF from the last recorded LH surge was 37.475 +/- 12.346 h (n

= 6). In animal 1, fluid evacuation from the follicle during ovulation was visualized and

photographed (Figure 2c,d). Complete evacuation of the follicular fluid from 1.5 cm to 0 cm was 4 h 20 min. A 2.0 cm corpus luteum (CL) was visible 6 d post-ovulation and a

3.0 cm CL was visible 22 d post ovulation (Figure 5a). An embryo was visualized 57 d

post ovulation (Figure 5b). CL’s were visualized at 100 and 142 d post ovulation to show

the relationship between fetal positioning and the ovary (Figure 5c,d).

Urinary LH was profiled before, during and after the ovulatory LH surge in 9

natural estrous cycles from 8 bottlenose dolphins using the canine LH assay (Table 2).

There was no significant score difference between results using nUr vs. cUr. A score of

0-4 was given to the darkness of the color band on the test strip (Figure 1). Mean time in

hours from the last tested baseline (score: 0–2) uLH sample to the first tested uLH surge

(score: 3-4) sample was 19.2 +/- 5.491 h (n=8). Mean uLH scores 7-24 h 30 min prior to

the LH surge were 0.5 +/- 0.5 (n=8). The LH surge (score: 3-4) duration from tested urine samples collected 2-4 x per day was 6.050 +/- 1.332 h (n = 6). Mean uLH surge scores

were 3.476 +/- 0.680 (n=21). Mean time from the last recorded peak uLH sample to the first baseline (score: 0-1) uLH sample was 19.393 +/- 3.797 h (n=7). Mean uLH scores

15- 23 h post the LH surge were 1.143 +/- 0.378 (n=7) (Figure 4).

Urinary progesterone (uP) was tested the day of the LH, surge and then again at 7-

10d, 11-17d and 18-24 d (Figure 3). Mean uP levels the day of the LH surge were 0.271

+/-0.125, at 7-10 d were 0.7 +/ 0.414 ng/ml, at 11-17 d were 1.188 +/- 0.482, and at 18-

24 d were 1.843 +/- 1.488.

Estrous cycles were ovulatory based on the disappearance of the POF on

ultrasound and subsequent rises in uP following ovulation (Figure 2d) (Figure 3). Animal

3 ovulated on day 21 following the end of altrenogest treatment and then ovulated again

naturally 32 days later based on days between peak size of the POF. No other animals in

this study received any altrenogest pre-treatment. Animal 5 cycled twice with 36 days

between peak LH levels. 6 of the ovulations occurred on the right ovary and 4 occurred

on the left. Of the two animals that experienced 2 successive ovulations, animal 3

ovulated both times on the left side and animal 5 ovulated both times on the right side.

The 9 natural cycles occurred in March (animal 4), May (animals 1-2), June (animal 6),

July (animal 5), August (animal 5), October (animals 7-8), and December (animal 3). The

one altrenogest-induced cycle occurred in November (animal 3).

2.4 Discussion

To effectively study the relationship between the LH surge, follicular dynamics

and ovulation of the bottlenose dolphin requires careful monitoring and multiple samples

taken daily. Identification of these fleeting processes would be impossible to achieve

without conditioned dolphins. One benefit to this project was the ability to monitor 40

natural cycles. This allowed for an accurate representation of the estrous cycle without influence of an artificial treatment regimen such as altrenogest. The study showed that the rise and fall of the dolphin ovulatory LH surge can be detected by the canine rapid ICG assay, and scoring the color band intensity can provide a semi-quantitative measurement tool. The POF during the LH surge was a turgid round structure with thickened walls and a mean diameter of 1.9 cm (Figure 2b). In all the study animals, the POF remained intact for the duration of the LH surge with ovulation occurring 24-48 hours later. Interestingly,

6 of the ovulations (4 conceptive) occurred on the right ovary and 4 ovulations (1 conceptive) on the left ovary. This finding is different from previously documented results of dolphin ovulations occurring 82% of the time on the left ovary (Robeck et al.,

2005). Of the 10 estrous cycles in this study, 4 were conceptive from artificial insemination (AI) and 1 from natural breeding. Of the remaining non-conceptive cycles,

2 were non-bred, 2 were AI and 1 was from a natural breeding. Further study will compare non-conceptive to conceptive cycles to look for potential infertility patterns in the endocrinology and physiology. Understanding the relationship between LH, POF’s, and ovulation will aid natural breeding programs and advance artificial insemination techniques.

Table 2.2 Dolphin LH and POF Data

Animal Date Timee LH Score Timef POF (cm) Conceptive

1 5/17/2009 1040 0 930 1.7 Yes–AI+ 1450 0 - - 5/18/2009 900 3 900 2 1130 2 1300 2.1

Table 2.2 (continued)

1400 2 - - 1500 4 - - 5/19/2009 900 2 840 1.5 1420 1 1305 0* 2 5/25/2009 1315 0 840 1.72 Yes–AI 5/26/2009 1100 4 1000 2 5/27/2009 1100 2 1030 0 3c 11/11/2005 - - - 1.8** Not Bred 11/12/2005 - - - 0** 3d 11/12/2005 930 1 1800 1.8 Yes–AI 12/12/2005 900 4 - - 1130 3 - - 1340 3 - - 1500 3 1800 1.9 12/13/2005 - - - - 4 2/28/2009 915 1 930 2.08 Yes–AI 1600 2 - - 3/1/2009 1300 4 1345 2.08 3/2/2009 945 1 1345 2.14 3/3/2009 - - 1625 0* 5a 7/6/2008 1400 1 - - Not Bred 7/7/2008 1130 4 - - 1400 3 1430 1.9 7/8/2009 1130 1 - - 7/9/2008 - - 1430 0* 5b 8/11/2008 - - 1600 1.8 Yes–NB++ 8/12/2008 1130 4 1030 1.9 8/13/2008 - - - - 6 6/11/2006 900 1 1030 1.5 No–NB

Table 2.2 (continued)

6/12/2006 800 4 1000 1.8 1200 4 - - 1400 4 1430 1.9 1600 3 - - 6/13/2006 900 1 930 2 6/14/2006 - - 930 0* 7 10/12/2004 800 0 - - No–AI 1500 0 - - 10/13/2004 800 3 - - 1500 4 1515 1.8 10/14/2004 800 1 - - 8 10/11/2004 800 0 - - No–AI 1500 0 - - 10/12/2004 800 0 1000 1.7 1500 4 1530 1.7 1930 4 1900 1.9 10/13/2004 800 1 900 1.7 10/14/2004 - - 900 0* Note. aFirst natural cycle. bSecond natural cycle. cAtrenogest cycle. dNatural cycle. eIndicates time of urine collection. fIndicates time of ultrasound exam.–Indicates no data collected. * Indicates POF had ovulated. **Data not included in calculated means. + Indicates artificial insemination. ++ Indicates natural breeding.

Figure 2.2 Dolphin ovarian follicular dynamics before during and after ovulation. Arrows indicate follicle

Figure 2a, 1.9 cm Preovulatory follicle (POF) before the luteinizing hormone (LH) surge. Figure 2b, 2.0 cm POF during the LH surge. Figure 2c, 1.5 cm follicle during ovulation. Figure 2d, fully evacuated follicle following ovulation.

Figure 2.3 Dolphin urinary progesterone levels showing increases in basal levels following ovulation

Urinary progesterone (uP) levels. 0 =Day of LH surge. 1 = 7-10 d. 2 = 11-17 d. 3 = 18–24 d.

Figure 2.4 Preovulatory follicle (POF) diameter before, during and after the luteinizing hormone (LH) surge.

Figure 2.5 Dolphin corpus luteum (CL) appearance during pregnancy from artificial insemination

White arrow with black outline indicates CL. Long white arrow indicates embryo/fetus. Figure 5a, CL at 22 days post-ovulation. Figure 5b, CL and embryo at 57 days post- ovulation. Figure 5c, CL and fetus at 100 days post-ovulation. Figure 5d, CL and fetus (lateral view of rostrum and skull visible) at 142 days post-ovulation.

CHAPTER III

URINARY PROLACTIN CONCENTRATIONS IN THE FEMALE BOTTLENOSE

DOLPHIN (TURSIOPS TRUNCATUS)

3.1 Introduction

Prolactin (prl) is a remarkable polypeptide hormone which controls a diverse amount of physiological processes in response to a wide array of physiological and environmental stimuli (Freeman, Kanyicska, Lerant & Nagy, 2000; Bernichtein, Touraine

& Goffin, 2010). For the purposes of this paper, prolactin’s role in lactation, reproduction, pregnancy and maternal behavior will be the main focus. No information about concentrations of dolphin prolactin was found in the literature. In one histological study, Cowan et al 2008, identified that prolactin could be found in the bottlenose dolphin pituitary gland (Cowan, Haubold & Tajima, 2008).

Prolactin is secreted primarily from lactotropes in in the anterior pituitary. The impulse is for continuous spontaneous secretion but this impulse is inhibited by dopamine from the hypothalamus (Leong, Frawley & Neill, 1983; Neill & Nagy, 1994). Lactation is prolactin’s most well-known role in mammals where it induces mammogenesis and synthesizes lactogenesis and maintains galactopoiesis (Freeman, Kanyicska, Lerant &

Nagy, 2000). As young suckle the mothers nipple, dopamine inhibition is reduced, and prolactin secretion is released in a series of bursts from the pituitary (Leong, Frawley &

Neill, 1983; Neill & Nagy, 1994; Bodnár, Bánky, Tóth, Nagy & Halász, 2002). Prolactin

is released 2-5 minutes after suckling starts, and if nursing continues for 20-30 minutes, prolactin levels may reach 60 times higher than baseline in humans (Leong, Frawley &

Neill, 1983).

In addition to lactation, prolactin plays a varied role in female reproduction. In the estrous cycle of the rat, prolactin secretion appears low until a preovulatory surge occurs similar in timing to the lutenizing hormone (LH) surge (Freeman, 1994; Butcher RL,

Collins WE & Fugo NW, 1974; Terry LC et al., 1977). There has been an observation of a slight mid-cycle increase in prolactin in primates (Aidara, Badawi, Tahiri-Zagret &

Robyn, 1981; Vekemans, Delvoye, L'Hermite & Robyn, 1977). During the breeding season of sheep, GnRH stimulates prolactin release (Henderson, Hodson, Gregory,

Townsend & Tortonese, 2008), and in mares, prolactin appears to play a role in follicular maturation and ovulation (Bennett-Wimbush, Loch, Plata-Madrid & Evans, 1998; King,

Roser & Jones, 2008). Prolactin is correlated with day length and regulation of reproduction in seasonally breeding animals (Fitzgerald, Davison & McManus, 2000).

Estrogen can stimulate prolactin production and secretion in various species (del Pozo &

Brownell, 1979). In the rat, estrogen is a critical factor in producing the proestrous prolactin surge (Nicoll, Meites & Blackwell, 1962; Lieberman, Maurer & Gorski, 1978;

Christensen, Zeng, Murawsky & Gregerson, 2011; Lawson, Haisenleder & Marshall,

1993). In the pig, prolactin surges at estrus are induced by estrogen (Stevenson, Cox &

Britt, 1981), and in the elephant, there is a prolactin rise during the follicular phase that may be estrogen dependent (Bechert, Swanson, Wasser, Hess & Stormshak, 1999). In the mare, a rise in prolactin coincides with a decline in estradiol that occurs before ovulation

(King, Roser & Jones, 2008). Prolactin can serve as a luteotropic agent in rat (Bouilly,

Sonigo, Auffret, Gibori & Binart, 2012), dog (Okkens, Bevers, Dieleman & Willemse,

1990), and spotted seal (Ishinazaka et al., 2002). In the rat, prolactin synthesizes progesterone and maintains the integrity of the corpus luteum (CL) (Freeman, Kanyicska,

Lerant & Nagy, 2000). In mares, a late luteal-phase prolactin surge may be associated with luteolysis (Roser, O'Sullivan, Evans, Swedlow & Papkoff, 1987). It can also be a luteostatic agent in some marsupials by limiting progesterone synthesis and secretion from the CL (Hinds & Tyndale-Biscoe, 2012). In the human, there is no evidence that prolactin plays a role in luteal development (Bouilly, Sonigo, Auffret, Gibori & Binart,

2012), but in human pregnancies, prolactin levels rise markedly at delivery (Kletzky,

Rossman, Bertolli, Platt & Mishell, 1985).

It is well documented that lactation, in particular the suckling stimulus, can inhibit folliculogenesis and ovarian function in most mammalian species (Bouilly, Sonigo,

Auffret, Gibori & Binart, 2012; McNeilly, Glasier, Jonassen & Howie, 1982; Rolland,

DeJong, Schellekens & Lequin, 1975; Wang, Hsueh & Erickson, 1980; Dorrington &

Gore-Langton, 1981; Jakubowski & Terkel, 1985). This is due in part to prolactin’s interference with follicle-stimulating hormone (FSH) on estrogen synthesis (Dorrington

& Gore-Langton, 1981) but primarily through suppression of LH secretion (McNeilly,

Glasier, Jonassen & Howie, 1982). Ovarian follicles may develop during lactation, but the increased prolactin levels from suckling can inhibit follicle maturation and result in an anovulatory state (Dorrington & Gore-Langton, 1981). In a study of captive lactating bottlenose dolphins, post-partum return to estrus occurred between 413 to 673 days (West et al., 2007). Studies of wild dolphins show that dolphins can continue lactation through ovulations and pregnancies (M. Yoshioka, K. Aida & I. Hanyu, 1989; West et al., 2000;

West et al., 2007), and this would allow the dolphin to maintain the average three year calving and three year nursing intervals (Wells, Scott MD & Irvine AF, 1987).

Maternal behavior in many species has been found to be induced and maintained by prolactin (Freeman, Kanyicska, Lerant & Nagy, 2000; Grattan, 2002). The maternal brain undergoes structural and functional modifications during pregnancy and lactation that ensure the demands of lactation can be met and to exhibit the appropriate behaviors to feed and care for the offspring (Pi & Voogt, 2000; Grattan, 2002). It has also been found that prolactin is associated with allomothering, when an individual other than the genetic parent provides care for conspecific young, in several bird and mammal species

(Ziegler, 2000; Soltis, Wegner & Newman, 2005; Schradin, Reeder, Mendoza &

Anzenberger, 2003). Allomothering behavior has been observed in wild and captive dolphins where non-genetic mothers escort a calf (Mann & Smuts, 1998; Simard &

Gowans, 2004; Tavolga, 1966; Gurevich, 1977; Leatherwood, 1977; Wells, 1991), while the genetic mother engages in social or resting behaviors (Mann & Smuts, 1998). Non- gravid, non-lactating dolphins in captivity have been documented to spontaneously lactate for a non-conspecific orphaned calf (Gaspar, Lenzi, Reddy & Sweeney, 2000)and have been induced to lactate after cohabitation with orphaned conspecific calves

(Ridgway, Kamolnick, Reddy, Curry & Tarpley, 1995). Howells et al (2009) observed a wild multiparous female adopt an orphaned calf and care for it for two years. Captive dolphins have also been observed to spontaneously lactate for non-orphaned calves in which they cohabitate (this paper).

Urinary prolactin has proven to be a reliable and non-invasive method to study prolactin dynamics in humans and primates (Alfredo Leanos-Miranda et al., 2008; Keely

& Faiman, 1994; Soltis, Wegner & Newman, 2005) (Alfredo Leanos-Miranda, Keely,

Soltis).

This study provides the first look at serial prolactin concentrations in the bottlenose dolphin and specifically during anestrus, estrous, pregnancy, lactation and when non-mothers cohabitate with mother-calf dyads.

3.2 Method

3.2.1 Experimental ethics

All animals, materials and methods used in the study were individually evaluated

by each facility’s Internal Animal Care and Use Committee (IACUC) to ensure safety,

health and well-being of the animals. Samples were taken during routine behaviors in

which the animals were conditioned to participate.

3.2.2 Animals

Bottlenose dolphins used in the study consisted of 7 females from 2 different

facilities (Table 1). Animals 1, 2 and 7 located at The Mirage Hotel (Mirage; Las Vegas,

Nevada USA), were housed in a 3,141,892-L manufactured seawater pool, maintained at

21 to 23° C year-round. Animals 1, 2 and 7 received a diet of frozen thawed herring (C.

harengus), capelin (M. villosus), sardine (Sardina pilchardus), and squid (L. opalescens)

for daily totals of 8.62, 7.26 and 8.16 kg, respectively. Animals 3-6, located at Six Flags

Discovery Kingdom (SFDK; Vallejo, California USA), were housed in a 2717.93-L

manufactured seawater pool maintained at 21-22o C year round. Animals 3-6 were fed a

diet of 10.4 and 5.9 kg, respectively, of whole frozen thawed herring (Clupea harengus)

and capelin (Mallotus villosus) daily.

Table 3.1 Animals in Study

Animal Birth Date Weight (kg) Repro History 1 5/1997 176 Nulliparous 2 3/2000 159 Parous 3 7/1979 184 Parous 4 7/1986 189 Parous 5 9/2004 137 Nulliparous 6 8/1979 196 Parous 7 11/1975 191 Parous

3.2.3 Statistics

Arithmetic means and standard deviations are presented as M +/- SD, and data

was tested for normality before calculating for comparisons, correlations or regression. P

values for prolactin and estradiol were calculated using student paired t tests, and P-

values <0.05 were considered significant. Correlations between estradiol and prolactin

were tested with Pearson’s Correlation Coefficient and presented as r = +/-. Polynomial

linear regression and studentized residual levels in regression diagnostics were used to

identify prl outliers during the follicular phase. Calculations were done using SYSTAT

software (Systat Software, Inc. 225 W Washington St., Suite 425, Chicago, IL 60606).

3.2.4 Behavioral conditioning

Operant conditioning techniques were used to train the dolphins for urine

sampling and ultrasound exams. Behaviors were positively reinforced and slow

approximations were taken until the dolphin was fully conditioned for daily sampling and

exams.

3.2.5 Urine collection

The dolphins were trained to urinate on cue. Initially the trainer placed firm hand pressure to the dolphin’s bladder, which often resulted in reflexive urination. This reflexive behavior was paired with primary reinforcement to increase the frequency. Over time, each dolphin was conditioned to urinate by the application of gentle pressure to the bladder. For animals 1, 2 and 7, the dolphins were partially pulled out of the water so that the genital slit was dry. First, the trainer stood on a dry ledge and grasped the tail flukes

of the dolphin that was stationary in a dorsal position floating at the pool surface. The dry

ledge was 7 to 10 cm higher than the water surface. Next, the trainer gently stepped

backward pulling the dolphin’s flukes backward and laterally over the ledge until the

dolphin’s genital slit was laterally out of the water. The dolphin’s head and pectoral

flippers remained in the water. The genital slit was wiped clean with gauze. A 50-ml

sterile plastic specimen cup was placed under the urethra and gentle pressure was applied

to the bladder. The urine was caught in the cup as it freely flowed from the urethra. For

animals 3-6, urine collection was achieved by placing the dolphin in a ventral position

and the genital slit was wiped clean with a dry cloth. A sterile plastic 25-ml syringe was

used to draw up the urine once it free flowed from the urethra. An average of 5-10 ml of

urine was collected from all study animals. Urine was immediately frozen at -17° C

(animals 1, 2 and 7) and -70° C (animals 3-6) in 3-5 ml increments in plastic storage

vials. Urine for the study was collected in the am (8 am–11 pm) for the study unless

noted.

