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Hormonal regulation of reproduction and the antler cycle in the male Columbian black-tailed deer Odocoileus… West, Nels O. 1975

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t HORMONAL REGULATION OF REPRODUCTION AND THE ANTLER CYCLE IN THE MALE COLUMBIAN BLACK-TAILED DEER (ODOCOILEUS HEMIONUS COLUMBIANUS) by NELS 0. WEST B.Sc., University of British Columbia, 1968 A THESIS SUBMITTED IN PARTIAL FULFILMENT OF - - THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY in the Department of Zoology We accept this thesis as conforming to the required standard The University of British Columbia June, 1975 In p r e s e n t i n g t h i s t h e s i s in p a r t i a l f u l f i l m e n t o f the r e q u i r e m e n t s f o r an advanced degree a t t h e U n i v e r s i t y o f B r i t i s h C o lumbia, I agree t h a t the L i b r a r y s h a l l make i t f r e e l y a v a i l a b l e f o r r e f e r e n c e and study . I f u r t h e r agree t h a t p e r m i s s i o n f o r e x t e n s i v e c o p y i n g o f t h i s t h e s i s f o r s c h o l a r l y purposes may be g r a n t e d by the Head o f my Department o r by h i s r e p r e s e n t a t i v e s . I t i s u n d e r s t o o d t h a t c o p y i n g o r p u b l i c a t i o n o f t h i s t h e s i s f o r f i n a n c i a l g a i n s h a l l not be a l l o w e d w i t h o u t my w r i t t e n p e r m i s s i o n . Department o f The U n i v e r s i t y o f B r i t i s h Columbia 2075 Wesbrook Plac'e Vancouver, Canada V6T 1W5 Date A</6V$T ^ 14?S 11 ABSTRACT The hormonal regulation of reproduction and the antler cycle in male Columbian black-tai led deer (Odocoileus hemionus columbianus) was investigated by measuring serum testosterone, testis volume, sperm production and the antler growth cycle of wild deer and of captive deer treated with methallibure and various hormones. A histological examina-tion of the p i tu i tary , adrenals, thyroids, testes and accessory sex glands of normal and methallibure-treated deer was also performed to study the functional relationships of these organs to reproduction and antler growth. Various constituents in the serum and urine of normal and methallibure-treated deer were measured to investigate the effects of methallibure on physiological function. Testosterone secretion, test is volume, sperm production, and the secretory act iv i ty of the accessory sex.glands were maximal in November, at the height of the reproductive season. During this period the mean serum testosterone level of the adult males was 10 ng/ml, test is volume 3 averaged 30 cm , and the concentration of sperm in the semen was 100 x 10^ to 700 x 10^/ml. In winter, the act iv i ty of the reproductive organs declined, u n t i l a minimum was reached i n February or March. The antlers were cast several weeks after the serum testosterone dropped below 1 ng/ml. In spring, a significant increase in spermatogenetic act iv i ty occurred, coincident with the i n i t i a t i o n of antler growth. Spermatogenesis declined in June and Ju ly , but the seminiferous tubules and accessory sex glands were s t i l l more active than in late winter. The serum testosterone level however, remained low (<1 ng/ml) throughout the antler growing period. i i i In late August or early September the testosterone level rose above 1.5 ng/ml, and velvet shedding ensued. Sperm production and the secre-tory activity of the seminal vesicles Increased markedly in the f a l l . The concentration and v i a b i l i t y of sperm was greatest between October and December, although some captive deer produced sperm a l l year round. These deer also had a higher serum testosterone level, and remained in rut longer than deer in the f i e l d . When methallibure, a non-steroidal inhibitor of gonadotrophin se-cretion, was applied in June, i t suppressed reproduction and arrested antler growth. PMS and HCG were capable of stimulating the testes of methallibure-treated deer to secrete enough testosterone to induce velvet shedding, but neither hormone by i t s e l f was effective in completely re-storing reproductive function. When methallibure was applied in Ap r i l , antler growth was prevented. The subsequent administration of prolactin, PMS, and some androgenic steroids during June and July failed to stimulate antler growth, but HCG did rejuvenate i t in one instance. During the normal period of antler growth, the testes of methallibure-treated deer did not respond to the exogenous administration of HCG by secreting testosterone, whereas both PMS and prolactin were effective in this respect. After the administration of PMS, HCG was capable of stimu-lating testosterone production, but i t was ineffective in the deer pre-viously treated with prolactin. Also, when methallibure treatment was ter-minated in the f a l l , testosterone production and spermatogenesis recovered, but the deer that had previously received prolactin did not produce mature sperm until the following spring. Thus, the testicular response may depend on the temporal sequence as well as the type of hormonal stimulation. The application of histochemical stains to the anterior pi tui tary revealed soven ce l l types, most of which, underwent cyc l i c changes that could be related to a seasonal pattern of functional act iv i ty . Hormonal release by the gonadotrophic cel ls was judged to be greatest in the f a l l , but some act iv i ty was also evident in spring and summer. The gonadotrophs of methallibure-treated deer were small and chromo-phobic, whereas other ce l l types appeared to be affected l i t t l e or not at a l l . The results of this study support the hypothesis that a gonadotrophin i s responsible for stimulating antler growth. V TABLE OF CONTENTS Page I. THE ANTLER CYCLE A. Introduction 1 B. Literature Review 4 1. Ontogeny of antlers 4 2. Sex hormones and the antler cycle 4 3. The thyroid gland and antler growth 6 4. The parathyroids and antler shedding 8 5. The adrenal glands and the antler cycle...... 8 6. The pituitary gland and antler growth 10 7. The gonadotrophins and antler growth 11 II. HORMONAL REGULATION OF REPRODUCTION AND THE ANTLER CYCLE 15 A. Introduction 15 B. Materials and Methods 21 1. Serum collection 21 2. Determination of serum testosterone by competitive-protein-binding 21 3. Estimation of testicular volume 22 4. Serum collection and evaluation 22 5. Methallibure-treatment and hormone injections 26 6. Routine measurements and tests of physiolo-gical function 27 7. Histology of the adrenals, thyroid, and re-productive organs 28 C. Results 30 1. Seasonal variation in serum testosterone and testis volume of wild deer 30 2. Seasonal variation in the histology of the testes of wild deer 30 3. Histology of the testes of methallibure-treated captive deer 39 v i Table of Contents (cont'd) Page 4. Seasonal variation in the histology of the accessory sex glands of wild deer and methallibure-treated captive deer 39 a. Seminal vesicles and ampulla 39 b. Prostate 45 c. Bulbo-urethral glands 46 5. Histology of the thyroid gland of wild deer and methallibure-treated captive deer 56 6. Histology of the adrenal cortex of wild deer and methallibure-treated captive deer 61 7. Seasonal variation in testis volume, serum testosterone, sperm production and the antler cycle of captive deer exposed to natural photoperiod and to continuous light 62 8. The effect of methallibure treatment and hor-mone injections on reproduction and the ant-ler cycle of captive deer 100 9. The effect of methallibure treatment and hor-mone injections on other aspects of physio-logical function 105 D. Discussion 110 1. The sexual cycle 110 2. The activity of the reproduction organs in the spring and i t s relationship to antler growth 113 3. The effect of methallibure on the reproductive organs 117 4. Testosterone and the antler cycle 118 5. The hormonal regulation of antler growth 120 I 6. The hormonal regulation of testicular functions 122 7. Seasonal variation in thyroid activity 126 8. The adrenal glands of wild deer and methalli-bure-treated captive deer 127 v i i Table of Contents (cont'd) Page 9. The effect of methallibure on physiological function . 128 III. CYTOLOGY OF THE ANTERIOR PITUITARY 133 A. Introduction 133 B. Materials and Methods 137 C. Results: Cytology of the pars anterior at dif-ferent times of the year in normal and me-thallibure-treated deer 139 D. Discussion 168 IV. SUMMARY AND CONCLUSIONS ..... 176 V. LITERATURE CITED 186 VI. APPENDIX A: Techniques 196 A. Testosterone determination by competitive-protein-binding ..... 