3.2.6 Hormone testing

3.2.6.1 Progesterone and estradiol

Urinary progesterone (P) and estradiol-17 beta (E2) was measured using an

Electrochemiluminescent Immunoassay (ECLIA) tested with the Elecsys 2010 instrument

(Roche Diagnostics, Mannheim, Germany) at Clinical Pathology Laboratory in Las

Vegas (4275 S. Burnham Ave, Suite 325, Las Vegas, NV 89119, USA) as previously reported (Muraco et al., 2009). Urine was shipped frozen to the laboratory from each facility.

3.2.6.2 Prolactin

Dolphin urinary Prolactin (Prl) concentrations in urine were determined by a modified homologous, double antibody RIA according to the methods of Roser et al.

(1984). Briefly, purified equine PRL (supplied by A.F. Parlow, National Hormone and

Peptide Program, Torrance, CA) was used for standards (0.125–32 ng/mL, AFP-7730B) and for iodination (AFP-8794B) by the iodogen method (Matteri, Roser, Baldwin,

Lipovetsky & Papkoff, 1987). Primary antibody was rat-anti-equine-prolactin antibody

(Parlow AFP-361687R; 1:38,000), second antibody was goat anti-rat IgG (Antibodies,

Inc., Davis, CA; 1:300). Sensitivity of the assay was 0.25 ng/mL, and inter- and intra- assay coefficients of variation were 12.7% and 4.9%, respectively. The standard curve was calculated using the four parameter logistic transformation technique, and sensitivity of the assay was 0.5 ng/ml.

3.2.6.3 Creatine (Cr)

For prolactin assays, concentrations of urinary hormones and metabolites were expressed as ng of prl per mg Cr excreted (Taussky, 1954).

3.2.7 Ultrasonography

A Sonosite Titan portable ultrasound machine with a 5-2 MHz curvilinear transducer (Sonosite Inc. 21919 30th Drive SE, Bothell, WA 98021-3904, USA) was used for the real-time B-Mode trans-abdominal ultrasound imaging. Images were digitally captured using the internal digital memory of the machine or by the use of an

Archos 605 Audio/Visual Player (Archos Inc. 7951 E.Maplewood Avenue #260

Greenwood Village, CO 80111 USA). For all animals, ultrasound exams were conducted as needed and exams typically lasted 3-5 minutes. For ovary and fetus visualization, the dolphins were trained to float laterally and stationary at the water’s surface. The blowhole was submerged, but the animal could lift her head and take a breath as needed. To visualize the ovaries and uterus, the ultrasound transducer was placed between the junction of the rectus abdominus muscle and the hypaxialis lumborum muscle (Brook,

2001). For mammary gland ultrasound, the dolphins were trained to float ventral up and remain stationary at the water’s surface. The blowhole was submerged, but the animal could maneuver to a lateral position and lift her head to take a breath as needed. To visualize the mammary gland parenchyma, the transducer was placed transversely on the dolphin approximately 2 cm anterolateral to the anterior line of the genital slit and moved cranially until the parenchyma was identified (Figure 1). A transverse diameter measurement was taken in the area of the medial parenchyma to record changes in size.

Figure 3.1 Starting position of ultrasound transducer for mammary gland parenchyma visualization

White arrows indicate the paired mammary slits

1.1.1.1 Anestrus

Dolphins can experience long term anestrus (>1 year) even when not post-partum or lactating (Sawyer-Steffan, Kirby & Gilmartin, 1983; Yoshioka, Hori, Tobayama, Aida

& Hanyu, 1986). In this study, anestrus was determined by no elevation in P and no follicular development on the ovary. Additionally, animals categorized as anestrus were also non-gravid, non-lactating and not living with a mother-calf dyad.

3.2.7.2 Seasonality and prolactin

Bottlenose dolphins can give birth in any season in aquaria (Urian, Duffield,

Read, Wells & Shell, 1996; Muraco, Clough, Teets, Arn & Muraco, 2010), and display a

broad, diffuse and flexible reproductive seasonality in the wild giving birth in all seasons

with a slight increase in births in spring and fall (Urian, Duffield, Read, Wells & Shell,

1996; Wells, Scott MD & Irvine AF, 1987; Victoria G. Thayer et al., 2003). Prl levels

were examined during Winter, Spring Summer and Fall in anestrus females to examine potential seasonal Prl changes.

3.2.7.3 Estrous

Analysis of serial P and E2 was used to determine follicular and luteal phases for animals 1 and 2 and ovarian ultrasound, P and E2 was used for animals 3 and 4.

3.2.8 Pregnancy

To determine pregnancy, P was analyzed for elevated levels and uterine

ultrasound was used to verify uterine fluid and fetal development. For the purposes of

this study, pregnancy was considered to have begun 20 days after ovulation.

3.2.9 Lactation

Daily observations were conducted on post-partum study females (animals

2,3,4,7) and calves for nursing frequency. Calves of mothers 3 and 4 were monitored for

24 hours during the first 15 days of life. Each nursing event was timed using a stopwatch

and recorded onto a spreadsheet showing duration. Non-mothers (animals 1,2,5,6), while

cohabitating with a calf, were periodically checked for signs of lactation through manual

expression of the mammary gland for milk secretion. Mammary gland expression

consisted of gently applying pressure anterior to the mammary slit and during lactation

milk would be secreted from the slit.

3.2.10 Cohabitation with mother and calf pairs

Females (1,2,4,5, and 6) were deemed non-gravid, non-lactating and anestrus

before moving them into a pool with a mother-calf dyad. Females 1 and 2 lived with a

newborn calf for 4 months, female 6 lived with two newborn calves for 4 months, female 58

5 lived with two 3 month old calves for 3 months and female 4 lived with an 18 month old calf for 2 months.

3.3 Results

Prolactin was detectable in dolphin urine from all 7 study animals (range 0.118–

3.790 ng prl/mg Cr). The lowest level was recorded during the follicular phase and the

highest level during the last trimester of pregnancy. Prl M +/- SD for each reproductive

state, lactation and cohabitation can be seen in table 2. Figure 2 shows mean and

maximum Prl levels during each studied phase. Figure 3 shows mean Prl during each

reproductive state and lactation.

Table 3.2 Study phases, reproductive state, lactation and cohabitation

Reproductive State, Lactation and uPrl Cohabitation with Mother/Calf pair Dolphin N N Mean +/- SD Anestrusa 3 37 0.757 +/-0.260 Follicular 4 61 0.510 +/-0.239 Luteal 3 41 0.578 +/-0.211 Pregnancy First Trimesterb 2 28 0.692 +/-0.180 Pregnancy Second Trimester 2 6 0.826 +/- 0.336 Pregnancy Third Trimester 3 25 0.957 +/- 0.617 Lactation 1 Month Post-Partum 1 3 0.785 +/- 0.228 Lactation 2 Months Post-Partum 3 16 1.100 +/- 0.304 Lactation 3 Months Post-Partum 3 8 1.852 +/- 0.971 Lactation 4 Months Post-Partum 3 3 1.450 +/- 0.288 Lactation 5 Months Post-Partum 1 1 2.440 Lactation 15 Months Post-Partum 1 5 0.986 +/- 0.182 Lactation 16 Months Post-Partum 2 4 0.844 +/- 0.440 Lactation 17 Months Post-Partum 2 4 0.806 +/- 0.343 Lactation 18 Months Post-Partum 1 2 0.416 +/- 0.158 Cohabitatec Days 1-30 4 84 0.706 +/- 0.408 Cohabitate Days 31-60 4 83 0.758 +/- 0.396 Cohabitate Days 61-127 1 16 1.050 +/- 0.495 Cohabitate with older calfd 1 6 0.213 +/- 0.047 a. No progesterone elevation no follicular development. b. Each trimester consists of 4 months. c. Non-gravid, non post-partum female cohabitate with calf < 3 months and the calf’s mother. d. Non-gravid, non-post-partum female cohabitate with 18 mo calf and the calf’s mother.

Figure 3.2 Prl means and maximum levels during studied phases a. Pregnant: Months 1-12 (n=62 0.849+/-0.458). b. Lactating: Months 1-5 (n=28 1.329+/- 0.700). c. Cohabitation: Days 1-127 (n=183 0.760+/-0.419).

Figure 3.3 Prl means during reproductive states and lactation.

1.1.1 Anestrus

Three dolphins (animals 1,2 and 6) were studied during anestrus (non-gravid, not post-partum or lactating and not cohabitating with a mother-calf dyad). Levels of prolactin were significantly higher during anestrus than the follicular and luteal phases (p

0.022 and p 0.000) (Figure 2) and cohabitation with a mother/calf pair for the first 30 days (p 0.000). Prolactin levels during anestrus were significantly lower than months 1-5 of lactation (p 0.000) (Figure 3) and days 60-127 of cohabitation with a mother/calf pair

(p 0.008) (Figure 2). Additionally, all study females were anestrus when cohabitating with a mother-calf dyad or when post-partum and lactating.

3.3.2 Seasonality and prolactin

Anestrus occurred during each season and prl levels were compared using paired t tests. There was no significant difference in anestrus prl levels between seasons. Prl means during each season are as follows: Winter (n=2 0.710+/-0.269), Spring (n=20

0.744+/-0.266), Summer (n=15 0.795+/-0.270) and Fall (n=2 0.640+/-0.057).

3.3.3 Estrous cycle

4 estrous cycles (animals 1,2,3 and 4) were studied. Estrous cycles occurred

synchronously in September for females 1 and 2 with ovulation occurring 17 and 14 days

respectively following removal from a mother-calf dyad. Female 4 cohabited with female

3 and her nursing 16 month old calf for two months. At 18 months, the calf was weaned,

and females 3 and 4 synchronized estrous in May with female 3 ovulating 22 days

following weaning and female 4 ovulating 30 days following the calf removal.

Follicular phases averaged 10 days and luteal phases averaged 20 days as previously reported (Muraco et al., 2009; Muraco H, Clough P, Teets V, Arn D &

Muraco M., 2010; Robeck, McBain, Mathey & Kraemer, 1998). Figure 4 shows Prl, E2 and P during the estrous cycle of animal 2.

Figure 3.4 Prl, uE2 and uP during the estrous cycle of animal 2.

*Ovulation

3.3.4 Follicular phase

Mean Prolactin levels were significantly lower during the follicular phase vs

anestrus (p 0.022), cohabitating with a mother-calf dyad (p 0.005) or when nursing a 15-

18 month old calf (p 0.073). However, during the follicular phase, prolactin rises were

detected and can be seen in Table 3 and figures 5-7.

Table 3.3 Prolactin rises detected during the follicular phase

Day of estrous Animal prl was elevated

1 -8, -1 2 -6 3 -7,-4 4 -5

Figure 3.5 Follicle growth and studentized residual prl rises during the follicular phase

Figure 3.6 Identification of Prl outliers using polynomial regression during the follicular phase

Figure 3.7 Ultrasound of follicular dynamics in animal 3 during prl rises a. Day -9, start of follicular phase, Prl 0.3623 ng/mg Cr. Blue circle indicates ovary. White arrow shows new follicle developing. b. Day -7, dominant follicle 1.2 cm dia, Prl 0.6029 ng/mg Cr. White arrows >follicle. c. Day -4, dominant follicle 1.6 cm dia, Prl 0.5921 ng/mg Cr. d. Day -1, periovulatory follicle 2.0 cm dia, Prl 0.3564 ng/mg Cr

3.3.5 Luteal phase

There was a slight increase in Prl means in the luteal phase vs the follicular phase

although it wasn’t significant (p 0.2957). However, there was a significant difference

between the luteal phase (first 20 days following ovulation) and first trimester of

pregnancy (21–120 days following ovulation) (p 0.0034).

3.3.6 Estradiol

Mean E2 from animal 2, was calculated during anestrus (n=10, 73.7000 +/-

17.8450), follicular phase (n=17, 99.235 +/- 32.011), cohabitation with a mother-calf

dyad (n=6, 79.667 +/- 16.379) and during the final 17 days of pregnancy (n=17, 139.706

+/- 91.649). Estradiol was significantly higher in the follicular phase vs anestrus (p

0.0305) and during the last trimester of pregnancy vs anestrus (p 0.0547). There was no

significant difference in estradiol levels between the follicular phase and the last trimester

of pregnancy (p 0.1333) or between cohabitation and anestrus (0.8609). Correlations

between E and Prl during anestrus (positive), cohabitation (neutral) and follicular phase

(negative) can be seen in figure 8 and during the final 17 days of pregnancy (negative) in

figure 9.

Figure 3.8 uE2 and prl during anestrus, cohabitation, follicular phase and ovulation a. anestrus (r= 0.3450). b. cohabitation with mother/calf (r= 0.0714). c. follicular phase and ovulation (r= -0.1111).*ovulation Place all detailed caption, notes, reference, legend information, etc here

Figure 3.9 E2 and prl in the final days leading to parturition

(r= -0.4232)

3.3.7 Pregnancy

Pregnancy was confirmed in animals 2,3 and 4 when a live fetus was visualized on ultrasound. Animal 1 aborted placental tissue positive for maternal and paternal DNA

(Duffield & Chamberlin-Lea, 1990) after 3 months of pregnancy.

Prl means increased with each trimester of pregnancy (Table 2). The last month of pregnancy showed the highest Prl means (n=3 m 1.073 +/- 0.038) (Figure 3).

There was a significant difference in Prl levels (p 0.0000) the first 55 days following ovulation between pregnancies of animal 1 (abnormal pregnancy) and animal 2

(normal pregnancy) (Figure 10).

Figure 3.10 Prolactin levels during the first trimester of pregnancy in a normal and abnormal pregnancy

*Abortion after abnormal pregnancy

3.3.8 Lactation

3.3.8.1 Post-partum

Post-partum mothers 2,3,4,7 all cared for and nursed their calves. Prl means

during different post-partum months of lactation can be seen in table 2. Prl was higher in

months 1-5 (n=31 1.341 +/- 0.669) vs months 15-18 (n=15 0.824 +/- 0.334) of lactation

(p 0.0897). During the first 5 months, mean prl levels were highest during month 3 (n=8

1.852 +/-0.971), however a higher singular level from month 5 (n=1 2.44) was recorded.

Figure 3 shows mean prolactin levels during early (months 1-5) and late (months

15-18) of lactation and how it compares to anestrus, estrous and pregnancy.

3.3.8.2 Nursing durations

The mean nursing duration per latch from two calves was n=4,481 5.718 +/-

2.602 seconds. Figure 11 shows mean nursing durations in minutes and number of latches

in a 24 hour period for the first 15 days of life from two calves.

Figure 3.11 Minutes of nursing and number of latches in a 24 hour period during the first 15 days of life for two calves

3.3.8.3 Lactation in non-mothers

Milk was expressed in non-mother 2 after three months of cohabitation with a calf but was not observed nursing the calf. Milk production was not identified in non-mothers

1, 4, 5 and 6. Figure 12 shows a comparison of Prl levels between animal 1 (non- lactating) and animal 2 (lactating) while cohabitating with a mother-calf dyad.

Figure 3.12 Prl levels during cohabitation with a mother/calf pair a. No calf. b. Calf born in adjacent pool to dolphins 1 and 2. c. Dolphins 1 and 2 cohabitate with newborn calf and calf’s mother. *Milk expressed from dolphin 2.

3.3.9 Cohabitation

Non-mothers 1,2,4,5,6 were all opportunistically observed interacting with

mother-calf dyads through group synchronized swimming and escorting the calves

without the mothers (Figure 13). No formal behavioral observations were conducted.

Figure 3.13 Non-mother cohabitation with mother-calf dyads a. Mothers and calves swimming in echelon position with non-mother (arrow) underneath. b. Mothers and calves swimming in slipstream position with non-mother carrying a toy. c. Non-mother with calf swimming in echelon. d. Non-mother with calf

Prl means for dolphins cohabitating with newborn calf/calves for 3 months

(animals 1,2) and 4 months (animal 6) are as follows: animal 1 (n=63 m 0.497 +/- 0.297), animal 2 (n=63 m 0.782 +/- 0.302), and animal 6 (n=43 m 1.010 +/- 0.424). Animal 5

who cohabitated with two 3 month old calves for three months (n=14 m 1.069 +/- 0.615) and for animal 4 who cohabitated with one 18 month old calf for 2 months (n=6 m 0.212

+/- 0.047). Figure 12 shows serial prl levels from animals 1 and 2 before and after

cohabitation with a calf.

Cohabitation with calves from newborn to 6 months had prolactin levels significantly higher than cohabitation with an older calf of 16 months (p 0.050). There was not a significant difference between days 90-127 of cohabitation with a mother/calf pair and months 1-5 of lactation (p 0.2253).

3.3.10 Mammary gland ultrasound

The dolphin has paired mammary glands with external openings located in two indented slits on either side of the genital groove. Connective tissue septa divide the parenchyma into lobules (Ridgway, 1972). Swollen dolphin mammary glands are easily visualized in lactating post-partum mothers anterolateral to the genital groove. Mammary gland ultrasound was performed on animal 3 (Figure 1), 5 and 6.

Animal 3 was monitored with ultrasound to identify changes in mammary gland parenchyma size and echogenicity during various stages of lactation concurrent with prolactin levels. Table 4 and Figure 14 shows Prl means and parenchyma size during resting, intermediate and lactating stages.

Table 3.4 Mammary gland size and state during pregnancy, lactation and post- weaning estrous

Stage Gland Statea Size cme Prl +/- SD

4 m preg Resting 1.2 0.0692 +/- 0.180 11 m preg Intermediate 3.2 0.957 +/- 0.617 6 d before parturition Lactating 3.8 0.834 +/- 0.118b 3 m lactation Lactating 4.8 1.852 +/- 0.971 18 m lactation Lactating 5.0 0.416 +/- 0.158 9 days post wean, no suckle Lactating 4.0 0.4756c 15 days post wean, cycle day - Intermediate 3.9 0.6029c 7 Cycle day +6 Intermediate 3.0 0.5195d a. Lactating: milk secreting; Intermediate: glandular lobules developed but not as developed as in lactation; Resting: involution of gland (Flock & Winter, 2006). b. Prl mean final 7 days before parturition (n=7) in animal 2. c. Prl level for the same day as ultrasound in animal 3. d. Prl level 8 days post ovulation in animal 3. e. Medial parenchyma cross sectional diameter.

Figure 3.14 Ultrasound of the mammary parenchyma

. White circle indicates mammary gland, white arrow shows lactiferous ducts. a. Resting mammary gland, 4 months pregnancy, Homogenous and small. Only hyperechoic ligaments and border visible.b. Lactating, 1 week before birth, homogenous and hyopechoic with small anechoic lactiferous ducts and hyperechoic ligaments.c. Lactating, 18 month post-partum, homogenous echogenicity with small anechoic lactiferous ducts and hyperechoic ligaments and parenchyma border. d. Lactating, no suckle two days, multiple large anechoic lactiferous ducts.

The parenchyma of a healthy mammary gland appears as a homogenous structure of average echogenicity with anechoic blood vessels and lactiferous ducts (Flock &

Winter, 2006).The non-lactating mammary parenchyma in the dolphin is difficult to visualize appearing with a homogenous echogenicity and hyper echoic border. Distinct hyperechoic ligaments can be visualized in the transverse view and these were used to identify the location of the mammary gland in non-lactating dolphins. The lactating mammary parenchyma was easy to visualize appearing with a hypoechoic homogenous

echogenicity with small anechoic lactiferous ducts and hyperechoic ligaments and border.