197 B. OX-AB-PAS-OG Histochemical staining procedures 213 C. OX-AT-PAS-OG Histochemical staining procedures 214 VII. APPENDIX B: Tabulated Deer Data • 216 A. Seasonal changes in body weight, serum testoster-one and the antler growth cycle of captive black-tailed deer under various experimental treatments 217 B. Seasonal changes in testis volume, sperm produc-tion, and semen quality of captive black-tailed deer under various experimental conditions 252 C. Serum enzymes, metabolic wastes, cholesterol, phosphorus, calcium, proteins and thyroxin of captive black-tailed deer under various experi-mental conditions 268 D. Urine output, specific gravity, inorganic ion concentration and osmolarity of normal and methal-libure-treated captive black-tailed deer 277 E. Serum enzymes, metabolic wastes, cholesterol, phos-phorus, calcium, proteins and thyroxin of wild black-tailed deer on Vancouver Island ..... 281 v i i i LIST OF TABLES TABLE Page I. Seasonal variation in serum testosterone concen-tration and testis volume of wiid black-tailed deer on Vancouver Island 31 II. Rf values of steroids in the chloroform: ethyl acetate (80:20) Gelman 1TLC-SAF thin layer chro-matography system 201 III. Precision of CPB testosterone determinations 210 IV. Specificity: Comparison of the CPB method to the double isotope derivative technique 211 VA to XXA. Seasonal changes in body weight, serum testoster-one and the antler growth cycle of captive black-tailed deer under various experimental treatments... 217-251 VB to Seasonal changes in testis volume, sperm produc-XXB. tion, and semen quality of captive black-tailed deer under various experimental conditions 252-266 VC to Serum enzymes, metabolic wastes, cholesterol, XIIC phosphorus, calcium, proteins, and thyroxin of captive black-tailed deer under various experimen-t a l conditions 267-276 VD and Urine output, specific gravity, inorganic ion VID concentration and osmolarity of normal and me-thallibure-treated captive black-tailed deer 277-280 XXI. Serum enzymes, metabolic wastes, cholesterol, phosphorus, calcium, proteins and thyroxin of wild black-tailed deer on Vancouver Island 281 LIST OF FIGURES FIGURE Page 1 An orchidometer, 23 2 Low power photomicrograph of the testis of an adult wild deer (No. 9) in March. 33 3 Photomicrograph of the testis of an adult wild deer (No. 27) in May. 33 4 Medium-power photomicrograph of an epididymal tubule of an adult wild deer (No. 27) in May 33 5 Photomicrograph of the testis of an adult wild deer (No. 12) in July. 33 6 Photomicrograph of the testis of an adult wild deer (No. 18) in September. 35 7 Photomicrograph of the testis of an adult wild deer (No. 4) in November. 35 8 Photomicrograph of the testis of an adult wild deer (No. 22) in January. 35 9 Photomicrograph of the testis of a methallibure-treated captive deer in November. 35 10 Low-power photomicrograph of the seminal vesicle of an adult wild deer (No. 9) in March. 41 11 Photomicrograph of the seminal vesicle of an adult wild deer (No. 27) in May. 41 12 Photomicrograph of the seminal vesicle of an adult wild deer (No. 15) in July. ..... 41 13 Photomicrograph of the seminal vesicle of an adult wild deer (No. 18) in September 41 14 Photomicrograph of the seminal vesicle of an adult wild deer (No. 4) in November 43 15 Photomicrograph of the seminal vesicle of an adult wild deer (No. 22) in January. 43 16 Photomicrograph of the seminal vesicle of a methallibure-treated captive deer in November 43 List of Figures (cont'd) x FIGURE Page 17 Photomicrograph of the ampulla of an adult wild deer (No. 4) i n November 43 18 Low-power photomicrograph showing the glandular tissue of the prostate and a duct which enters the pelvic urethra. 47 19 Medium-power photomicrograph of the prostate gland of an adult wild deer (No. 9) in March 47 20 Photomicrograph of the prostate of an adult wild deer (No. 26) in May. 47 21 Photomicrograph of the prostate of an adult wild deer (No. 15) i n July. 47 22 Photomicrograph of the prostate of an adult wild deer (No. 18) in September 49 23 Photomicrograph of the prostate of an adult wild deer (No. 6) in November. 