Immediately following weaning of an 18 mo calf, lactiferous ducts filled due to the lack of nursing (Figure 14d).

Animals 5 and 6 were monitored via ultrasound for signs of mammary development while cohabitating with a calf but development and lactation was not detected.

3.4 Discussion

This study aimed to provide preliminary data on prolactin’s role in the female

bottlenose dolphin.

3.4.1 Pregnancy

Prl levels rose as pregnancy progressed with the highest levels occurring during

the last month. This differs from findings in rats where prl levels remain low during the

latter half of pregnancy (Morishige, Pepe & Rothchild, 1973; Amenomori, Chen &

Meites, 1970) but similar to humans where there is an increase in prl as pregnancy

progresses with a peak at delivery (Hwang, Guyda & Friesen, 1971; Kletzky, Rossman,

Bertolli, Platt & Mishell, 1985). Based on ultrasound, the dolphin mammary gland

enlarges and changes in echogenicity during the last month of pregnancy so increased prl

levels are most likely involved in lobuloalveoli development (Harris et al., 2006).

Prl levels during the first trimester were higher than during the luteal phase and

higher in normal healthy pregnancies vs the abnormal/unhealthy pregnancy. This

suggests that prl may play a role in corpus luteum maintenance during pregnancy such as

been found in ferrets and dogs (Murphy, 1979; P.W. Concannon, V.D. Castracane, M.

Temple & A. Montanez, 2009).

3.4.2 Lactation

Dolphin mother and calf nursing behaviors have been well documented in the literature (Cockcroft & Ross, 1990; Mann & Smuts, 1998; Mann, 1999; Wells, Scott MD

& Irvine AF, 1987; Peddemors, Fothergill & Cockcroft, 1992; A. Eastcott & T.

Dickinson., 1987; Reid, Mann, Weiner & Hecker, 1995), and dolphins have been

compared to primates in terms of long term young dependency (Oftedal, 1997). Wild

dolphin mothers will typically nurse calves for three years (Wells, Scott MD & Irvine

AF, 1987; Oftedal, 1997) although it has been documented to continue for over 5 years

(Wells, Scott MD & Irvine AF, 1987; Mann, Connor, Barre & Heithaus, 2000). The need

to teach young how to forage in a challenging environment may influence lactation duration in cetaceans (Whitehead & Mann, 2000).

Dolphin calf nursing behavior has been described as suckling bouts which includes multiple 5 second nipple latches within a 1 to 5 minute period, followed by an interval phase where no suckling occurs (Eastcott & Dickinson, 1987). During the first two weeks of life, a neonate dolphin will nurse in 5 second intervals totaling over 30 minutes and over 100 nipple latches in a 24 hour period (Figure 11). However, after the first two weeks, total nursing durations drop off dramatically (A. Eastcott & T.

Dickinson., 1987; Peddemors, Fothergill & Cockcroft, 1992; Reid, Mann, Weiner &

Hecker, 1995). By month three, nursing durations may be less than 7 minutes in a 24 hour period (Peddemors, Fothergill & Cockcroft, 1992). Interestingly, Prl levels were higher in months 3-5 of post-partum lactation than months 1 and 2 when nursing 77

durations were more frequent (Table 2 and Figure 3). During the first 20 days post- partum, milk is spontaneously expressed into the water but as lactation continues, spontaneous milk discontinues (Peddemors, Fothergill and Cockcroft, 1992). Peddemors,

Fothergill and Cockcroft (1992) also noted that after the first month of nursing, calf suckling was often preceded by the calf massaging the mammary glands with the dorsal fin or top of the melon and this occurred more often before the first suckle of each bout.

It has been suggested that the massaging behavior may be required to initiate milk letdown as lactation stages progress (Cockcroft & Ross, 1990). In piglets, mammary massage prior to suckling causes an increase in prolactin levels (Algers, Madej,

Rojanasthien & Uvnäs-Moberg, 1991). It’s possible that after the first month of

spontaneous milk flow ends and as dolphin calf mammary massage increases/improves,

that could account for increased prl levels in months 3-5. Additionally, numerous studies

have been conducted on the milk composition of the bottlenose dolphin and show a high

level of variance in composition depending on the stage of lactation (L Eichelberger, E.

S. Fetcher, E. M. K. Geiling & and B. J. Vos, 1940; Ackman, Eaton & Mitchell, 1971;

Yablokov, Belkovich & Borisov, 1974; Pervaiz & Brew, 1986; Peddemors, de

Muelenaere & Devchand, 1989; Ridgway, Kamolnick, Reddy, Curry & Tarpley, 1995;

West et al., 2007; Oftedal, 1997). As lactation stages progress, there is a tendency

towards higher fat and protein and less water content (West et al., 2007).

Overall, mean Prl levels were higher during the first 5 months of post-partum

lactation vs months 15-18. Captive dolphin calves in this paper began eating fish as a

supplement to nursing at approximately 6 months of age and by 18 months were

receiving 3% of body weight in fish. This may reduce the amount of nursing by an older calf and also subsequently lead to lower prl levels.

Mammary gland ultrasound provided a valuable tool to monitor changes over the stages of lactation, through weaning and to check for lactation in non-mothers.

Echogenicity changes and size of the parenchyma showed a positive correlation with changes in prolactin levels (Figure 7).

3.4.3 Cohabitation

In this study, when non-mothers cohabitated with mother-calf dyads they were

assumed to allomother in some aspect. Although no formal behavioral observations were

conducted, opportunistic observations indicated that non-mothers interacted positively

with mother/calf pairs and/or calves alone (Figure 13). Prolactin levels rose markedly

when non-mothers cohabitated with mothers and young calves of less than 6 months for

at least 60 days. One 7 year old nulliparous study female (animal 2) began lactating after

approximately 3 months of cohabitation with her mother and her mother’s newborn calf.

No observations were made of the non-mother nursing the calf although she was

observed escorting the calf. This is consistent with Mann and Smuts, 1998, finding that

wild nulliparous inexperienced female dolphins were more likely to allomother than

parous females. As with primates, (Ziegler, 2000; Soltis, Wegner & Newman, 2005;

Schradin, Reeder, Mendoza & Anzenberger, 2003), prolactin appears to be associated

with maternal behavior in dolphins in both the mother and associated non-mothers.

Additionally, based on the nulliparous female that began lactating, prolactin is most

likely involved in dolphin spontaneous and/or induced lactation. Each study dolphin

experienced anestrus while cohabitating with a mother/calf pair. 79

3.4.4 Estrous Cycle

During the follicular phase, all four dolphins experienced a rise in prolactin between days–8 and -5 and two dolphins experienced a second rise on days -4 and -1

(Table 3). The prolactin rise was positively correlated with selection of a dominant

follicle in animal 3 (Figure 7). This suggests that prolactin may play a role in follicular

maturation as seen in horses (Bennett-Wimbush, Loch, Plata-Madrid & Evans, 1998;

King, Roser & Jones, 2008). Prolactin was negatively correlated with estradiol during the

follicular phase, however Figure 8 shows that the prl rise at day –6 in animal 2 came the

day after a rise in estradiol. So it is possible that the rise in prl was in part due to a

previous rise in estradiol.

Prolactin is an important regulator in seasonal breeding species. There is a clear

seasonal correlation of prolactin in horses and sheep with elevated prolactin during the

breeding season of horses and low prolactin levels during the breeding season of sheep

(Gerlach & Aurich, 2000). In aseasonal domestic cattle, there is an annual prolactin

rhythm related to photoperiod and temperature with peaks in the summer (Tucker, 1982;

D. Petitclerc et al., 1983). In contrast, there is no is no correlation between prolactin and

photoperiod in humans or domestic pigs (Reinberg et al., 1978; Martikainen, Tapanainen,

Vakkuri, Leppäluoto & Huhtaniemi, 1985; Ravault et al., 1982). Dolphins do not appear

to have an annual prolactin rhythm and may not use photoperiodic cues for reproductive

regulation.

This study showed an association between weaning and a return to estrous similar to pigs (Armstrong & Britt, 1985) as well as a similar “male effect” where boar contact with weaned sows can advance estrus by several days (Langendijk, Soede & Kemp,

2000; Pearce & Pearce, 1992). The male effect in terrestrial mammals involves an external chemical stimulus that immediately modulates the activity of the hypothalamic gonadotrophin-releasing hormone pulse generator (Hamada, Nakajima, Takeuchi &

Mori, 1996). After spending three months with a mother/calf dyad, animals 1 (non- lactating) and 2 (lactating non-mother) were moved into a social group with an adult male with whom they had previously been living separately from. Ovulation occurred in 14

(animal 2) and 17 (animal 1) days and both animals conceived from natural breeding.

This quick time to ovulation in the presence of a mature male following weaning is similar to the “male effect” seen in pigs. In contrast, animals 3 (lactating mother) and 4

(non-lactating) lived in a social group with no adult male following the weaning of animal 3’s calf. Ovulation occurred 22 and 30 days respectively following weaning, and both animals conceived from artificial insemination. This slower post-weaning ovulation could be the result of not having contact with a mature male. Prolactin levels were elevated above basal levels (Table 2) due to lactation or cohabitation with a calf, prior to

3 of the 4 studied estrous cycles (animals 1,2 and 3). Removal of the animals from the respective calves resulted in a drop in prolactin levels and a return to estrous (Figures 8 and 12). Animal 4 did not have markedly elevated prolactin levels when living with the

16 month old calf prior to removal from the calf, and she took the longest time to ovulation at 30 days. There was also a natural paired estrous synchrony between animals

1 and 2 and animals 3 and 4. Natural estrous synchrony irrelevant to seasonality has been documented in elephants (Weissenböck, Schwammer & Ruf, 2009; Rasmussen &

Krishnamurthy, 2000), and typically pheromonal and/or odorous cues are the mechanisms by which synchronicity is achieved (Rasmussen & Schulte, 1999; Stern &

McClintock, 1998; McClintock, 1998). There have been suggestions that dolphins may have the ability to use chemoreception in communication (Yablokov, 1961; Yablokov,

Belkovich & Borisov, 1974), and dolphins have the ability to taste salty, sweet, sour and bitter, with most sensitivity to bitter (Nachtigall & Hall, 1984; Nachtigall, 1986). Dolphin urine, semen, blood, feces, prostate glandular extract and perianal glandular secretions were analyzed using gas chromatography and mass spectrometry in a study by the Naval

Ocean Systems Center to identify chemoreceptively active compounds (Ceruti,

Fennessey & Tjoa, 1985). That study found that dolphin urine contained compounds that were sweet, sour and bitter, and that one or more of these compounds could account for the dolphin’s ability to detect urine during behavioral taste studies (Kuznetzov, 1990). It is unknown how, or to what extent, dolphins may utilize this sense in their daily life, and it is also unknown how dolphins signal reproductive condition to one another.

3.4.5 Anestrus

The positive correlation between prolactin and estradiol during anestrus is

interesting and suggestive that the prolactin rhythm during anestrus could be resultant of

an estradiol rhythm (del Pozo & Brownell, 1979). Anestrus in dolphins could be similar

to swine where estradiol provides a negative feedback and prevents FSH and LH release

(Almond & Dial, 1990).

All study females remained anestrus while lactating or when cohabitating with a

mother-calf dyad and none of the females had access to a mature male during that time

period. Previous studies that showed a return to estrous while lactating (M. Yoshioka, K.

Aida & I. Hanyu, 1989; West et al., 2000; West et al., 2007) all involved the lactating

females having or gaining access to mature males. This may be further evidence of a possible “male effect” on female dolphin reproduction.

3.5 Conclusion

Prolactin is detectable in dolphin urine, and levels during pregnancy are higher

than during the luteal phase. Normal term pregnancies have higher levels than those in

abortive pregnancies which may indicate a role in luteal maintenance. There is a prolactin

rise at the end of gestation that is most likely involved in mammary development.

Prolactin levels become elevated in both nulliparous and parous non-mothers when they

cohabitate with mother-newborn calf dyads. During lactation, prolactin levels are higher

during months 1-5 of lactation vs months 15-18. Anestrus dolphins have higher prolactin

levels than during the follicular phase however, there are prolactin rises that occur during

the follicular phase that may assist in follicle maturation. There is a similarity between

dolphins and sows with a return to estrous following weaning, and dolphins experience

lactational anestrous. A male effect was observed that shorted the days to ovulation

following weaning when females cohabitated with a mature male. Spontaneous lactation

occurred in a nulliparous female when cohabitating with a mother-newborn calf dyad and

was positively correlated with elevated prolactin levels. Finally, there was no positive

correlation between season and prolactin levels suggesting that dolphins may not use

photoperiod in reproduction regulation as is observed in horses and sheep.

CHAPTER IV

CONCEPTIVE ESTRUS BEHAVIOR IN THREE BOTTLENOSE DOLPHINS

(TURSIOPS TRUNCATUS)

Muraco, H. & Kuczaj, S. (2015). Conceptive Estrus Behavior in Three Bottlenose Dolphins (Tursiops truncatus). Animal Behavior and Cognition. 2, 30–48.

4.1 Introduction

Little is known about conceptive dolphin estrus behavior despite the fact that dolphins routinely engage in overtly sexual interactions (Ridgway, 1972). Most observations of sexual behavior involve socio-sexual behaviors rather than true conceptive mating behaviors (Connor et al., 1992; Dudzinski, 1998; Puente & Dewsbury,

1976; Wells, 1984). Dolphin non-conceptive socio-sexual behaviors may involve immature individuals, same sex individuals, and copulation during the non-conceptive period (Furuichi, Connor, & Hashimoto, 2014). In addition, male and female dolphins masturbate with objects and the appendages of other dolphins (Ridgway, 1972).

Reflexive penile erection occurs in male dolphin calves within 48 hours of birth, and male calves are able to assume the copulatory position within weeks (Ridgway, 1972). A dependent male calf will often copulate with his mother (Ridgway, 1972). Dolphins are not the only animals to participate in non-conceptive sexual behaviors. Many social mammals engage in these sorts of socio-sexual behaviors, including mounting by or on

pre-pubertal animals, mounting between same sex animals, mounting without thrusting, and intromission and ejaculation (Hanby & Brown, 1974).

There are few studies that document conceptive dolphin sexual behavior. Connor et al. (1992) noted that male bottlenose dolphins (Tursiops sp.) in Shark Bay Australia

form alliances to cooperatively herd and control females. Reproductive states of the

herded females could not be determined, but the authors hypothesized that the females

were in estrus. Dudzinski (1998) studied Atlantic spotted dolphins (Stenella frontalis) in

the Bahamas, and noted that much of the affliative social behavior included genital

oriented behaviors, rubbing and body to body contact. Most of the socio-sexual behaviors

in this study occurred between animals of the same age range and of the same sex, but

reproductive states of the animals could not be specified.

In a study of captive spinner dolphins (Stenella longirostris), tactile, contact and

socio-sexual behaviors were observed along with one estradiol sample (Wells, 1984). The

sexual behaviors observed appeared to serve social functions rather than reproductive

ones (Wells, 1984). Puente and Dewsbury (1976) conducted observations on captive

bottlenose dolphins and documented nine potential courtship patterns that involved

mouthing, nuzzling, rubbing, stroking, leaping, chasing, head butting, display and yelping

as well as copulation. However the reproductive status of the female dolphins was

unknown. Female dolphins being monitored for artificial insemination showed estrus

behaviors of “listing on the water surface, sinking and reduced responsiveness during

training sessions” 12-48 hrs prior to ovulation (Robeck et al., 2005, p. 668).

In mammals, the period of female sexual receptivity, called estrus, heat, or

lordosis occurs when the female will accept the male for copulation (Bearden, Fuquay, &

Willard, 2004; Hunter, 1980; Pineda & Dooley, 2003), and estrus behaviors imply the act of mounting, intromission and pelvic thrusts (Austin & Short, 1972). Additionally, estrus behaviors in many mammals may include immobility, raising the hindquarters, arching the back, restlessness, irritability and excitability (Bearden et al., 2004; Hunter, 1980).

Cows in estrus will solicit mounts and mount other cows, smell the vulva of other cows, raise and switch their tail and spend more time walking and less time resting and feeding

(Bearden et al., 2004). Estrus pigs will assume a rigid stance, called the lordosis reflex, when pressure is applied to their back (Austin & Short, 1972).

To truly understand if the sexual behaviors of dolphins are for conceptive purposes, serial endocrinology and known reproductive states are critical, and to date no detailed sexual behavioral study has been conducted on dolphins with a known reproductive state. The work presented in this paper is the first to analyze the sexual behavior of bottlenose dolphins at two different aquaria during three conceptive estrous cycles, two (one nulliparous and one parous) from natural breeding and one (nulliparous) from artificial insemination.

4.2 Method

4.2.1 Experimental ethics

All animals, materials and methods used in the study were individually evaluated

by the facility’s Internal Animal Care and Use Committee (IACUC) to ensure safety,

health and well-being of the animals. Samples were taken during routine behaviors in

which the animals were conditioned to participate.

4.2.2 Animals

Animals 1-5 (Table 1) were housed at facility A, a 3,141,892-L manufactured

seawater pool, maintained at 21 - 23° C year-round. Animal diets consisted of frozen

thawed herring (C. harengus), capelin (M. villosus), sardine (Sardina pilchardus), and

squid (L. opalescens) for daily totals of 5–10 kg per animal. Animals 6 and 7 (Table 1)

were located at facility B and housed in a 2717.93 L manufactured seawater pool

maintained at 21-22o C year round and fed a diet of whole frozen thawed herring (Clupea

harengus) and capelin (Mallotus villosus) ranging from 8–15 kg daily.

Table 4.1 Animals in study

Animal Facility Sex Birthdate Weight (kg) Repro Hist

1 A F 11/1975 191.86 Parous 2 A F 5/1997 176.44 Nulliparous 3 A F 3/2000 159.66 Immature 4 A M 2/2003 165.15 Immature 5 A M 8/1989 201.35 Proven Sire 6 B F 9/2004 134.00 Nulliparous 7 B F 8/2003 163.00 Nulliparous 8 B M 11/1969 210.00 Proven Sire 9 B F 7/1998 220.00 Parous 10 B F 10/2005 130.00 Nulliparous

1.1.1.1 Statistics

Arithmetic means and standard deviations are presented as M +/- SD.

Calculations were done using SYSTAT software (Systat Software, Inc. 225 W

Washington St., Suite 425, Chicago, IL 60606). Inter-rater reliability for coding

behaviors was calculated using one hour of continuous footage that was independently

rated by each coder.

4.2.2.2 Behavioral Conditioning

Operant conditioning techniques were used to train the dolphins for urine and vaginal swab sampling. Behaviors were positively reinforced and slow approximations were taken until the dolphin was fully conditioned for daily sampling and exams.

4.2.2.3 Urine Collection

The dolphin was trained to urinate on cue by applying hand pressure to the bladder resulting in reflexive urination. The reflexive behavior was paired with primary reinforcement to increase the frequency, and over time the dolphin was conditioned to urinate by the application of gentle pressure to the bladder. For collection, the dolphin would beach laterally on a shallow ledge so that the genital slit was wiped clean and dry with gauze. A 50-ml sterile plastic specimen cup was placed under the urethra and gentle pressure was applied to the bladder. The urine was caught in the cup as it freely flowed from the urethra. An average of 5-10 ml of urine was collected from all study animals.

Urine was frozen at -17 oC in 5-ml increments immediately following collection. Urine for the study was collected in the AM (8 AM–11 AM) for the study unless noted.

4.2.2.4 Ultrasonography

Real-Time B-Mode trans-abdominal ultrasound imaging was utilized. For all animals, ultrasound exams were conducted as needed and exams typically lasted 3-5 min.

For ovary and fetus visualization, the dolphins were trained to float laterally and stationary at the water’s surface. The blowhole was submerged, but the animal could lift her head and take a breath as needed. For animal 1, a Sonosite 180 Plus ultrasound machine with a 5-2 MHz transducer (Sonosite Inc., 21919 30th Drive SE, Bothell, WA

98021-3904, USA) was used and images recorded using either the internal digital memory of the machine or by the use of an Archos 605 Audio/Visual Player (Archos

Inc., 7951 E.Maplewood Avenue #260 Greenwood Village, CO 80111 USA). For animals 6 and 7, a Sonosite Titan portable ultrasound machine with a 5-2 MHz curvilinear transducer (Sonosite Inc. 21919 30th Drive SE, Bothell, WA 98021-3904,

USA) was used. Images were digitally captured using the internal digital memory of the machine or by the use of an Archos 605 Audio/Visual Player (Archos Inc. 7951

E.Maplewood Avenue #260 Greenwood Village, CO 80111 USA).

4.2.2.5 Hormone Testing Progesterone, Estradiol and Lutenizing Hormone

Urinary progesterone (uP) and estradiol-17 beta (uE2) was measured using an electrochemiluminescent immunoassay (ECLIA) tested with the Elecsys 2010 instrument

(Roche Diagnostics, Mannheim, Germany) at Clinical Pathology Laboratory in Las

Vegas (4275 S. Burnham Ave, Suite 325, Las Vegas, NV 89119, USA) as previously reported (Muraco et al., 2009). Fresh collected chilled or frozen urine was transferred to the laboratory via courier. Urinary luteinizing hormone (uLH) was measured using the

ICG Canine Witness® Luteinizing Hormone Assay (Witness Synbiotics Corp., Kansas

City, MO, USA). The assay provides a rapid, semi-quantitative visible color band in the presence of dolphin LH (Muraco et al., 2009). Dolphin urine was applied to the test strip either directly following collection, or, if frozen, after thawing. A 100-µl volume of neat urine was placed onto the test strip and results were obtained in less than 1 hr after application. The LH surge was identified when the test band became darker than the control band.

4.2.2.6 Social Grouping

Animal 5, a mature proven sire, arrived to facility A on 4/9/06 and remained fully separated from animals 1-4 for quarantine purposes. On 4/26/06, the quarantine ended and dolphins 1-5 were able to see and hear one another for the first time. However, they remained separated by a nine meter canal and two closed see-through metal gates. On

5/16/06, animal 4 was introduced to animal 5, and animals 1-3 can see and hear animals 4 and 5 through closed see-through metal gates. Beginning 5/18/06, all the gates were opened, and all the animals (1-5) were able to physically interact with one another. On

6/8/06, animal 1 was separated from animals 2-5. At facility B, animal 6 was paired with animals 8 and 10, and animal 7 with animal 9 for the duration of the data collection period.

4.2.2.7 Behavioral Recording and Analysis

Animals 1-5 were monitored with continuous time stamped digital video recorded from an underwater viewing window using 3 Honeywell video surveillance cameras attached to a Honeywell digital recording device (Honeywell, Louisville, KY USA).

Daylight video footage was analyzed (7 AM–6 PM) when the animal resided in the pool with the cameras, and amount of time the animals resided in the study pool each day ranged from 0-6 hrs. Video was downloaded and stored on an external hard drive

(Maxtor OneTouch III 750GB Hard Drive, Seagate, Cupertino, CA USA).

Animals 6, 8, and 10 were videotaped from the surface for 1 hr each day using a

Canon HF200 HD camcorder (Canon USA Inc., Melville, NY) and footage downloaded onto an external hard drive (Seagate, Cupertino, California USA). Animals 7 and 9 were

videotaped opportunistically from the surface using an iPhone 4S (Apple Inc, Cupertino,

CA USA). All video footage was coded by the authors.

4.2.2.8 Proof of Ejaculation via the Identification of Spermatozoa in Vaginal Fluid

Vaginal fluid was collected daily from animal 1 to identify spermatozoa

indicating ejaculation had occurred. To collect vaginal fluid, the dolphin was partially

pulled out of the water so that the genital slit was dry. First, the trainer stood on a dry

ledge and grasped the tail flukes of the dolphin that was stationary in a dorsal position

floating at the pool surface. The dry ledge was 7 to 10 cm higher than the water surface.

Next, the trainer gently stepped backward pulling the dolphin’s flukes backward and

laterally over the ledge until the dolphin’s genital slit was laterally out of the water. The

dolphin’s head and pectoral flippers remained in the water. The genital slit was wiped

clean with gauze. The genital slit was gently spread apart using a gloved hand and a

sterile 15.24-cm cotton tipped swab with a plastic shaft (Puritan Medical Products,

Guilford, Maine, USA) was inserted 14 cm into the vaginal vault, twisted 360o, and

removed. The swab was immediately rolled onto a 75 mm x 25 mm glass microscope

slide and allowed to air dry for 24 hrs before reading. A phase contrast compound light

microscope (LW Scientific, Lawrenceville, GA USA) at 400x power was used to analyze

the slides. All slides were examined for the presence of spermatozoa by the author

Muraco using previously published methods (Muraco, Coombs, Procter, Turek, &

Muraco, 2012).

4.3 Results

4.3.1 Ultrasonography

Ultrasound exams were conducted by the author Muraco using previously

published methods (Muraco, Clough, Teets, Arn, & Muraco, 2010) to identify the pre-

ovulatory follicle (POF) and pregnancy in Animals 1, 6 and 7. For animal 1, a POF was

identified on 5/27/06 on the left ovary and the follicle was gone on 6/8/06 indicating

ovulation (Figs 1a and b). Uterine fluid and a fetus were identified on 8/15/06 (Figure

1c). For animal 6, a POF was identified on 1/11/13 and a fetus on 4/3/13 (Figure 2a and

2b). For animal 7, a POF was identified on 2/3/14 and ovulation occurred on

2/4/14(Figure 3a and 3b). Uterine fluid and an embryo were identified on 4/7/14(Figure

3c).

Figure 4.1 Animal 1 ultrasound a. white arrow points to pre-ovulatory follicle. b. white arrow points to ovulated follicle. c. white arrow points to uterine fluid and embryo 92

Figure 4.2 Animal 6 ultrasound a. white arrow points to pre-ovulatory follicle. b. white arrow points to fetus.

Figure 4.3 Animal 7 ultrasound a. white arrow points to pre-ovulatory follicle. b. white arrow points to corpus hemorrhagic

4.3.2 Endocrinology

Urine estradiol (uE2) and progesterone (uP) was tested in 28 samples surrounding

social change, estrus, ovulation and pregnancy in animal 1 (Figure 4). In animal 1, LH

was tested for 10 days, and the surge was identified in the am urine sample on 6/6/06.

Ovulation occurred 24 - 36 hrs later. Urinary progesterone was elevated (> 1.0 ng/ml) 14

days following ovulation.

Urinary LH was tested in animals 6 and 7 to identify the LH surge. For animal 6,

the LH surge was identified at 8 am on 1/13/13, and ovulation was assumed to have

occurred 24–36 hrs later based on previous studies (Muraco et al., 2010). For animal 7,

ovulation (11 am 2/4/14) occurred 27 hours after the LH surge was identified (9 am

2/3/14).

4.3.3 Reproductive behavioral ethogram

Video recordings from animals 1-5, 6, 8 and 10, and 7 and 9 were analyzed to

develop a reproductive behavioral ethogram. For animal 1, 22 days (49 total study hours)

of underwater behavioral footage was analyzed (2.227 +/- 1.602 hrs daily) starting the

day animals 1-4 were introduced to animal 5 and ending when animal 1 was separated

from the group following ovulation. For animal 6, 3 days (3 total study hours) of (above

the surface) behavioral footage was analyzed (1 hr daily) the day before the LH surge

through the day of ovulation while she was paired with a mature male and nulliparous

female (animals 8 and 10). Additionally, two minutes of underwater footage was taken of

animals 6, 8 and 10. Two days (4 min and 10 min) of opportunistic (above the surface)

behavioral footage and observations were conducted for animal 7 when she was paired

with a mature female (Animal 9) the day before and of the LH surge.

For the ethogram, 20 focal reproductive behaviors were identified from 52 total

hrs of recorded footage from animals 1-10 (Table 2). Focal behaviors were selected after

review of previously published dolphin ethograms and behavioral observations

(Dudzinski, 1998; Furuichi et al., 2014; Puente & Dewbury, 1976; Wells, 1984) and

estrus behaviors of domestic hoofstock (Bearden et al., 2004; Helmer & Britt, 1985).

Inter-rater reliability was 92.6%. 94

Figure 4.4 Animal 1 urinary estradiol, progesterone and luteinizing hormone

4.3.4 Reproductive State

For animal 1 (Table 3), behaviors observed following the introduction to the mature male(days -21 through -16) were considered anestrus due to low levels of uE2, basal levels of uP, and no follicular development. Behaviors observed during days -15 through -12 were considered socio-sexual based on an increase in sexual behaviors and increasing uE levels but without the presence of sperm on the vaginal fluid slides. Days -

11 through 0 are considered estrus based on elevated uE2 levels, copulatory behavior and the presence of sperm on the vaginal fluid slides. All sexual behaviors observed from animals 6 and 7 are considered estrus due to the close proximity to ovulation.

4.3.5 All-occurrences of reproductive behavior

One hour each day (12 total hours) was analyzed for all-occurrences of reproductive behavior by or towards the focal multiparous female animal 1 (Table 3), three total hours were analyzed for animals 6, 8 and 10, and 14 total minutes were analyzed for animals 7 and 9. Additionally, all-occurrences of atypical swimming patterns by the mature male, animal 5, were recorded.

Table 4.2 Ethogram of reproductive behavior

Behavior Description

1 Avoid (A) Female dolphin turns her genital slit away from a male dolphin (Fig 17) 2 Copulation (COP) Male dolphin thrusts penis into the genital slit of another dolphin (Fig 14) 3 Dorsal Mount (DM) One dolphin’s dorsal fin goes into the genitals of another dolphin (Fig 13) 4 Erection (E) Penis oriented towards another dolphin or object 5 Fluke mount (FM) One dolphin places a fluke into or against another’s genital slit 6 Flutter (FLU) Male dolphin performs an atypical swim pattern where the tail flukes are rapidly moved up and down in a flutter-like motion 7 Genital on head (GOH) Melon/head of a dolphin touches the genitals of another dolphin

8 Genital tracking (GT) One dolphin swims so that the rostrum is oriented towards the genitals of another without touching (Fig 11)

9 Goose (GOS) Rostrum touching a genital slit of another dolphin (Fig 12) 10 Group-on-one-sex (GPS) Two dolphins sexually thrust and rub on a single dolphin 11 Immobility (IM) Dolphin remains immobile in a lateral or upright position at the surface, in the water column or at the bottom of the pool for 3 or more seconds, often with eyes closed and mouth slightly open (Fig 6) 12 Line-up (LU) Three or more dolphins swim in a line with rostrums oriented towards the genital slit often with lead dolphin slightly raising up tail fluke (Fig 15) 13 Masturbate (MB) Single dolphin touches and rubs an object (i.e. toys) or structures in the environment (i.e. filtration grates) on genitals 14 Mount (MNT) One dolphin positions themselves onto the back of another dolphin 15 Pec mount (PMT) One dolphin places pectoral fin into or against another’s genital slit 16 Pursuit (PUR) Male dolphin quickly chases a female dolphin (Fig 16) 17 Push (P) One dolphin pushes another dolphin displaying immobility 18 Rubbing (R) Two or more dolphins rub one another with pectoral fins or body-to-body 19 Shark (SRK) Male dolphin performs an atypical swim pattern where the peduncle and flukes move left and right in a shark-like motion 20 Thrust (THR) An individual thrusts genitals towards another

1.1.1.1 Anestrus

Active behaviors by the focal female (animal 1) that were oriented towards the mature male (animal 5) following the introduction and during anestrus included avoid(3) and genital tracking (2), and no behaviors were observed by the multiparous female towards animals 2-4. In total, 5 behaviors were oriented towards the mature male during anestrus with avoid being most frequent.

4.3.5.2 Socio-Sexual

No behaviors were oriented towards the mature male or immature female (animal

3) during the socio-sexual period. Behaviors towards the immature male (animal 4) included genital tracking (1), group-on-one sex (1), dorsal mount (3), push (1), mount (1), thrust (1), rub(1), and fluke mount(1). Behaviors towards the nulliparous mature female

(animal 4) included group-on-one sex (1). In total, 10 behaviors were oriented towards the immature male and 1 towards the nulliparous female. Dorsal mount was the most

frequent.

4.3.5.3 Estrus

No behaviors were oriented towards the immature female during estrus.

Behaviors towards the mature male included avoid (2), genital tracking (1), goose (1),

and copulation(2). Behaviors towards the nulliparous mature female included dorsal

mount (12), genital tracking (1), thrust (5), rub (5) and goose (3). Behaviors towards the

immature male included dorsal mount (1). In total, 6 behaviors were oriented towards the

mature male, 26 towards the nulliparous female and 1 towards the immature male. Dorsal

mount was the most frequent.

4.3.6 Passive behaviors towards animal 1 (Figure5)

4.3.6.1 Anestrus

No behaviors were oriented towards the focal female by animals 2-4 during anestrus. Behaviors by the mature male towards the focal animal following the introduction and during anestrus included pursuit (3) and genital tracking (2). In total, 5 behaviors were oriented towards the focal animal by the mature male with pursuit being the most frequent.

4.3.6.2 Socio-sexual

No behaviors were oriented towards the focal female by the mature male, the immature female or the nulliparous female during the socio-sexual period. Behaviors by the immature male towards the focal female during the socio-sexual period included genital tracking (2). In total, 2 genital tracking behaviors were orientated towards the focal female by the immature male.

4.3.6.3 Estrus

Behaviors by the mature male towards the focal female during estrus included pursuit(2), genital tracking(6), goose(1), push(1), mount(1), genital-on-head(1), erection

(2), line-up(1) and copulation(2). Behaviors by the immature male towards the focal female included genital tracking (3) and goose(1). Behaviors by the nulliparous female towards the focal female included genital tracking(5), goose(13), dorsal mount(34), push(1), mount(2), thrust(2), rub(4), fluke mount(1), pec mount(1), and genital-on- head(1). No behaviors were oriented towards the focal female by the immature female. In

total, 17 behaviors were oriented towards the focal female by the mature male, 4 by the immature male, and 64 by the nulliparous female. Dorsal mount was the most frequent.

Figure 4.5 Percentage of reproductive behaviors during each reproductive phase for animal 1

. a. active. b. passive

4.3.7 Behaviors enacted by and towards the focal nulliparous females (animals 6 and 7) during estrus

4.3.7.1 Active reproductive behaviors by animals 6 and 7

Behaviors enacted by the focal nulliparous female (animal 6) towards the mature

male (animal 8) included avoid (41) and copulation (6). Behaviors by animal 6 towards

the nulliparous female (animal 10) included rub (1) and towards toys included masturbate

(6). In total, 47 behaviors were orientated towards the mature male by the focal female

and 1 towards the nulliparous female.

Behaviors enacted by the focal nulliparous female (animal 7) towards the

multiparous pregnant female (animal 9) included genital tracking (1), genital-on-head (1)

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and goose (1). In total, 3 behaviors were orientated towards the pregnant female by the focal nulliparous female.

4.3.7.2 Passive reproductive behaviors by animals 6 and 7

Behaviors by the mature male towards the focal nulliparous female (animal 6) included pursuit(48), push(16), genital tracking(2), copulation(6), mount(6), goose(1), line-up(1), and erection(1). In total, 81 behaviors were orientated towards the focal nulliparous female by the mature male and 0 by the nulliparous female. Avoid and pursuit was the most frequent.

4.3.8 Behaviors from animal 9 towards animal 7 included push (1)

4.3.8.1 Immobility

All three focal females (animals 1, 6 and 7) demonstrated bouts of immobility

(IM) during which they became motionless in the water, often with eyes closed and mouth open (Figure 6). This behavior occurred when the animal was alone (i.e., not in close proximity of any other dolphins) as well as when interacting with other dolphins.

The behavior was observed to occur throughout the water column from the surface to the bottom of the pool and in both vertical and horizontal positions.

Animal 1 demonstrated immobility 31 times between 5/28/06 and 6/8/06 (6 hrs) totaling 397 s (Figure 7), with the longest continuous immobility bout being 34 s. Animal

6 demonstrated immobility 20 times between 1/12/13 and 1/14/13 (3 hours) totaling 247 s with the longest continuous immobility bout being 34 s. Animal 7 demonstrated immobility 12 times between 2/2/14 and 2/3/14 (14 min) totaling 502 s with the longest continuous immobility bout being 150 s.

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Figure 4.6 Immobility (IM) behavior

Lateral position with eyes closed and mouth slightly open

Figure 4.7 Animal 1 immobility bouts, total duration in seconds and maximum bout duration in seconds

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4.3.9 Behavioral Correlations to Endocrinology

4.3.9.1 Estradiol

Urinary E2 levels and the combined total number of reproductive behaviors by and towards the focal female (animal 1) during each reproductive state were compared

(Table 4). A trend was observed showing an increase in the number of sexual behaviors as mean uE2 levels rose (Figure 8).

Table 4.3 Animal 1 number of reproductive behavioral occurrences and uE2 levels

Mean uE2 Max uE2 Repro State Beh Oca pg/ml pg/ml

Anestrus 10 66.00 +/-2.517b 69.00 Socio-Sexual 13 73.00 +/- 1.414c 75.00 Estrus 118 75.667 +/- 2.521d 85.00 a. combined active and passive behavioral occurrences. b. n 3. c. n 4. d. n 12

4.3.9.2 Luteinizing hormone (LH)

Total bouts of the immobility behavior (IM) reached maximum numbers during

the LH surge in all three focal females (animals 1, 6 and 7) (Figure 8 and 9).

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Figure 4.8 Animal 1 immobility, uE2, uLH and regression

Figure 4.9 Animals 6 and 7 immobility duration and uLH correlation

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4.3.10 Copulation observations and spermatozoa in vaginal fluid

Only two copulations (5/31/06, 6/8/06) were observed in 12 hours of behavioral

footage between animals 1 and 5, however vaginal fluid collected for 18 consecutive days

(5/22/06–6/8/06) showed spermatozoa on 7 days between 5/30/06 and 6/7/06 (Table 3).

The copulation on 5/31/06 started with the male genital tracking the female, the female

showing immobility, the male goosing the female, followed by copulation occurring with

the animals’ perpendicular to one another near the water’s surface. The copulation on

6/8/06 started with the female (animal 1) immobile and the nulliparous female (animal 2)

pushing, dorsal mounting, thrusting and mounting animal 1. The male (animal 5) was

nearby and swam in and copulate with animal 1 who was immobile.

Surface behavioral recordings revealed six copulations between animal 6 and 8.

Each successful copulation was preceded by a similar pattern in which the female was in

a lateral surface immobility position, the male approached and genital tracked, goosed or

pushed, followed by copulation near the surface with the animals perpendicular to one

another (Figure 14). During unsuccessful copulation attempts the male would pursue a

female that was not demonstrating the immobility behavior, and she would avoid his

pursuits by turning her genital slit away from him or swimming away.