49 24 Photomicrograph of the prostate of an adult wild deer (No. 22) in January 49 25 Photomicrograph of the prostate of a methallibure-treated captive deer in November 49 26 Low-power photomicrograph of the bulbo-urethral gland of an adult wild deer (No. 9) in March 51 27 Photomicrograph of the bulbo-urethral of an adult wild deer (No. 28) in May. 51 28 Photomicrograph of the bulbo-urethral of a six-year-old wild deer (No. 15) in July. 51 29 Photomicrograph of the bulbo-urethral of an adult wild deer (No. 18) in September. 51 30 Photomicrograph of the bulbo-urethral of an adult wild deer (No. 4) In November. ..... 53 31 Photomicrograph of the bulbo-urethral of an adult wild deer (No. 25) in January. 53 32 Photomicrograph of the bulbo-urethral of a methalli-bure-treated captive deer in July. 53 L i s t o f Figures (cont'd) x i FIGURE Page 33 Photomicrograph o f the b u l b o - u r e t h r a l o f a w e t h a l l i b u r e - t r e a t e d c a p t i v e deer i n November 53 34 Low-power photomicrograph o f the t h y r o i d o f an a d u l t w i l d deer (No. 11) i n March. ..... 57 35 Photomicrograph o f the t h y r o i d o f an adu l t w i l d deer (No. 20) i n May. 57 36 Photomicrograph o f the t h y r o i d o f a s i x - y e a r -o l d w i l d deer (No. 15) i n J u l y . 57 37 Photomicrograph o f the t h y r o i d o f an adu l t w i l d deer (No. 18) i n September. 57 38 Photomicrograph of the t h y r o i d o f an a d u l t w i l d deer (No. 4) i n November. 59 39 Photomicrograph o f the t h y r o i d o f an adu l t w i l d deer (No. 22) i n January. 59 40 Photomicrograph o f the t h y r o i d o f a m e t h a l l i b u r e -t r e a t e d c a p t i v e deer i n J u l y . 59 41 Photomicrograph of the t h y r o i d o f a m e t h a l l i b u r e -t r e a t e d c a p t i v e deer i n November. 59 42 Seasonal v a r i a t i o n i n serum t e s t o s t e r o n e , t e s t i s volume, sperm co n c e n t r a t i o n and the a n t l e r c y c l e o f No. U-26, a c a p t i v e deer exposed to outdoor c o n d i t i o n s . ..... 64 43a Seasonal v a r i a t i o n i n serum t e s t o s t e r o n e and the a n t l e r c y c l e o f c a p t i v e deer No. W-l du r i n g and a f t e r exposure t o continuous l i g h t ; temperature h e l d constant at 25°C. 66 43b Seasonal v a r i a t i o n i n serum t e s t o s t e r o n e , t e s t i s volume, sperm c o n c e n t r a t i o n and the a n t l e r c y c l e o f c a p t i v e deer No. W-l b e f o r e , during and a f t e r treatment w i t h m e t h a l l i b u r e , p r o l a c t i n , HCG and LH. 68 44a Seasonal v a r i a t i o n i n serum te s t o s t e r o n e and the a n t l e r c y c l e o f c a p t i v e deer No. W-3 du r i n g and a f t e r exposure t o continuous l i g h t ; temperature h e l d constant at 25°C. 70 List of Figures (cont'd) x i i FIGURE Page 44b. Seasonal variation in serum testosterone, testis volume, sperm concentration and the antler cycle cf captive deer No. W-3 before .and during methal-libure treatment. 72 45 Seasonal variation in serum testosterone, testis volume, sperm concentration and the antler cycle of captive deer No. Y-9 before, during and after methallibure treatment. 74 46 Seasonal variation in serum testosterone, testis volume, sperm concentration and the antler cycle of captive deer No. Y-16 before, during and after treatment with methallibure, PMS and HCG. 76 47 Seasonal variation in serum testosterone and the antler cycle of No. Y-23, a captive deer exposed to natural photoperiod with a r t i f i c i a l light; tem-perature held constant at 25°C. 78 48 Seasonal variation in serum testosterone, testis volume, sperm concentration and the antler cycle . of No. Z-l, a captive deer exposed to outdoor con-ditions. 80 49 Seasonal variation in serum testosterone, testis volume, sperm concentration and the antler cycle of No. 2-1, a captive deer exposed to outdoor conditions. 82 50 Seasonal variation in serum testosterone, testis volume, sperm concentration and the antler cycle of captive deer No. B-1 before, during and after j treatment with methallibure , prolactin and HCG 84 51 Seasonal variation in serum testosterone, testis volume, sperm concentration and the antler cycle of captive deer No. B-2 before, during and after methallibure treatment. 