4.3.11 Mature male (animal 5) display

The mature male (animal 5) demonstrated two distinct types of atypical

swimming patterns, shark and flutter which appeared to be used as a form of courtship

display. The shark swim pattern included the male moving the peduncle and tail flukes in

a side-to-side motion. The flutter involved moving the tail flukes up and in down rapidly

without a proportional forward momentum. 105

On 5/19/06, the day after the introduction of the focal female (animal 1) to the mature male (animal 5), the male swam under the focal female and genital tracked.

Immediately after the genital tracking, the male swam in a shark-like motion in front of the focal female (animal 1).

On 5/23/06, two atypical swim patterns were observed. The first involved the immature male (animal 4). The immature male swam around the mature male (animal 5) who was positioned vertically in the water column. The mature male performed the shark swim pattern while vertical in the water column. The second atypical swim pattern involved the focal female. The mature male genital tracked the focal female, and the focal female responded by avoiding the male; the male then positioned himself horizontally near the water surface and swam in a flutter-like motion.

On 5/31/06, two atypical swim patterns were observed. The first occurred when the male swam under and in front of the focal female and swam in the shark swim pattern. The second occurred when the mature male genital tracked the focal female followed by the focal female showing immobility. The male swam in a flutter-like motion near the water’s surface, and seven seconds later copulation occurred.

On 6/1/06, one atypical swim pattern was observed. The mature male genital tracked the focal female, and the female avoided the male. The male then positioned himself near the surface of the water and swam in a flutter-like motion.

4.4 Discussion

Our intent was to define reproductive behaviors in dolphins of known

reproductive states and identify behaviors that could be exclusive to estrus. Dolphins

evolved from artiodactyls (even-toed ungulates including cows, pigs hippos and camels) 106

(Milinkovitch & Thewissen, 1997), and because of the monophyletic grouping of cetaceans and artiodactyls, a dolphin is more closely related to a cow than a horse is.

Perhaps some terrestrial reproductive processes have been conserved in the dolphin that we can use to improve our understanding of this species. For the purposes of this study, the estrus period was defined as the period between the female first exhibiting immobility until ovulation, and this is similar to what is accepted as the criteria for the estrus period in cattle (Helmer & Britt, 1985). Estrus duration and the timing of ovulation after the start of estrus is species dependent, varies from one female to another, and differs with nulliparous or parous animals. In cows, estrus lasts from ~18-19 hrs with ovulation occurring 10-11 hrs after estrus end; in the sow, estrus lasts 48-72 hrs with ovulation occurring 35-45 hrs from the beginning of estrus; and in the mare, estrus lasts 4-8 days with ovulation occurring 1-2 days before estrus end (Hafez & Hafez, 2000). The average follicular and luteal phase for a dolphin lasts 10 and 20 days respectively (Muraco et al.,

2010). There was an increase in reproductive behavioral occurrences during estrus as estradiol and luteinizing hormone levels peaked, which is consistent with patterns in domestic species (Austin & Short, 1972; Bearden et al., 2004; Hunter, 1980; Pineda &

Dooley, 2003). In the dominant, multiparous dolphin (animal 1), estrus spanned the duration of the 10 day follicular phase and peak behaviors occurred during the LH surge.

She primarily sexually interacted with the nulliparous mature female during her estrus period with active and passive dorsal mounting seen most frequently. In the nulliparous, sub-dominant female (animal 6), copulation was only successful during LH and ovulation periods when the female displayed immobility. When not engaged in sexual interactions with the mature, dominant male (animal 8), she was observed masturbating with toys.

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Two unique behaviors observed in the study were genital tracking and immobility. The genital tracking behavior was first observed upon introduction of the female 1 to the mature male 5 with the behavior increasing in frequency during estrus.

The genital tracking behavior is similar in appearance to “genital sniffing” behavior observed in terrestrial mammals, and it is possible that dolphins may utilize

chemosensory abilities for reproduction. Studies conducted by the United States Navy

showed that dolphins have the ability to distinguish salty, sweet, sour and bitter, with

most sensitivity to bitter (Nachtigall, 1986; Nachtigall & Hall, 1984), and Russian

scientists hypothesized that dolphins may use taste reception in a form of chemically-

mediated communication (Kuznetsov, 1990; Yablokov, 1961). Perhaps dolphins are able

to gain information about another dolphin’s physiological state via a form of chemo-

communication. Although it is possible that the dolphins were looking at the genital slit

to gain a visual cue, there was no obvious change, such as swelling, observed at any point

in the study. The genital slit occasionally turned a light shade of pink following bouts of

rubbing, but since dolphins are colorblind (Peichl, Behrmann, & Kröger, 2001), this

would not work well as a visual cue. There was no evidence in this study that the

dolphins were echolocating on the genital slit when engaged in genital tracking, although

no audio recordings were formally analyzed. The genital tracking behavior often

progressed to the goose behavior, which is similar in appearance to “vulva nuzzling”

observed in terrestrial mammals. The goose behavior has been reported as part of dolphin

socio-sexual repertoires (Dudzinski, 1998), and it has been observed in adult females

towards their offspring (Figure 10) (Ridgway, 1972).

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Figure 4.10 A dolphin mother gooses her 3 month old female calf.

The immobility behavior was observed in all three focal females during estrus, and maximum immobility duration occurred during the LH surge. The estrous cycle is regulated by endocrine and neuroendocrine mechanisms including hypothalamic hormones, gonadotropins and gonadal steroids (Bearden et al., 2004; Hafez & Hafez,

2000). Luteinizing hormone-releasing hormone (LHRH) is primarily responsible for the ovulatory surge of LH and facilitation of lordosis (estrus) in rats (Foreman & Moss,

1979; Moss & McCann, 1975; Moss & Foreman, 1976; Pfaff, 1973), and in sows, maximum LH levels being present at the first observance of behavioral estrus (Tilton,

Foxcroft, Ziecik, Coombs, & Williams, 1982). Bottlenose dolphins have pre-ovulatory

LH surges that last 12-24 hours with peak levels lasting 6 hrs, and ovulation following peak LH levels within 24-48 hrs (Muraco et al., 2010). In this study, immobility occurred prior to or during each observed copulation which suggests that dolphins demonstrate a novel form of lordosis and/or standing heat estrus that assists in copulation success. The

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genital tracking and goose behaviors may serve to communicate reproductive readiness

(or lack thereof), while immobility demonstrates copulatory receptivity.

The dominant female (animal 1) demonstrated a period of active socio-sexual behavior with the immature male following anestrus and prior to estrus. Her estradiol levels were above anestrus levels during this time period which may have contributed to the behavior. However, she was observed aggressively herding the immature male away from the mature male, so the socio-sexual behavior may have been a display of dominance. Unlike the gonadal steroid estrogen, and the protein hormone LH, which contribute to conceptive estrus behavior (Foreman & Moss, 1979; Moss & McCann,

1975; Moss & Foreman, 1976; Pfaff, 1973), socio-sexual behavior can be mediated by oxytocin (Carter, 1992; Kendrick et al., 2000). Oxytocin is a nonapeptide produced primarily in the supraoptic (SON) and paraventricular (PVN) nuclei of the hypothalamus

(Carter, 1992). Pulses of oxytocin are released into systemic circulation from the posterior pituitary following tactile stimulation in rats and during sexual arousal in humans (Carter, 1992). It is probable that oxytocin may be responsible for the highly tactile socio-sexual interactions of dolphins, while estrogen and LH contribute to estrus behaviors.

Finally, the proven sire (animal 5) demonstrated atypical swim patterns that appeared to act as a form of display or courtship. The male (animal 5) was new to the social group where the focal female (animal 1) was dominant. He was observed displaying the unusual swim patterns in several contexts: (1) following the introduction to the focal female, (2) after the focal female demonstrated avoidance behavior, and (3) once prior to copulation.

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Encountering sexual partners though mutual searching, identification of physiological states and sequential interactions that result in copulation are essential steps to ensure the survival of any species, and we have a fairly clear understanding of terrestrial animal reproductive biology, behavior and ecology. However, the holaquatic world of the dolphin remains steeped in mystery, and our knowledge of reproduction in

Cetacea is rudimentary at best. Future studies should focus on sex hormones, oxytocin levels and behavior during non-conceptive socio-sexual scenarios, conceptive estrus cycles, and male courtship displays to help determine the relative roles of these variables on dolphin reproductive behavior.

Figure 4.11 Genital tracking (GT) of animal 1 by animal 4

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Figure 4.12 Goose (GOS) of animal 1 by animal 4

Figure 4.13 Dorsal Mount (DM) of animal 1 by animal 2

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Figure 4.14 Copulation (COP) between animal 8 and animal 6

Figure 4.15 Line Up (LU) animal 1 at the front with animals 4 and 5 behind

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Figure 4.16 Mount (MNT) by animal 8 to animal 6.

Figure 4.17 Avoid (A) by animal 6 of animal 8.

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CHAPTER V

CHANGES IN A BOTTLENOSE DOLPHIN (TURSIOPS TRUNCATUS) VAGINAL

FLUID ARBORIFORM ARRANGEMENT DURING ANESTRUS AND THE

ESTROUS CYCLE

5.1 Introduction

In the human, vaginal fluid is composed of several components including: glandular vulvar secretion, vaginal wall transudate, exfoliated cells, cervical mucus and endometrial and oviductal fluids (Huggins & Preti, 1981). The viscoelastic cervical mucus component of vaginal fluid is produced by secretory epithelial cells of the cervix which secrete glycoproteins made up of amino acids and carbohydrates (Menarguez,

Pastor & Odeblad, 2003). There are endocrinological cyclic variations in the arrangement and viscosity of the cervical mucus which will help or impede the penetrability of spermatozoa through the cervical canal and provide a barrier to prevent the pathogens entering the endometrium (Menarguez, Pastor & Odeblad, 2003; Brunelli et al., 2007).

During estrus and ovulation, estrogenic cervical mucus forms an enlarged meshwork so that sperm has adequate space to progress into the uterus, and in contrast, during the gestagenic luteal phase, the mesh becomes increasingly smaller preventing sperm transport (Bearden, Fuquay, & Willard, 2004; Hafez & Hafez, 2000). When cervical mucus is allowed to dry on a microscope slide, a crystallization of the mucus forms which creates an arboriform or ferning pattern (Menarguez, Pastor & Odeblad, 2003;

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England & Allan, 1989). Studies in humans, cows, and dogs show cervical mucus and/or vaginal fluid will show different types of crystallization, arboriform arrangement, and ultrastructure depending on the cycle phase (England and Allan, 1989; Menarguez, Pastor

& Odeblad, 2003; Noonan, Schultze & Ellington, 1975). Four primary types of cervical mucus secretions have been described in humans including: G (gestagenic), L (loaf), S

(string) and P (peak) mucus, each with a different viscosity and morphology in the crystallization (Menarguez, Pastor & Odeblad, 2003).

The bottlenose dolphin female reproductive anatomy consists of an external mid- ventral genital slit which houses the internal urethral orifice, the clitoris and vulva.

Mature dolphins have a 15-20 cm vagina with thick muscular walls and extensive deep longitudinal rugae. At the vaginal fornix are numerous valve-like vaginal folds and a pseudocervix just caudal to the true cervix forming a spermathecal recess (Harrison,

1969; Ridgway, 1972). The dolphin true cervix ranges from 3.8-5 cm in length with a smooth canal, and delineates the uterine body from the vagina. The short uterine body joins two horns forming a bicornuate uterus with compressed fallopian tubes on the anterior ends of the horns. The ovaries are located posterior to the kidneys and are held in place by a broad ligament and each ovary is covered by a large fallopian funnel

(Ridgway, 1972).

Nothing was found in the literature about dolphin vaginal fluid or cervical mucus,

although mucus, presumably originating from the cervix, has been noted flowing from

the genital slit in dolphins.

This study looked at the arboriform arrangement of vaginal fluid during and

anestrus, estrus, ovation and early luteal phase of a conceptive cycle, as well

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documenting the gross appearance of the estrogenic vagina and cervix through the use of vaginal endoscopy.

5.2 Method

5.2.1 Experimental ethics

All animals, materials and methods used in the study were individually evaluated

by the facility’s Internal Animal Care and Use Committee (IACUC) to ensure safety,

health and well-being of the animals. Samples were taken during routine behaviors in

which the animals were conditioned to participate.

5.2.2 Study animals

Animal 1 (Table 1) was housed at The Mirage Hotel (Mirage; Las Vegas, Nevada

USA), in a 3,141,892-L manufactured seawater pool, maintained at 21 to 23° C year-

round. The animal diet consisted of frozen thawed herring (Clupea harengus), capelin

(Mallotus villosus), sardine (Sardina pilchardus), and squid (L. opalescens) for daily

totals of 10 kg. Animal 2 (Table 1) had a vaginal endoscopy performed for the purposes

of artificial insemination and was housed at Six Flags Discovery Kingdom (SFDK;

Vallejo, California USA) in a 2717.93-L manufactured seawater pool maintained at 21-

22o C year round. Animal 2 ate a diet of whole frozen thawed herring (Clupea harengus)

and capelin (Mallotus villosus) with totals of 8 kg per daily.

Table 5.1 Animals in study

Weight Reproductive Animal Birth Date (kg) History 1 11/1975 191.86 Parous 2 5/1986 189.00 Parous

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1.1.1.1 Behavioral Conditioning

Operant conditioning techniques were used to train the dolphins for urine, vaginal swab and vaginal endoscopy. Behaviors were positively reinforced and slow approximations were taken until the dolphin was fully conditioned for daily sampling and exams.

5.2.2.2 Vaginal Endoscopy

Animal 2 was trained to allow a flexible video endoscope (Olympus America,

Center Valley, PA 18034 USA) enter approximately 15 cm into the vaginal vault and images were recorded with an Archos digital recording device (Archos, Irvine, CA 92618

USA).

5.2.2.3 Vaginal fluid collection and analysis

The dolphin was partially pulled out of the water so that the genital slit was dry.

First, the trainer stood on a dry ledge and grasped the tail flukes of the dolphin that was stationary in a dorsal position floating at the pool surface. The dry ledge was 7 to 10 cm higher than the water surface. Next, the trainer gently stepped backward pulling the dolphin’s flukes backward and laterally over the ledge until the dolphin’s genital slit was laterally out of the water. The dolphin’s head and pectoral flippers remained in the water.

The genital slit was wiped clean with gauze. The genital slit was gently spread apart using a gloved hand and a sterile 15.24 cm cotton tipped swab with a plastic shaft

(Puritan Medical Products, Guilford, Maine, USA) was inserted 14 cm into the vaginal vault, twisted 360o, and removed. The swab was immediately rolled onto a 75 mm x 25

mm glass microscope slide and allowed to air dry for 24 hours before reading.

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A phase contrast compound light microscope (LW Scientific, Lawrenceville, GA

USA) at 400x power was used to analyze the slides. All slides were examined for ferning and for the presence of spermatozoa. Dolphin ferning morphology was identified on each slide according to Menarguez, Pastor & Odeblad (2003) as types G (gestagenic), L (loaf),

S (string), P (peak) and P4 (peak subtype 4). A series of digital photos (Am Scope, Irvine,

CA USA) was taken of each type of mucus morphology that appeared on each slide.

5.2.2.4 Hormone Testing Progesterone, Estradiol and LH

Urinary progesterone (P) and estradiol-17 beta (E2) was measured using an

Electrochemiluminescent Immunoassay (ECLIA) tested with the Elecsys 2010 instrument

(Roche Diagnostics, Mannheim, Germany) at Clinical Pathology Laboratory in Las

ICG Canine Witness® Luteinizing Hormone Assay (Witness Synbiotics Corp., Kansas

City, MO, USA). The assay provides a rapid, semi-quantitative visible color band in the presence of dolphin LH (Muraco et al., 2009). Dolphin urine was applied to the test strip either directly following collection, or, if frozen, after thawing. A 100-µl volume of neat urine was placed onto the test strip and results were obtained in less than 1 h after application. The LH surge was identified when the test band became darker than the control band.

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5.2.2.5 Ultrasonography

Real-Time B-Mode trans-abdominal ultrasound imaging was utilized. The dolphins were trained to float laterally and stationary at the water’s surface. The blowhole was submerged, but the animal could lift her head and take a breath as needed. To visualize the ovaries and POF, the ultrasound transducer was placed between the junction of the rectus abdominus muscle and the hypaxialis lumborum muscle (Brook, 2001). For animal 1, a Sonosite 180 Plus ultrasound machine with a 5-2 MHz transducer (Sonosite

Inc., 21919 30th Drive SE, Bothell, WA 98021-3904, USA) was used and images recorded using either the internal digital memory of the machine or by the use of an

Archos 605 Audio/Visual Player (Archos Inc., 7951 E.Maplewood Avenue #260

Greenwood Village, CO 80111 USA). Ultrasound exams typically lasted 3-5 minutes for both the left and right lateral positions. For animal 2, a Sonosite Titan portable ultrasound machine with a 5-2 MHz curvilinear transducer (Sonosite Inc. 21919 30th Drive SE,

Bothell, WA 98021-3904, USA) was used. Images were digitally captured using the internal digital memory of the machine or by the use of an Archos 605 Audio/Visual

Player (Archos Inc. 7951 E.Maplewood Avenue #260 Greenwood Village, CO 80111

USA).

5.3 Results

5.3.1 Vaginal fluid arboriform (ferning) arrangement

61 air dried vaginal fluid slides were analyzed from May 1, 2006–June 30, 2006

encompassing a period of anestrus and subsequent estrus and luteal phases of a

conceptive ovulatory cycle in animal 1 for the presence of ferning patterns. All 4 mucus

ferning patterns that have been described in humans by Menarguez, Pastor & Odeblad 120

(2003) (Table 2) were identified in the dolphin along with 1 distinct subtype (Figures 1-

5). Ferning patterns showed a distinct difference depending on cycle phase (Table 3 and

Figure 6). Additionally slides were scanned for the presence of sperm which indicated

breeding activity (Figure 7).

Table 5.2 Cervical mucus types

Mucus Type Description

Gestagenic (G) A free crystal content with no predetermined form Loaf (L) Typical ferning morphology with straight or curved central axis and branches protruding from it at a 90o angle String (S) A parallel arrangement of crystals close to one another but not joined Peak (P) A star-like morphology with or without well-defined axes Subtype Peak 4 (P4) 2 fundamental and well-defined axes with a 90o angle between them and branches projecting from the main axis at a 60o angle

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Figure 5.1 Dolphin vaginal fluid type G

Figure 5.2 Dolphin vaginal fluid type L

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Figure 5.3 Dolphin vaginal fluid type S

Figure 5.4 Dolphin vaginal fluid type P

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Figure 5.5 Dolphin vaginal fluid type P4

Table 5.3 Number of dolphin vaginal fluid arboriform types identified during anestrus, follicular and luteal phases

Luteal phase Mucus Anestrus Follicular (Days +1– Type (22 days) (Days -16–0) +22)

G 13 11 20 L 12 9 8 S 4 15 3 P 4 13 4 P4 0 3 2

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Figure 5.6 Vaginal fluid arboriform types and spermatozoa during anestrus, follicular and luteal phases

X = sperm

Figure 5.7 Spermatozoa on Vaginal Fluid Slide With Type P Mucus.