86 52 Seasonal variation in serum testosterone, testis volume, sperm concentration and the antler cycle of captive deer No. B-3 before, during and after treatment with methallibure, PMS and HCG 88 53 Seasonal variation in serum testosterone, testis volume, sperm concentration and the antler cycle of captive deer No. C-l before, during and after treatment with methallibure, HCG and dehydroepi-androsterone. 90 List of Figures (cont'd) x i i i FIGURE Page 54 Seasonal variation in serum testosterone, testis volume, sperm concentration and the antler cycle of captive deer No. C-2 before, during and after treatment with methallibure, HCG and androstene-dione. 92 55 Seasonal variation in serum testosterone, testis volume, sperm concentration and the antler cycle of captive deer No. C-3 before, during and after treatment with methallibure, HCG, 19-nortestoster-one, and testosterone. ..... 94 56 Seasonal variation in serum testosterone, testis volume, sperm concentration and the antler cycle of captive deer No. C-5 before, during and after treatment with methallibure and HCG. 96 57 Seasonal variation in serum testosterone, testis volume, sperm concentration and the antler cycle of captive deer No. C-6 before, during and after treatment with methallibure, PMS, HCG, FSH and testosterone. 98 58 Photograph of a methallibure-treated captive deer, showing the inhibitory effect that methalli-bure had on antler growth. , 106 59 Photograph of a methallibure-treated captive deer showing the amount of antler growth that oc-curred during the administration of HCG in June and July. 106 60 Photograph showing the antlers of methallibure-treated captive deer No. W-l after they were cast... 106 61 Photograph showing the antlers of an untreated captive deer. 106 62 Diagram of a median section of the deer hypophy-sis i l l u s t r a t i n g the zonation of acidophilic and basophilic c e l l s , and the plane of sectioning 140 63 Photomicrograph of the pars d i s t a i i s of an adult male deer (No. 26) in May. 142 64 The pars d i s t a l i s of a male fawn (No. 1) in No-vember. 142 65 The pars d i s t a l i s of an adult male (No. 9) in March 144 List of Figures (cont'd) xiv FIGURE Page 66 The pars d i s t a l i s of an adult male (No. 9) in March. ..... 144 67 The pars di s t a l i s of a methallibure-treated deer in December. 146 68 The pars d i s t a l i s of an adult male (No. 26) in May. 147 69 A f o l l i c l e of Type III red-orange cells from the pars d i s t a l i s of an adult male (No. 26) in May. 148 70 The pars di s t a l i s of a methallibure-treated deer in July. 148 71 The pars d i s t a l i s of an adult male (No. 18) in September. 150 72 The pars d i s t a l i s of an adult male (No. 18) in September. ..... 150 73 The pars dist a l i s of an adult male (No. 6) t in November. ..... 152 74 The pars d i s t a l i s of a male fawn (No. 1) in November. 152 75 The pars di s t a l i s of an adult male (No. 26) in May. 154 76 The pars d i s t a l i s of an adult male (No. 12) in July. 154 77 The pars d i s t a l i s of a methallibure-treated deer in July. 156 78 The pars dist a l i s of a methallibure-treated deer in July. 156 79 The pars d i s t a l i s of an adult male (No. 26) in May. 158 80 The pars d i s t a l i s of an adult male (No. 18) in September. 158 81 Adjacent areas of the neurohypophysis and pars intermedia of an adult male (No. 18) in September. 160 82 The pars tuberalis of a methallibure-treated deer in December. 160 List of Figures (cont'd) x v FIGURE Page 83 Hypothetical scheme i l l u s t r a t i n g how various hormones may act to regulate reproduction and the antler cycle. 182 x v i ACKNOWLEDGEMENTS I am d e e p l y i n d e b t e d t o my s u p e r / i s o r , D r . H . C . Nordan , s f o r h i s p a t i e n c e , u n d e r s t a n d i n g , and f i n a n c i a l s u p p o r t d u r i n g the extended c o u r s e o f t h i s s t u d y . Of the many o t h e r peop le who a s s i s t e d me i n v a r i o u s ways, the f o l l o w i n g d e s e r v e s p e c i a l m e n t i o n : M r . P . Whi tehead , C a n a d i a n W i l d l i f e S e r v i c e , V a n c o u v e r , f o r h i s a s s i s t a n c e d u r i n g the i n i t i a l s tages o f t h i s i n v e s t i g a t i o n , and f o r h i s h e l p f u l s u g g e s t i o n s d u r i n g p r e p a r a t i o n o f the manu-s c r i p t . D r . J . M. T a y l o r and D r . W. S. H o a r , Department o f Z o o l o g y , f o r t h e i r p o s i t i v e c r i t i c i s m o f the m a n u s c r i p t . M r . I . Smith and M r . G.W. S m i t h , F i s h and W i l d l i f e B r a n c h , Nanaimo, f o r t h e i r a u t h o r i z a t i o n and c o o p e r a t i o n i n the c o l l e c -t i o n o f s t u d y spec imens . D r . D. C . Thomas, C a n a d i a n W i l d l i f e S e r v i c e , Ot tawa, f o r h i s a d v i c e and a s s i s t a n c e i n the i n i t i a l c o l l e c t i o n o f h i s t o l o g i c a l m a t e r i a l from the f i e l d , and t o M r . G . J o n e s , M r . R. Ledoux , M r . R. McLennan and M r . M. Michaud f o r t h e i r a s s i s t a n c e on subsequent c o l l e c t i n g t r i p s . Ms. D. H a r d s , f o r her c o o p e r a t i o n and t e c h n i c a l a s s i s t a n c e i n the p r o c e s s i n g and s t a i n i n g o f h i s t o l o g i c a l m a t e r i a l , e s p e c i a l l y f o r her e f f o r t s w i t h the p i t u i t a r y s t a i n s . D r . R . L . H . Honore , Department o f P a t h o l o g y , f o r h i s c o n s u l -t a t i o n on p i t u i t a r y c y t o l o g y . x v i i M r . S. G i n g r a s , M r . J . J y r k a n n e n , M r . Y . P e r r o n , M r . A . T e p p e r , and my b r o t h e r , M r . C . West , f o r t h e i r a s s i s t a n c e i n c o l l e c t i n g s e r e o f the b l o o d and semen samples from the c a p t i v e d e e r . D r . M. T a i t , Department o f A n i m a l S c i e n c e , f o r h i s c o o p e r a -t i o n and a s s i s t a n c e i n p r e p a r i n g the m e t h a l l i b u r e - t r e a t e d f e e d . D r . W.D. K i t t s , Department o f A n i m a l S c i e n c e , f o r g r a n t i n g p e r m i s s i o n t o use t h e i r e l e c t r o e j a c u l a t o r . D r . F . D i l l , Department o f M e d i c a l G e n e t i c s , f o r the use o f t h e i r p h o t o m i c r o s c o p e . D r . Ian McTaggart -Cowan, Dean o f Graduate S t u d i e s , f o r the i n s p i r a t i o n he p r o v i d e d e a r l y i n my c a r e e r , f o r h i s encouragement and h e l p f u l s u g g e s t i o n s d u r i n g my s t u d y , and f o r h i s c o n t i n u e d w i l l i n g n e s s t o extend p e r s o n a l s u p p o r t . 1 I. THE ANTLER CYCLE A. Introduction The Coast deer, or Columbian black-tailed deer, (Odbcoileus hemionus  cdlumbianus) inhabits the western slope of the Pacific Coast Range of North America and the adjacent offshore islands. Like other members of the Cervidae l i v i n g i n North America, the black-tailed deer is a seasonal breeder. At latitude 49°N, the bucks begin rutting in September and mating usually occurs in November. In January, as the rut declines, the bucks be-gin to drop their antlers, and by the middle of March antlers are rarely seen. New antler growth is not conspicuous unt i l late A p r i l , but by the end of July the antlers are f u l l y grown. When the antlers have reached their f u l l size they undergo a process of internal ossification (matura-tion) . By late August or early September, the velvet is shed, and the antlers are f i n a l l y transformed into dense, compact bone. Following this, each antler gradually "dies" back toward i t s base, but they remain firmly attached, and are retained in this condition until the end of the rutting season. The capacity of the male deer (and female reindeer and caribou) to regenerate bone in the form of antlers i s a remarkable phenomenon. During the month of June, the rapidly growing antlers of an adult black-tailed buck may attain a growth rate of 1 cm/day, and reach a f i n a l length of 60 cm. or more (personal observations). However, this growth rate i s slow compared to that of the antlers of a bu l l elk (Cervus canadensis) or caribou (Rarigifer arcticus). In these species, the antlers may grow as much as 3 cm/day, and reach a length of 2 meters (Goss, 1963; personal observations). 