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5.3.2 Endocrinology

57 urine estradiol and progesterone samples were tested during anestrus, estrus, ovulation and early luteal phase in animal 1 (Figure 8). Ovulation occurred on June 8. uP was elevated (>1.0 ng/ml) 14 days following ovulation.

Animal 2 had a positive LH on 2/23, and a vaginal endoscopy was performed for the purposes of artificial insemination.

Figure 5.8 uE2 and uP levels of animal 1

5.3.3 Ultrasonography

Ultrasound exams were conducted to identify the POF and ovulation in animals 1 and 2.

For animal 1, a POF was identified on May 27 on the left ovary and the follicle

was gone on June 8 indicating ovulation (Figures 9 a & b).

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For animal 2, a POF was imaged on 2/23 and ovulation occurred on 2/24 (Figures

10 a & b).

Figure 5.9 Animal 1 ultrasound

. a. POF. b. ovulation.

Figure 5.10 Animal 2 ultrasound a. POF. b. ovulation

5.3.4 Vaginal Endoscopy

Vaginal endoscopy was performed for the purposes of artificial insemination, and opportunistic photos were taken of the internal reproductive anatomy (Figures 11-14).

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Images were taken of the vaginal walls (Figure 11), pseudocervix (Figure 12), spermathecal recess (Figure 13) and external cervical os of the true cervix (Figure 14) of an LH positive dolphin (animal 2) on 2/23. Vaginal fluid with viscoelastic mucus can be

visualized covering the tissues (Figures 11-14).

Figure 5.11 Vaginal walls

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Figure 5.12 Pseudo-cervix/vaginal flap

Figure 5.13 Spermathecal Recess

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Figure 5.14 True cervix

5.4 Discussion

This study is the first to report on an arboriform arrangement in bottlenose

dolphin vaginal fluid. Although there was only one animal studied, it showed the

potential application for identifying the state of fertility. As seen in humans, mucus type

“S”, with a parallel arrangement and large open mesh, was prevalent during the

estrogenic follicular phase, which would allow sperm to penetrate through the cervical

canal (Menarguez, Pastor & Odeblad, 2003; Hafez & Hafez, 2000). During anestrus and

the luteal phases gestagenic mucus type “G” , with no distinct crystallization and closed

mesh, was the primary form observed which would make it difficult for sperm to

penetrate (Menarguez, Pastor & Odeblad, 2003; Hafez & Hafez, 2000). Mucus type “L”

was seen more often during anestrus and luteal phases, and type “P” and sub-type “P4”

was present primarily during the follicular phase. Future studies will encompass more 130

animals during both conceptive and non-conceptive cycles and calculate the percentage of each mucus type present on each microscope slide.

In human and bovine cervical mucus studies, mucus was collected directly from the external os to see the maximal crystallization potential (Cortes et. al., 2012;

Menarguez, Pastor & Odeblad, 2003). In a study of anterior vaginal fluid in dogs, arborization patterns were identified that were positively correlated with proestrus and estrus, but there were some variations in the crystallization results between the study animals (England and Allan, 1989). England and Allan (1989) suggested that the result variations in the dog could be due to the use of vaginal fluid instead of cervical mucus. In the dolphin, the true cervix is covered by the pseudocervix and surrounded by multiple vault-like folds. The anatomy is such that the only way to collect cervical mucus from the os would be through endoscopy. That isn’t a realistic option for serial studies. The dolphin vaginal fluid in this study was collected at a depth of 14 cm into the 15-20 cm long vagina, which would put the collection device close to the vaginal fold arrangements just caudle to the true cervix. Gross anatomical examination via vaginal endoscopy of an

LH positive female dolphin showed ample mucus on the vaginal folds and pseudocervix

(Figures 11-14).

This study shows the future potential of utilizing a non-invasive collection of vaginal fluid for the purposes of identifying fertile periods of the dolphin. This process could be utilized by field researchers during catch and release studies to give further information about the reproductive phase of the animals.

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CHAPTER VI

C18 REVERSED-PHASE HIGH-PERFORMANCE LIQUID CHROMATOGRAPHY

(RP-HPLC) OF BOTTLENOSE DOLPHIN (TURSIOPS TRUNCATUS) URINE TO

IDENTIFY POTENTIAL CANDIDATE PROTEIN AND/OR PEPTIDE

PHEROMONES

6.1 Introduction

Dolphins are social mammals that utilize a variety of sensory abilities to survive

in their holaquatic world. Vocalizations, hearing, echolocation, visual acuity and tactile

sense are well-studied sensory mechanisms utilized by dolphins (Harrison & Bryden,

1986; Ridgway & Harrison, 1999). However, chemoreception (taste, olfaction,

pheromones) has had very little study, and it is unknown if, how, or to what extent

dolphins may utilize these senses in their daily life. It is also unknown how dolphins

signal reproductive condition to one another (Ridgway, Leatherwood & Reeves, 1990;

Connor, Smolker & Richards, 1992).

Chemoreception plays an important reproductive role in almost all species

(Gorman et al., 1974, 1978; Rasmussen and Krishnamurthy, 2000; Rekwot at el., 2001),

and pheromones have been identified from species in every animal phylum (Wyatt,

2010). It is likely that the majority of species across the animal kingdom use pheromones

for communication (Wyatt, 2010).

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Chemical substances that convey information among individuals of the same species can be defined as pheromones (Brennan & Keverne, 2004; Baxi, Dorries &

Eisthen, 2006). Originally the term pheromone was created based on the identification of a volatile sex attractant, bombykol, which was released by the female silk moth that elicits the sexual behavior of male moths. However, recent definitions for pheromones don’t contain the words ‘odor’ or ‘odorant’ because the chemical messenger does not have to be odorous or volatile to transfer signals between conspecifics (Touhara, 2008), and any bodily secretion can be a potential route for pheromonal communication

(Brennan & Keverne, 2004).

In terrestrial mammals, pheromones are typically processed by two systems; the vomeronasal system, which mediates responses to molecules of low volatility, and the olfactory system, which mediates responses to volatile molecules (Baxi, Dorries &

Eisthen, 2006). Many pheromonal responses are mediated by the accessory olfactory organs rather than main olfaction (Singer, 1991; Dulac & Torello, 2003), and some pheromones can be detected by the taste system, a possibility that has not yet been explored in vertebrates (Baxi, Dorries & Eisthen, 2006).

Pheromones are used by most species, from single-cell organisms to mammals, and some components of pheromone signaling systems have been conserved through evolution from insects to mammals (Dulac & Torello, 2003). Pheromones can elicit both short and long lasting effects (Singer, 1991; Dulac & Torello, 2003). Long lasting effects alter the endocrine state of the recipient via the hypothalamic pituitary (HP) axis with lutenizing hormone (LH) and prolactin (Prl) playing key roles (Dulac & Torello, 2003).

For example, in female mice, the physiological response to male pheromones can cause

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an increase in LH and reduction of Prl (Dulac & Torello, 2003). Additionally, pheromones can elicit rapid behavioral responses such as regulation of inter-male aggression, initiation of male and female copulatory behavior and parent-infant interactions; these immediate behavioral responses are also modulated by the endocrine status of the animal (Dulac & Torello, 2003).

Pheromones can be conveyed in aqueous solution via proteins and peptides. In contrast to the well-known volatile pheromones, which are produced metabolically, relatively large non-volatile compounds, such as proteins and peptides, can also be pheromones (Singer, 1991; Dulac & Torello, 2003; Touhara, 2008). Touhara (2008) reports that that secreted peptides or proteins can possibly function not only as hormones in the body but also as chemosensory pheromone signals that convey sociosexual information between individuals. Unlike volatile pheromones, peptides and proteins are genetically coded, stable and not airborne, so the information they convey, such as species, sex and identity of the individual, remains constant throughout the lifespan of the animal. Peptide and protein pheromones have been identified in vertebrates (amphibians, rodents), worms, insects, mollusks and in prokaryotic and eukaryotic species (bacteria, fungi); it is likely additional novel pheromones in other species will be discovered

(Touhara, 2008).

Many aquatic animals utilize protein and peptide chemosignals, and water is an ideal medium to disseminate aqueous chemosensory cues over large areas (Dulac &

Torello, 2003). In the aquatic environment, solubility is important, so high molecular weight peptides and proteins have an attractant role (Brennan & Zufall, 2006). Chemical signals are especially important in aquatic environments where visual and auditory

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stimuli may be limited, and aquatic cues are easily transmitted with a high signal-to-noise ratio (Belanger & Corkum, 2009). A decapeptide, sodefrin, was isolated from the cloacal abdominal gland of the male red-bellied newt (Cynops pyrrhogaster), and both the native peptide and a synthetic one had female-attracting activity (Kikuyama et al., 1995;

Brennan & Zufall, 2006). In the lamprey, a mixture of sulphated steroids and bile acids are utilized as migratory pheromones (Brennan & Zufall, 2006). In anurans (frogs and toads), both larvae and adults respond to chemical signals such as recognizing predators, kin, and injured conspecifics (Belanger & Corkum, 2009). Peptide pheromones or chemoreceptive cues have been identified in ragworms, polychaetes, amphibians,

Aplysia, settlement cues in barnacles, and ciliates and marine bacteria (Painter et al.,

1998; Hardege, 1999). Aquatic animals use small molecules for long-distance communication such as the sex pheromones in the lamprey, and large soluble peptide pheromones for close range communication such as those found in the sea-slug mollusk and nereid worms (Wyatt, 2010). Goldfish steroid hormone products have been identified as pre-ovulatory sex pheromones and are released in various combinations depending on reproductive status (Dulac & Torello, 2003).

Terrestrial mammalian pheromonal communication via aqueous solutions is well known and plentiful in the literature. Urine and other aqueous bodily secretions are rich in semiochemical cues (Dulac & Torello, 2003). An elephant pheromone, (Z)-7-dodecen-

1-yl acetate, is found in female pre-ovulatory urine, is active in water and causes male

elephant sexual behavior (Dulac & Torello, 2003). Chemical communication in the pig is

mediated by saliva, and steroid hormones of mature males have been shown to act as

pheromones that initiate cycling in the female (Marchese et al., 1998). The Bruce effect,

135

when a pregnant female undergoes spontaneous abortion when exposed to an unfamiliar male, was initiated in a pregnant female mouse when a MHC peptide from an unfamiliar male was added to a familiar male’s urine (Touhara, 2008). When guinea pigs (Cavia

porcellus) are given free access to conspecific urine containing a nonvolatile fluorescent

dye, the dye is transferred to the vomeronasal and septal organs but not the olfactory

epithelium (Baxi, Dorries & Eisthen, 2006). The vaginal discharge of estrous female

hamsters contains volatile compounds that signal the presence of a female, and

nonvolatile compounds that elicit mating behaviors in males (Baxi, Dorries & Eisthen,

2006). In the mouse, detection of male pheromones by females can result in the onset of

puberty, induction of estrus and termination of pregnancy (Dulac & Torello, 2003).

Urine and vaginal mucus contain high levels of proteinaceous compounds known

as MUPs (major urinary proteins) or lipocalins that work well for pheromonal

transmission (Marchese et al., 1998; Dulac & Torello, 2003; Brennan & Keverne, 2004;

Baxi, Dorries & Eisthen, 2006). MUPs are a large family of structurally homologous

proteins with a molecular mass of 15 and 30 kDa, and the interior pockets form a highly

apolar “basket” ideally suited to bind and transport small hydrophobic molecules

(Flower, 1996; Cavaggioni & Tirindelli, 1999). The large proteinaceous compounds

appear to serve several functions including holding smaller molecules in place releasing

them over time, transporting smaller molecules into the vomeronasal organ, and large and

small molecules can form complexes with receptors (Dulac & Torello, 2003; Baxi,

Dorries & Eisthen, 2006). Since several species in a given environment may use

structurally similar compounds as pheromones, it is hypothesized that pheromone signals

are not single compounds but a blend of chemosignals (Dulac & Torello, 2003). A

136

lipocalin called aphrodisin, found in the vaginal fluid of female hamsters, attracts the attention of males and promotes copulatory activity. Aphrodisin binds volatile pheromones found in the vaginal secretions of estrus hamsters, but aphrodisin itself may have pheromonal activity (Brennan & Keverne, 2004; Baxi, Dorries & Eisthen, 2006).

Mouse urinary volatile pheromones are bound to lipocalins and these proteins are found at levels of up to 70 mg/ml in male mouse urine, reflecting their importance in chemosensory communication (Brennan & Keverne, 2004). Boar saliva contains two proteins with binding activity called pheromaxeins. This polypeptide molecule is sex- specific and isn’t found in females (Marchese et al., 1998). Preliminary bird studies show that sexual behaviors seem to be influenced by possible chemical communication roles

(Wyatt, 2010).

Anecdotally it has been reported that dolphins may utilize pheromones for communication, but there have been no reports in the literature. If a dolphin were to utilize pheromones for communication, it could be in the form of proteins and peptides acting singly or in combination with other molecules. Reversed-phase high-performance liquid chromatography (RP-HPLC) is a commonly used method for the analysis and purification of peptides and proteins (Aguilar & Hearn, 1996). In a study by Painter et al.

(1998, 2004), specimens of Aplysia were induced to lay eggs, the egg cordons collected and eluted, and the eluates fractionated by Cl8 reversed-phase HPLC to identify peptide pheromones followed by N-terminal sequence analysis for compound identification.

For this study, dolphin urine from one parous female of known reproductive state was fractionated by reversed-phase HPLC in an attempt to identify potential candidate

137

peptide or protein pheromones and serves as a preliminary investigation into the possibility of dolphin pheromonal use.

6.2 Method

6.2.1 Experimental ethics

All animals, materials and methods used in the study were individually evaluated

by the facility’s Internal Animal Care and Use Committee (IACUC) to ensure safety,

health and well-being of the animals. Samples were taken during routine behaviors in

which the animals were conditioned to participate.

6.2.2 Animals

Animals in the study (Table 1) were housed at The Mirage Hotel (Mirage; Las

Vegas, Nevada USA), a 3,141,892-L manufactured seawater pool, maintained at 21 to

23° C year-round. Animal diets consisted of frozen thawed herring (C. harengus), capelin

(M. villosus), sardine (Sardina pilchardus), and squid (L. opalescens) for daily totals of

5–10 kg per animal.

Table 6.1 Animals in study

Reproductive Animal Sex Birth Date Weight (kg) History

1 F 11/1975 191.86 Parous 2 F 3/2000 159.66 Immature 3 F 5/1997 176.44 Nulliparous 4 M 2/2003 165.15 Immature 5 M 8/1989 201.35 Parous

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1.1.1.1 Behavioral Conditioning

6.2.2.2 Urine Collection

The dolphin was trained to urinate on cue. Initially the trainer placed firm hand pressure to the dolphin’s bladder, which often resulted in reflexive urination. This reflexive behavior was paired with primary reinforcement to increase the frequency. Over time, the dolphin was conditioned to urinate by the application of gentle pressure to the bladder. The dolphin was partially pulled out of the water so that the genital slit was dry.

First, the trainer stood on a dry ledge and grasped the tail flukes of the dolphin that was stationary in a dorsal position floating at the pool surface. The dry ledge was 7 to 10 cm

higher than the water surface. Next, the trainer gently stepped backward pulling the

dolphin’s flukes backward and laterally over the ledge until the dolphin’s genital slit was

laterally out of the water. The dolphin’s head and pectoral flippers remained in the water.

The genital slit was wiped clean with gauze. A 50-ml sterile plastic specimen cup was

placed under the urethra and gentle pressure was applied to the bladder. The urine was

caught in the cup as it freely flowed from the urethra. An average of 5-10 mls of urine

was collected from all study animals. 5 ml increments of urine were immediately frozen

following collection at -17 oC. Urine for the study was collected in the am (8 am–11 pm)

for the study unless noted.

139

6.2.2.3 Ultrasonography

Real-Time B-Mode trans-abdominal ultrasound imaging was utilized. Dolphin 1 was trained to float laterally and stationary at the water’s surface. The blowhole was submerged, but the animal could lift her head and take a breath as needed. A Sonosite

180 Plus ultrasound machine with a 5-2 MHz transducer (Sonosite Inc., 21919 30th Drive

SE, Bothell, WA 98021-3904, USA) was used and images recorded using either the internal digital memory of the machine or by the use of an Archos 605 Audio/Visual

Player (Archos Inc., 7951 E.Maplewood Avenue #260 Greenwood Village, CO 80111

USA). Ultrasound exams typically lasted 3-5 minutes for both the left and right lateral positions.

6.2.2.4 RP-HPLC Analysis

RP-HPLC was performed to fractionate potential peptides and proteins in dolphin urine samples from animal 1 (Department of Neuroscience and Cell Biology, Medical

Research Building, University of Texas Medical Branch, Galveston, TX 77555-1069). 5 mls of dolphin urine was immediately frozen at -30 in 5 ml polypropylene cryovials following collection, and shipped frozen on dry ice to the laboratory. Samples were thawed, and 0.8 mls of urine was fractionated by C 18 RP-HPLC as described by Painter et. al. (1998). Hardcopies of the results were scanned into PDF digital copies using a

Fujitsu multi-sheet document scanner (Fujitsu Limited, New York, NY USA). Peaks greater than 0.1 arbitrary units (AU) and between 20 and 250 minutes were identified in each sample. Minute ranges for each peak was approximated based on visual analysis of the scanned RP-HPLC readout and recorded.

140

6.2.2.5 Amino Acid Sequencing

The sequence analysis was performed using a Perkin-Elmer Procise Sequencer

(Perkin-Elmer, 940 Winter Street, Waltham, Massachusetts 02451 USA) and methods were optimized for separation of phenylthiohydantoin (PTH) amino acid derivatives

(Department of Neuroscience and Cell Biology, Medical Research Building, University of Texas Medical Branch, Galveston, TX 77555-1069). The N-terminal sequences were compared with protein sequences stored in the University of Texas protein database to search for homologous sequences.

6.2.2.6 Progesterone, Estradiol and Lutenizing Hormone

Urinary progesterone (P) and estradiol-17 beta (E2) was measured using an

Electrochemiluminescent Immunoassay (ECLIA) tested with the Elecsys 2010 instrument

(Roche Diagnostics, Mannheim, Germany) at Clinical Pathology Laboratory in Las

ICG Canine Witness® Luteinizing Hormone Assay (Witness Synbiotics Corp., Kansas

City, MO, USA). The assay provides a rapid, semi-quantitative visible color band in the presence of dolphin LH (Muraco et al., 2009). Dolphin urine was applied to the test strip either directly following collection, or, if frozen, after thawing. A 100-µl volume of neat urine was placed onto the test strip and results were obtained in less than 1 h after application. The LH surge was identified when the test band became darker than the control band.

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6.2.2.7 Identification of spermatozoa in vaginal fluid

To collect vaginal fluid, the dolphin was partially pulled out of the water so that the genital slit was dry. First, the trainer stood on a dry ledge and grasped the tail flukes of the dolphin that was stationary in a dorsal position floating at the pool surface. The dry ledge was 7 to 10 cm higher than the water surface. Next, the trainer gently stepped backward pulling the dolphin’s flukes backward and laterally over the ledge until the dolphin’s genital slit was laterally out of the water. The dolphin’s head and pectoral flippers remained in the water. The genital slit was wiped clean with gauze. The genital slit was gently spread apart using a gloved hand and a sterile 15.24 cm cotton tipped swab with a plastic shaft (Puritan Medical Products, Guilford, Maine, USA) was inserted

14 cm into the vaginal vault, twisted 360o, and removed. The swab was immediately rolled onto a 75 mm x 25 mm glass microscope slide and allowed to air dry for 24 hours before reading. A phase contrast compound light microscope (LW Scientific,

Lawrenceville, GA USA) at 400x power was used to analyze the slides. All slides were

examined for the presence of spermatozoa.

6.2.2.8 Social Grouping

Animal 5 arrived to the facility on April 9, 2006 and remained separated from

animals 1-4 for quarantine purposes. On April 26 the water from animal 5’s pool is mixed

with animals 1-4 so the animals can see and hear one another for the first time through

closed see-through metal gates and a 30 foot canal. On May 16, animal 4 is introduced to

animal 5, animals 1-3 can see and hear animals 4 and 5 through closed see-through metal

gates. On May 18, all the animals were allowed together. On June 8, animal 1 was

separated from animals 2-5. 142

6.3 Results

6.3.1 Ultrasonography

Ultrasound exams were conducted to identify the pre-ovulatory follicle (POF) and

pregnancy in Animal 1. The POF was identified on May 27 on the left ovary and the

follicle was gone on June 8 indicating ovulation (Figures 1 & 2). Uterine fluid and an

embryo was identified on August 15 (Figure 3).

Figure 6.1 Animal 1 POF

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Figure 6.2 Animal 1 Ovulation

Figure 6.3 Animal 1 uterine fluid and embryo

6.3.2 Endocrinology

28 urine estradiol and progesterone samples were tested surrounding social

change, estrus, ovulation and pregnancy in animal 1 (Figure 4). The LH surge was

identified 24 hours prior to ovulation. uP was elevated (>1.0 ng/ml) 14 days following

ovulation.

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Figure 6.4 Animal 1 uE2, uP and uLH

6.3.3 Spermatozoa in vaginal fluid

Sperm was observed on vaginal fluid slides for 7 days between May 30 and June

6.3.4 RP-HPLC

Initially 10 samples from animal 1 were fractionated during anestrus with

lactation and no social change for the purposes of testing the RP-HPLC process on

dolphin urine (Figure 6). The process worked well and was followed by serially testing

an additional 25 urine samples taken during anestrus, social change (introduction to a

mature male), estrus, ovulation and pregnancy. An anestrus pre-social change sample was

chosen to be a baseline and 49 total peaks and 12 major peaks with sizes over 0.1

arbitrary units (AU) between 20 and 250 minutes were identified (Figure 5).

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Figure 6.5 Animal 1 baseline RP-HPLC peaks

Major baseline peaks occurred at approximate ranges of 25, 30, 35, 40, 45, 105, 115, 135, 140, and 160 minutes

In subsequent samples, peaks that occurred at differing minute ranges from baseline were considered novel. Novel peak minute ranges were compared to social grouping, reproductive status and breeding activity (Table 2). The number of novel peaks in each tested sample (n = 25) varied from 0–8 peaks (Table 3) (Figures 6 & 7).

146

Table 6.2 RP-HPLC results

L Day H of uE2 +/ cycl Soc Sp Date a - e Grpb c RP-HPLC readoutd

0 5/10 75 An 1-4, 5

5/11 76 An 1-4, 5

5/12 82 An 1-4, 5

5/13 69 An 1-4, 5 No Sample

147

Table 6.2 (continued)

5/14 66 An 1-4, 5

5/15 66 An 1-4, 5

5/16 66 An 1-4, 5

5/17 70 An 1-3, No Sample 4-5

1-3 view of 4-5

148

Table 6.2 (continued)

5/18 61 An 1-3, AM 4-5

1-3 view of 4-5

5/18 Intro No sample PM 1-5

5/19 1-5 No Sample

5/20 1-5 No Sample

5/21 1-5 No Sample

5/22 68 1-5

5/23 69 1-5

149

Table 6.2 (continued)

5/24 75 -15 1-5

5/25 69 -14 1-5

5/26 73 -13 1-5

150

Table 6.2 (continued)

5/27 75 -12 1-5

5/28 65 -11 1-5

5/29 79 -10 1-5 No Sample 5/30 70 - -9 1-5 S

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Table 6.2 (continued)

5/31 84 - -8 1-5 S

6/1 68 - -7 1-5 S

6/2 82 - -6 1-5

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Table 6.2 (continued)

6/3 84 - -5 1-5 S

6/4 77 - -4 1-5

6/5 82 + -3 1-5 S

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Table 6.2 (continued)

6/6 58 + -2 1-5 S

6/7 85 - -1 1-5 S

6/8 74 - 0 1-5

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Table 6.2 (continued)

6/22 uP +12 Preg 1.0 1-5 ng/ ml

a. Urinary estradiol pg/ml. b. social group. c. sperm on vaginal fluid slide. d. RP-HPLC readout with novel peaks circled

Figure 6.6 Novel peaks per minute range per day of study

Two unique novel peaks were only observed once each, a peak between 90-100 minutes occurred on May 18 when animal 1 had a view of animals 4 and 5 through the

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gate, and a peak between 35-45 minutes occurred on the same day the first copulation was recorded (Figure 6).

June 4 and 5 had the most novel peaks at 9 each (Figure 7), and novel peaks between the minute ranges of 115-175 occurred more often when the LH was positive

(Table 3).

Figure 6.7 Number of novel peaks per day

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Table 6.3 Novel peak minute range and reproductive status

Novel Peak Minute range Dates seen Reproductive status

25-35 June 3 Day -5, -LH, Sperm June 4 Day -4, -LH

30-40 May 31 Day -8, -LH, Sperm June 1 Day -7, -LH, Sperm June 2 Day -6, -LH June 4 Day -4, -LH June 5 Day -3, -LH, Sperm

35-45 May 30 Day -9, -LH, First Day Sperm

90-100 May 18 View of animal 4 with animal 5

115-125 June 2 Day -6, -LH June 4 Day -4, -LH June 5 Day -3, +LH, Sperm June 6 Day -2, +LH, Sperm

165-175 May 21 First sample following introduction of all animals June 3 Day -5, -LH, Sperm June 5 Day -3, -LH, Sperm June 6 Day -2, +LH, Sperm

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6.3.5 Amino-terminal (N-terminal) sequence analysis

10 samples were fractionated by RP-HPLC during lactational anestrus and static social grouping and a novel peak was identified in two of the samples in the 250 minute range (Figures 8 & 9). Amino-terminal (N-terminal) sequence analysis was conducted on the two novel samples, and the results found a milk protein fragment due to milk

contaminating the urine sample.

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Figure 6.8 RP-HPLC of samples taken during lacational anestrus and static social grouping of animal 1

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Figure 6.9 Milk protein fragment in dolphin urine

6.4 Discussion

This study showed that a healthy dolphin has peptides and proteins in her urine,

with peaks that change throughout the estrous cycle and with changes in social grouping.

The amino acid sequence of the milk protein fragment found in the urine samples taken

when she was lactating showed that RP-HPLC peaks in the urine could represent a

protein compound.

Interestingly, only a small fraction of dolphin urine, 0.8 mls, was needed to

achieve multiple peaks in each analysis, where-as a similar study in humans required over

100 mls (Heine, Raida & Forssmann, 1997). Novel peaks appeared during key points of

the social and reproductive status including: (1) when the focal animal had visual and

hearing access to a mature male dolphin she had never met, (2) when she was introduced

to the male and (3) during estrus (Tables 2 & 3) (Figures 6 & 7). This is compelling

evidence that if dolphins utilize a form of chemoreception, urine could be an ideal

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medium, but clearly much more study is needed. Future studies will include more subject animals and amino acid sequencing of all novel peaks.

Cetaceans share a common ancestor not shared with any other group of mammals, the artiodactyls (even-toed ungulates including pigs, hippos, camels and ruminants)

(Milinkovitch & Thewissen, 1997). Because of the monophyletic grouping of cetaceans and artiodactyls, a dolphin is more closely related to a cow than a horse is. Given the common ancestry to a highly chemoreceptive group like the artiodactyls, have Cetaceans completely abandoned the use of chemical communication? If dolphins do use pheromones, what mechanism would they use to send and receive them?

Urine could be an effective medium for pheromonal transmission in dolphins.

Dolphins have a very small bladder relative to their size, and they urinate frequently.

Ridgway (1972), reports that in a study by Prescott, dolphins were fed methylene blue and found to urinate at intervals of every 35 minutes. In a different study by Ridgway

(1972), urine was collected in 9 different dolphins over a 22-24 hour period resulting in

an average of 2790 mls collected per dolphin. When these two studies are combined, it

suggests that dolphins may urinate 58 mls of urine 48 times over a 24 hour period. This

could allow a dolphin to communicate pheromonal information frequently, but if urine

was a medium for pheromone transmission, how would it be received? What is known

about dolphin chemoreception?

In terrestrial mammals, chemical communication such as pheromones, acting

singly or in combination with olfaction, auditory, visual or tactile stimuli, can transmit

information such as long- and short-range navigation, relationship recognition, and inter-

and intra-sexual exchange of reproductive condition, metabolic state and social status

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(Rasmussen & Krishnamurthy, 2000). Two olfactory systems are utilized by most terrestrial vertebrates, the main olfactory system and the vomeronasal system (VNS). The main olfactory system consists of the main olfactory epithelium and olfactory bulb, which detects volatile odor molecules. The VNS includes the vomeronasal organ (VNO) and the accessory olfactory bulb, which primarily detects fluid-phase odorants such as pheromones (Marriot et al., 2013). It has been reported that aquatic mammals of the order

Cetacea do not have a vomeronsasal organ (Mackay-Sim, Duvall & Graves, 1985;

Oelschlager & Buhl, 1985), but they do possess an accessory olfactory bulb (Filimonov,

1965; Jacobs et al., 1971; Ridgway, 1990).

It is often assumed that aquatic and semi-aquatic species have a rudimentary or fully lacking sense of olfaction but this isn’t always the case. Olfactory bulbs are reduced in toothed whales before birth, leading to the conclusion that they have no olfaction sense

(Sinclair, 1966: Oelschlager and Buhl, 1985). However, olfactory brain structures are present in dolphins, and they have a well-developed olfactory tuberculum (Filimonov,

1965). According to Ridgway (1990), in the early fetal stages of the bottlenose dolphin, the olfactory bulb, nerve and tracts are present, but as the fetus develops, the structures degenerate and are absence from the mature brain. However, although the peripheral olfactory components of the cetacean nervous system are absent or rudimentary, other brain structures traditionally thought to be involved in olfaction are present in the mature dolphin. The olfactory lobes and septal areas are large whereas the hippocampus and subiculum are small. The rhinencephalon displays the same basic structural arrangement as in primates or carnivores. Ridgway states: “the seemingly paradoxical presence of a

“nosebrain” without peripheral connections are sensory olfactory endings in intriguing”.

162

Despite the absence of olfactory nerves, the tiny terminal nerves that have been observed to accompany the olfactory nerves in many vertebrate species are present in fetal and adult dolphins. Kuznetzov (1974) theorized that dolphins possess quasi-olfaction. This was based on experiments that showed dolphins could detect odorous substances such as carbonic acids. Kuznetzov further theorized that dolphins may use quasi-olfaction in chemical communication to direct behavioral actions. Kuznetzov noted that the islands of

Calleja, structures located in the forebrain, are found in dolphin olfactory tubercles

(Jacobs et al., 1971). The islands of Calleja are responsible for some odorous derived reproductive behavioral actions in some mammals (Millhouse, 1987; Talbot et al., 1988).

Aquatic mammals that forage for food underwater are unable to use olfaction in the recognizable sense, but several species do seem to utilize a form of olfaction and have chemosensory abilities. Recent advances have shown that bowhead whales have a histologically complex olfactory bulb making up approximately 0.13% of brain weight and 51% of the olfactory receptor genes were intact. This suggests that bowhead whales have a sense of smell possibly used to find krill aggregations on which they feed

(Thewissen, George, Rosa & Kishida, 2011). Semi-aquatic mammals such as the platypus have a large repertoire of vomeronasal receptors, and the star-nosed mole and water shrew are capable of detecting underwater semiochemicals (Marriot et. al, 2013). In pinnipeds (seals, sea lions and walrus), early anatomical examinations suggested that the sense of smell is of little importance for these aquatic mammals. It was noted: ‘The olfactory apparatus is variably reduced, but more reduced in phocids than in otariids and

Odobenus’ (Harrison & Kooyman 1968). Later anecdotal reports suggested that olfaction may be important for mother-pup recognition, olfactory site-recognition, and migration;

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and harbor seals (phocids) have been shown to have a very high olfactory sensitivity relevant to their sensory ecology (Kowalewsky, Dambach, Mauck & Dehnhardt, 2006).

Taste and olfaction often work together for chemoreception, and it was initially thought that cetaceans may not have taste ability based on the anatomical lacking of taste papillae on the cetacean tongue as is seen in terrestrial mammals (Sonntag, 1922). Later anatomical studies reported structures that resemble taste buds inside small pits in the

root region of dolphin tongues (Valiulina and Khomenko, 1976; Donaldson, 1977).

Finally it was concluded that anatomically, odontocete chemoreception was highly

specialized and unlike other mammals due to the reduced structure and number of lingual

taste receptors (Agarkov and Gilevich, 1979). Studies conducted by the United States

Navy proved that dolphins have the ability to distinguish salty, sweet, sour and bitter,

with most sensitivity to bitter (Nachtigall and Hall, 1984; Nachtigall, 1986). Russian

scientists hypothesized that dolphins may use taste reception in a form of chemically-

mediated communication (Yablokov, 1961; Kuznetsov, 1990). In humans,

electrophysiological studies show that the trigeminal nerve, which innervates somato-

sensation on the tongue, modulates the taste neurons which arise from the cranial nerve

VII (Sanders, 2010), and anosmia humans can distinguish between taste substances

(Laska, Distel & Hudson, 1997). The dolphin trigeminal nerve has the greatest number of

axons of any other dolphin cranial nerve (Jansen and Jansen, 1969), and the upper

rostrum and lower jaw have a rich somatic innervation (Ridgway, 1990).

Recently, great strides have been made in our understanding of vertebrate

chemoreception thanks to molecular biology (Bargmann, 2006). Odorant receptor genes

that allow chemosensation were discovered in 1991, and it is now known that olfaction

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begins with the detection of odors by G-protein-coupled odorant receptors (Bargmann,

2006). Two smaller GPCR families mediate sweet and bitter taste, and the odorant receptor family exists from fish through to mammals (Bargmann, 2006). No recent studies on dolphin chemoreception abilities utilizing molecular biology have been conducted.

Dolphins can distinguish tastes, have accessory olfactory brain structures, and a rich somatic innervation of the trigeminal nerve around their mouth, so perhaps one or a combination of these chemoreceptive abilities could receive pheromonal information.

Future testing will purify and characterize the novel peaks N-terminal amino acid microsequence analyses and mass spectrometry, so that the full-length amino acid sequence of candidate peptide/protein compounds could be identified.

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CHAPTER VII

VIBRISSAL CRYPTS OF THE BOTTLENOSE DOLPHIN (TURSIOPS TRUNCATUS)

7.1 Introduction

Most mammals, except humans, have vibrissae (whiskers, sensory hairs) on the face, and they serve as an important tactile sensory apparatus (Van Horn, 1970).

Vibrissae are used a variety of ways depending on the species and can include judging surface texture, object recognition, searching for food, prey attack, facial expression in intra-species communications, dispersion of pheromones, environmental monitoring, maintaining head position while swimming, and for underwater navigation in seals

(Moore, 2004).

The vibrissae in semi-aquatic pinnipeds (seals, sea lions, walrus) are part of an intricately structured and densely innervated sensory organ, the follicle–sinus complex

(FSC). The FSCs of pinnipeds larger than those found in terrestrial mammals, and they contain up to ten times more nerve fibers than the rat and cat (Glaser et. al, 2011). Studies on pinnipeds have shown they use their vibrissae to easily discriminate shape, size and texture of objects in the environment and work as vibrotactile sensors (Dehnhardt &

Kaminski, 1995).

All cetaceans possess some form of hair-like vibrissae of epidermal origin at some stage of their ontogeny but the functions of these structures remain unclear. King (1977) provided a review of research that had been conducted in the early to mid-1900’s on

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Mysticeti and Odontoceti vibrissae. He found that vibrissae only occur on the head of cetaceans and are primarily distributed along the margins of the upper and lower jaws and around the blowhole of some baleen whales. The rorquals (Balaenopteridae) have high numbers of vibrissae with a thick patch of 20-30 hairs arranged in two vertical rows on the chin, 15 along the edge of the lower lip and 15 more on each side of the dorsal surface of the head between the blowhole and tip of the upper jaw with an additional 6-7 hairs on either side of the blow hole. The humpback whales (Megaptera novaeangliae) possess the largest number of hairs of any cetacean with the typical pattern of the rorquals plus additional vibrissae located on the characteristic nodules of the jaws and head.

In toothed whales, vibrissal hairs or follicular crypts vary from 4–20 along the upper jaw. The Amazon River dolphin (Inia geoffrensis), Risso's dolphin (Grampus griseus) and white-beaked dolphin (Lagenorhynchus albirostris) have vibrissae that persist past infancy and often into adulthood (Harrison, 1977). Bottlenose dolphins are born with vibrissal hairs but lose them postnatally leaving behind an array of 5-8 pits or crypts on each side of the upper rostrum.

Odontoceti follicular crypts have been described as vestigial structures of the integument, having no apparent blood sinuses, blood vessels or nerve fibers (King, 1977;

Yablokov & Klezeval, 1969). However, in contrast, Japha (1907), showed that the crypts

of Cetacea are actually highly innervated, specialized organs with complex blood sinuses

similar to those of sensory hair follicles in other mammals, with the fundamental

difference that whale vibrissae are immobile (Harrison, 1977).

More recent studies of vibrissal crypts in odontocetes continue to suggest that

they are in fact functional organs but the exact function remains debatable (Czech-Damal

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et al., 2012; Mauck, Eysel and Dehnhardt, 2000). Studies of an adult Guiana dolphin

(Sotalia guianensis) suggest they are utilized in electroreception (Czech-Damal et al.,

2012), whereas studies in the Tucuxi dolphin (Sotalia fluviatilis guianensis) hypothesize they are used as a hydrodynamic receptor system (Mauck, Eysel and Dehnhardt, 2000).

Anatomically, the crypts in Czech-Damal et al.’s (2012) study of the Guiana dolphin consisted of an invagination of the epidermal integument from 4.1–7.1 mm in length and a 1.2–4.3 mm width encircled by a capillary network derived from large muscle-bound blood vessels. A dense innervation by infraorbital branches of the trigeminal nerve were identified showing 300 axons per crypt, and most nerve endings

were free nerve endings and lanceolate endings, concentrated in the lower two-thirds of

the crypt. Merkel cells were evenly scattered along the epidermal basal layer.

Intraepithelial nerve fibers frequently distributed throughout the crypts, which have not

been described in FSC’s of other mammals so far (Czech-Damal et al., 2012).

For this study, the vibrissal crypts of a 1.5 year old expired male bottlenose

dolphin were photographed in-situ, preserved and examined microscopically. Natal

vibrissae and adult crypts were photographed opportunistically while the animals

engaged in routine activities.

7.2 Method

7.2.1 Experimental ethics

All animals, materials and methods used in the study were individually evaluated

by the facility’s Internal Animal Care and Use Committee (IACUC) to ensure safety,

health and well-being of the animals. Photos of live animals were taken during routine

168

behaviors in which the animals were conditioned to participate. Photos from deceased animals were taken during necropsy.

7.2.2 Animals

Animals were located at Six Flags Discovery Kingdom (SFDK; Vallejo,

California USA) and housed in a 2717.93-L manufactured seawater pool maintained at

21-22o C year round. Animal 1 was fed a diet of whole frozen thawed herring (Clupea

harengus) and capelin (Mallotus villosus) for totals of 9 kg daily. Animal 2 was nursing.

Animal 3 was a 1.5 year old nursing male who expired of pneumonia.

Table 7.1 Animals in study

Reproductive Animal Sex Birth Date Weight (kg) History

1 F 5/1986 189.00 Parous 2 M 6/2010 13.6 Newborn 3 M 5/2010 90.00 Immature

1.1.1.1 Vibrissae

Vibrissal crypts were sectioned from formalin preserved tissue of an expired 1.5

y.o. male dolphin (Six Flags Discovery Kingdom, Vallejo, CA USA). The sectioned

crypts were analyzed using a compound light microscope (Accu-Scope, Commack, NY

USA) at 100x power. Photos were taken with an iPhone 4S (Apple Inc, Cupertino, CA

USA) with the iMicroscope software application (Perceptive Development, Woodland

Hills, CA USA).

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Natal vibrissae were opportunistically photographed underwater in a 2 day old bottlenose dolphin (Six Flags Discovery Kingdom, Vallejo, CA USA and vibrissal crypts in a 24 y.o. female bottlenose dolphin (Six Flags Discovery Kingdom, Vallejo, CA USA) using a digital SLR camera (Canon U.S.A, Melville, NY).

7.2.3 Results

7.2.3.1 Vibrissal hairs and crypts

Vibrissal hairs in a 2 day old dolphin (Animal 2) consisted of two bilateral rows of 5-6 slightly curled white hairs on each side of the upper jaw along the rostrum (Figure

1). Bilateral rows of 7 vibrissal crypts with no visible vibrissal hairs were identified in

animal 1 (Fig 2).

Figure 7.1 Natal vibrassal hairs

White arrows point to vibrassal hairs

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Figure 7.2 Mature dolphin vibrissal crypts

White circle show vibrissal crypts

Seven crypts from formalin preserved tissue were sectioned (Figure 3). Crypts

consisted of an invagination of the epidermal integument with a dense capillary network

at the terminus of the crypts (Figures 3-5) At the bottom of each crypt, an amber colored

1 mm x 3 mm piece of vibrissal hair was found (Figures 4,6, & 7). A vibrassal hair was

sliced and photographed microscopically showing an amber colored keratinous structure

(Figure 7) and a possible associated nerve fiber (not imaged).

171

Figure 7.3 Vibrissal crypts

Figure 7.4 In-situ vibrissae inside the crypts

172

Figure 7.5 Capillary bed at the crypt terminus

Figure 7.6 Formalin preserved vibrissae

173

Figure 7.7 Sectioned formalin preserved vibrissae

7.3 Discussion

Dolphins are social mammals that utilize a variety of sensory abilities to survive

in their holaquatic world. Vocalizations, hearing, echolocation, visual acuity and tactile

sense are known sensory mechanisms utilized by dolphins. However, chemoreception

(taste, olfaction and trigeminal chemosensory system) and vibrissae function has had very

little study, and it is unknown how, or to what extent, dolphins may utilize these senses in

their daily life.

In the Guiana dolphin, follicular crypts had a dense innervation by infraorbital

branches of the trigeminal nerve (Czech-Damal et al., 2012). In the bottlenose dolphin,

the trigeminal nerve has the greatest number of axons of any other dolphin cranial nerve

174

(Jansen and Jansen, 1969), and the upper rostrum and lower jab have a rich somatic innervation (Ridgway & Carder, 1990).

The trigeminal nerve is the largest of the cranial nerves carrying motor supply for mastication and transmits sensory information from the face and head (Woolfall &

Coulthard, 2001). The trigeminal nerve is responsible for somatosensory sensations such as touch, temperature, pain and the perception of humidity (Brand, 2006). Additionally, separate from taste and olfaction, the trigeminal chemosensory system detects chemical stimuli (Purves et al., 2001; Silver & Finger, 2009; Brand, 2006). The trigeminal chemosensory system consists of polymodal nociceptive neurons and axons in the trigeminal nerve typically activated by chemicals such as irritants (e.g. air pollutants, ammonia, ethanol, vinegar, menthol, and capsaicin a chili pepper compound) ( Purves et al., 2001; Silver & Finger, 2009). Interestingly, many of the compounds classified as irritants can also be recognized as odors or tastes but the trigeminal system can respond to the compounds with a much higher threshold (Purves et al., 2001). In studies of patients with anosmia perceptual thresholds for irritating compounds are 100 times higher than normal subjects who perceive the compounds as odors (Purves et al., 2001). In humans, electrophysiological studies show that the trigeminal nerve, which innervates somato-sensation on the tongue, modulates the taste neurons which arise from the cranial nerve VII (Sanders, 2010), and anosmia humans can distinguish between taste substances

(Laska, Distel & Hudson, 1997).

More study is needed, but it’s plausible that trigeminal nerve endings located in dolphin follicular crypts could act similarly to those found in the nasal mucosa (Purves et al., 2001) of terrestrial species and respond to chemical stimuli.

175

CHAPTER VIII

CONCLUSION

For cetaceans, surviving in an aquatic world has required some extreme adaptations in many aspects of anatomy and physiology including: voluntary breathing with low breath per minute ratios, bradycardia, vasoconstriction, redistribution of blood flow, and thermoregulation (Ridgway, 1972). Reproduction adaptions include a valve- like vagina to prevent salt water contamination of spermatozoa and precocious young immediately able to swim, breathe and nurse after birth. Yet many aspects of reproduction has been conserved through evolution including the attachment of the

dolphin penis retractor muscle to the hipbone (Dines et al., 2014), endocrinology,

behavior, and possibly the use of chemoreception.

This long-term study utilized conditioned dolphins in aquaria to better understand

female dolphin reproductive biology and provide a hypothesis of how dolphins may

communicate reproductive readiness to one another. Through serial urine sampling for

endocrinology, adapting a rapid lutenizing hormone assay for use with urine, ovarian

ultrasound, vaginal fluid collection and behavioral observations during non-treatment

induced estrous cycles, new insights to dolphin reproduction were discovered including

natural estrous synchrony, a return to estrous following calf weaning, a hastening of

ovulation in the presence of a mature male, standing heat estrus behavior, and vaginal

mucus conducive to sperm transport during estrus.

176

Dolphins have a 10 day estrogenic follicular and 20 day luteal phase. During the

12-24 hour LH surge, pre-ovulatory follicles reach an average of 2.0 cm and ovulation

occurs within 24-48 hours. Ovulated follicles can be cleared of all fluid in less than 2

hours. Prolactin is involved with lactational anestrus, mammary development and

lactation, mothering behavior in mothers and non-mothers, spontaneous lactation in non-

mothers, and showed no correlation between seasons indicating that dolphins don’t use

photoperiod to regulate reproduction.

Conceptive reproductive behaviors increased with estradiol levels and peak

occurrences of behaviors were observed during the luteinizing hormone surge. Two novel

behaviors, genital tracking, an investigatory-type behavior and immobility, a novel form

of standing heat estrus, were observed; these behaviors appeared conducive to

communicating reproductive readiness and increasing copulation success. A total of 314

occurrences of estrus behavior were recorded in 10 hours of footage from the three focal

females, and copulation spanned from day -9 to day 0 in one dominant female. Sexual

interactions during estrus included female-to-female, immature male-to-female, mature

male-to-immature male and masturbation with toys. During estrus, focal females received

more behavioral attention than they initiated, and passive and active dorsal fin mounting

between females was the most frequent behavior. These dolphins showed behavioral

patterns similar to those reported in estrus cows where genitals are nuzzled, females

mount and are mounted by other females, and standing heat intensity increases as LH

levels rise.

Dolphin vaginal fluid forms arboriform arrangements upon drying and can be

conductive in identifying fertility. As seen in humans, mucus type “S”, with a parallel

177

arrangement and large open mesh, was prevalent during the estrogenic follicular phase, which would allow sperm to penetrate through the cervical canal (Menarguez, Pastor &

Odeblad, 2003; Hafez & Hafez, 2000). During anestrus and the luteal phases gestagenic mucus type “G” , with no distinct crystallization and closed mesh, was the primary form observed which would make it difficult for sperm to penetrate (Menarguez, Pastor &

Odeblad, 2003; Hafez & Hafez, 2000). Mucus type “L” was seen more often during anestrus and luteal phases, and type “P” and sub-type “P4” was present primarily during the follicular phase.

Through the use of RP-HPLC, a healthy dolphin has generous amounts of

peptides and proteins present in only a small fraction of urine, with peaks that change

throughout the estrous cycle and changes in social grouping. Novel peaks appeared

during key points of the social and reproductive status including: (1) when the focal

animal had visual and hearing access to a mature male dolphin she had never met, (2)

when she was introduced to the male and (3) during estrus. Proteins and peptides in urine

could be an effective medium for pheromonal/chemical transmission in dolphins.

Dolphins have a very small bladder relative to their size, and may urinate up to 58 mls of

urine 48 times over a 24 hour period. To receive and process chemical communication,

Cetacea possess an accessory olfactory bulb (Filimonov, 1965; Jacobs et al., 1971;

Ridgway, 1990), and they have a well-developed olfactory tuberculum (Filimonov,

1965). According to Ridgway (1990), in the early fetal stages of the bottlenose dolphin,

the olfactory bulb, nerve and tracts are present, but as the fetus develops, the structures

degenerate and are absence from the mature brain. However, although the peripheral

olfactory components of the cetacean nervous system are absent or rudimentary, other

178

brain structures traditionally thought to be involved in olfaction are present in the mature dolphin. The olfactory lobes and septal areas are large whereas the hippocampus and subiculum are small. The rhinencephalon displays the same basic structural arrangement as in primates or carnivores. Ridgway states: “the seemingly paradoxical presence of a

“nosebrain” without peripheral connections are sensory olfactory endings in intriguing”.

Despite the absence of olfactory nerves, the tiny terminal nerves that have been observed

to accompany the olfactory nerves in many vertebrate species are present in fetal and

adult dolphins. Kuznetzov (1974) theorized that dolphins possess quasi-olfaction. This

was based on experiments that showed dolphins could detect odorous substances such as

carbonic acids. Kuznetzov further theorized that dolphins may use quasi-olfaction in

chemical communication to direct behavioral actions. Kuznetzov noted that the islands of

Calleja, structures located in the forebrain, are found in dolphin olfactory tubercles

(Jacobs et al., 1971). The islands of Calleja are responsible for some odorous derived

reproductive behavioral actions in some mammals (Millhouse, 1987; Talbot et al., 1988).

Taste and olfaction often work together for chemoreception, and it was initially thought

that cetaceans may not have taste ability based on the anatomical lacking of taste papillae

on the cetacean tongue as is seen in terrestrial mammals (Sonntag, 1922). Later

anatomical studies reported structures that resemble taste buds inside small pits in the

root region of dolphin tongues (Valiulina and Khomenko, 1976; Donaldson, 1977).

Finally it was concluded that anatomically, odontocete chemoreception was highly

specialized and unlike other mammals due to the reduced structure and number of lingual

taste receptors (Agarkov and Gilevich, 1979). Studies conducted by the United States

Navy proved that dolphins have the ability to distinguish salty, sweet, sour and bitter,

179

with most sensitivity to bitter (Nachtigall and Hall, 1984; Nachtigall, 1986). Russian scientists hypothesized that dolphins may use taste reception in a form of chemically- mediated communication (Yablokov, 1961; Kuznetsov, 1990). In humans, electrophysiological studies show that the trigeminal nerve, which innervates somatosensation on the tongue, modulates the taste neurons which arise from the cranial nerve

VII (Sanders, 2010), and anosmia humans can distinguish between taste substances

(Laska, Distel & Hudson, 1997). In the Guiana dolphin, follicular crypts, also known as vibrissal follicles, had a dense innervation by infraorbital branches of the trigeminal nerve (Czech-Damal et al., 2012). In the bottlenose dolphin, the trigeminal nerve has the greatest number of axons of any other dolphin cranial nerve (Jansen and Jansen, 1969), and the upper rostrum and lower jab have a rich somatic innervation (Ridgway & Carder,

1990). Dolphins can distinguish tastes, have accessory olfactory brain structures, and a rich somatic innervation of the trigeminal nerve around their mouth, so perhaps one or a combination of these chemoreceptive abilities could receive and process pheromonal/chemical reproductive information.

180

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DOLPHIN SAMPLE COLLECTION METHODS

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A.1 Voluntary Collection Techniques

The bottlenose dolphins in this study were behaviorally conditioned using positive reinforcement to participate in the serial sampling. The dolphins always had a choice of whether or not they participated in sample collection sessions. Urine, blood, and vaginal fluid samples were taken, and ultrasound exams conducted, when the dolphins voluntarily participated.

Figure A.1 Sample collection photos a. urine collection. b. blood collection. c. vaginal fluid collection

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Figure A.2 Ultrasound exams a. ultrasound exam using a waterproof box to protect the machine. b. showing location of the ovaries

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ULTRASOUND CASE STUDIES

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B.1 Dolphin ovulation

Figure B.1 Dolphin ovulation sequence a. 5/18, 900 h, preovulatory follicle 2.0 cm, LH surge. b. 5/18, 1300 h, LH surge peaking. c. 5/19, 900 h, LH surge ending, white arrow shows ovulating follicle, white outlined arrow shows fluid leaving the follicle. d. 5/19, 1300 h, ovulation complete

B.2 Two consecutive ovulations and conception

A 20 year old parous female had baseline progesterone and no follicular growth upon arriving at a new facility. She lived with two mature females with visual and hearing access to two mature males. The males and females were separated by a net fence.

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Figure B.2 Preliminary ovarian ultrasound upon arrival showed no follicular activity on either ovary. a. left ovary circled. b. right ovary circled

Figure B.3 Follicular activity c. 6/14, 30 days after arriving to the new facility follicular growth was seen. d. 6/30, white arrow shows dominant follicle, white outlined arrow shows thickened endometrium. e & f 1.3 cm dominant follicle on 7/2 and 7/3. 210

Figure B.4 Ovulation and CL development during the non-mated cycle g. 7/7, LH surge, Follicle 1.8 cm. h. 7/10, 2 days post ovulation. i. 7/18, day 10, irregular CL measuring 2.1 cm, thickened endometrium, serum progesterone 4.4 ng/ml, urine progesterone 0.5 ng/ml. j. 7/25, day 17, white arrow shows 2.8 cm CL.

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Figure B.5 CL regression and new follicular development k. 7/28, day 20, urine progesterone 1.0 ng/ml, white arrow shows 2.0 cm CL. l and m. 7/30, day 22, new follicular growth is seen. n. 8/4, urine progesterone 0.3 ng/ml, day -9, a dominant follicle is visible

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Figure B.6 Pre-ovulatory follicle during the LH surge o. 8/6, day -7, she is placed with a mature male for breeding. p. 8/9, day -4. q. 8/11, day - 2, POF 1.8 cm. r. 8/12, day -1, POF 2 cm, LH surge

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Figure B.7 Ovulation and CL development during the conceptive cycle s. 8/13, day 0, ovulation. t. 8/20, day 7, CL visible. u. 8/24, day 11, CL 2.8 cm. v. 9/4, day 22, CL 3.0 cm, urine progesterone 3.8 ng/ml

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Figure B.8 Early pregnancy w. 9/11, CL 3 cm, urine progesterone 5.3 ng/ml. x. 9/25, day 43, uterine fluid visible, urine progesterone on day 35 was 36.7 ng/ml. y. 10/9, uterine horns visible in cross section with fluid. z. 10/18, day 66, fetus and heartbeat visible.

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Dolphin fetal development

Figure B.9 Fetal development 4–24 weeks a. embryo 4 weeks. b. embryo 9 weeks. c. fetus 11 weeks. d. fetus 24 weeks

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Dolphin ovarian cysts

Figure B.10 Dolphin ovarian cysts a. paraovarian cysts. b. follicular cyst arrow, and follicle. c. paraovarian cysts. d. luteal cyst and corpus albacans

Resolution of luteal cyst using prostaglandin in a 31 year old multiparous female dolphin

Introduction

A 31 year old multiparous female dolphin showed a history of a chronic luteal

cyst. From February 2008–May 2009 she maintained an elevated progesterone ranging

from 0.5–2.2 ng/ml. This cyst resolved naturally, and she entered into a period of anestrus and baseline progesterone from June 2009–July 2010.

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Figure B.11 Serum progesterone levels in a dolphin with a reoccurring chronic luteal cyst.

In August 2010, an ultrasound showed a new follicular cyst on her right ovary, a poorly defined cl on her left ovary and elevated progesterone. The follicular cyst slowly regressed leaving behind an apparent luteal cyst along with the cystic CL on her left ovary. Elevated progesterone remained through April 2011 with levels ranging from 0.7–

1.6 ng/ml.

A prostaglandin treatment protocol was initiated using lutalyse in an attempt to resolve the cyst. When lutalyse failed, estrumate was used and the cyst was resolved.

Lutalyse and Estrumate are PGF2 α products that are widely used in the cattle and equine industries for estrus synchronization and resolution of luteal cysts. Lutalyse is a naturally occurring PGF2 α (dinoprost), whereas the synthetic Estrumate is a more potent analog of

PGF2 α (cloprostenol).

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Treatment

March 9, 2011

Serum progesterone 1.0 ng/ml.

March 10, 2011

10 mg IM injection of lutalyse

March 16, 2011

Serum progesterone 0.7 ng/ml

March 24, 2011

15 mg IM injection of lutalyse

March 31, 2011

Serum progesterone 1.6 ng/ml

April 5, 2011

1 cc/ml IM injection of estrumate

April 9, 2011

1 cc/ml IM injection of estrumate

April 14, 2011

Serum progesterone 0.7 ng/ml

April 27, 2011

Serum progesterone 0.2 ng/ml

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Figure B.12 Follicular and luteal cyst a. 8/23, follicular cyst right ovary, serum progesterone 1.2 ng/ml. b. 8/31, fluid reabsorption. c. 9/21, luteal cyst, serum progesterone 1.3 ng/ml. d. 10/14, luteal cyst, serum progesterone 0.7 ng/ml

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Figure B.13 Luteal cyst during treatment period a. 4/12, luteal cyst during treatment, progesterone 0.7 ng/ml. b. 4/19, luteal cyst during treatment. c. 4/22, luteal cyst during treatment. d. 4/25 progesterone 0.2 ng/ml

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FETAL SEXING, CERVICAL DILATION, PARTURITION AND POST-PARTUM

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Fetal sexing in-utero with ultrasound

Figure C.1 Fetal sexing a. male fetus with extended penis. b. female fetus with genital slit

Dilated cervix 1 hour before birth

Figure C.2 Dilated cervix

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Parturition

Figure C.3 Emerging flukes

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Figure C.4 Expulsion of calf

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Figure C.5 Delivery

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Figure C.6 Calf’s first breath m. birth. n-o. calf takes her first breath unassisted by the mother

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Post-partum

Figure C.7 Mammary gland appearance before and after nursing a. full mammary glands prior to nursing. b. mammary glands following nursing

Figure C.8 Behavior a. mom investigating calf by goosing genital slit. b. mom and calf sleeping 228