2 The growing a n t l e r s o f a b u l l moose (Alces a l c e s ) , the l a r g e s t member o f the deer f a m i l y , may i n c r e a s e i n weight by over 300 gm/day (Bubenik, 1966), and reach a t o t a l weight o f 35 kilograms. The a n t l e r s o f the red deer s t a g (Cervus elaph u s ) , although s m a l l e r , are a l s o i m p r e s s i v e . In f o u r months a mature male may grow a set o f a n t l e r s equal to one-half i t s s k e l e t a l weight! (Bubenik, 1966). To grow a n t l e r s of t h i s s i z e i n such a s h o r t time undoubtedly places a c o n s i d e r a b l e burden on the animal's mineral metabolism; nev e r t h e l e s s these e x t r a o r d i n a r y s t r u c t u r e s are l o s t and r e p l a c e d each year. The renewal of these unique appendages has aroused the c u r i o s i t y o f s c i e n t i s t s s i n c e the time o f A r i s t o t l e , because i t i n d i -cates that deer have c e r t a i n regenerative powers not found i n other mammals. The seasonal nature and p r e c i s e t i m i n g of c e r t a i n events w i t h i n the a n t l e r c y c l e s t r o n g l y i m p l i c a t e the involvement o f one o r more environmen-t a l f a c t o r s i n i t s c o n t r o l . These s t i m u l i probably act through v a r i o u s p h y s i o l o g i c a l mechanisms to r e g u l a t e a l s o a number of other c y c l i c events such as moulting, body weight changes, and v a r i o u s a c t i v i t i e s a s s o c i a t e d w i t h r e p r o d u c t i o n . Since i t appears t h a t at l e a s t some o f these events may be under the i n f l u e n c e o f the same hormones ( i . e . the gonadotrophins and t e s t o s t e r o n e ) , i t i s reasonable to c o n s i d e r both r e p r o d u c t i o n and the a n t l e r c y c l e as r e l a t e d phenomena that are r e g u l a t e d by an i n t e g r a t e d system o f hormonal c o n t r o l . One might then expect to f i n d c e r t a i n features about the hormonal c o n t r o l of r e p r o d u c t i o n which provide clues to the r e s p e c t i v e r o l e s of the p i t u i t a r y and gonadal hormones i n r e g u l a t i n g the a n t l e r c y c l e . Some o f the endocrine i n f l u e n c e s on reproduction and the a n t l e r c y c l e are w e l l documented. Goss (1963, 1964, 1968a, 1968b,1969, 1970) has e x t e n s i v e -l y reviewed a l l aspects o f a n t i e r o g e n e s i s i n deer, and i n a d d i t i o n has 3 performed a number of experiments r e l a t e d to a n t l e r growth and a n t l e r shedding. Previous experiments i n d i c a t e chat c e r t a i n events within the a n t l e r c y c l e , such as the maturation of a n t l e r s , shedding of the v e l v e t , and the maintenance of antlers i n the hard, fu n c t i o n a l condition, are probably dependent on the sex hormone , testosterone. Those factors responsible f o r i n i t i a t i n g new a n t l e r growth however, have yet to be de-termined. There i s reason to b e l i e v e that the stimulation of a n t l e r growth i s d i r e c t l y c o n t r o l l e d by the p i t u i t a r y gland, because castrates grow antlers whereas hypophysectomized deer do not (Goss, 1963; H a l l , Ganong and T a f t , 1966). Several p i t u i t a r y hormones are probably involved, e i t h e r d i r e c t l y or i n d i r e c t l y , i n the regulation of both the gonadal and a n t l e r c y c l e s . The purpose o f my study was to t e s t several hypotheses concerning the r o l e of the gonadotrophins and testosterone i n reg u l a t i n g these c y c l e s . Some of these hypotheses and my j u s t i f i c a t i o n f o r t e s t i n g them are presented i n the next s e c t i o n , along with a summary and a review o f the l i t e r a t u r e on a n t l e r c y c l e s . Since t h i s s e c t i o n contains background information concerned mainly with the a n t l e r c y c l e , those who are f a m i l i a r with research i n t h i s area may wish to proceed d i r e c t l y to Section II (p. 15 ) where some of the l i t e r a t u r e on reproduction i s reviewed, and the s p e c i f i c objectives of thf.s study are presented. 4 B. Literature Review 1. Ontoger." of Antlers During the first year the frontal bones of male fawns develop a pair of bony protruberances called pedicles, which later give rise to antler tissue. If a fawn is castrated within a few months of birth the pedicles fa i l to develop unless testosterone is administered (Wislocki et^  al_., 1947). If testosterone is administered to female fawns they will also develop pedicles, but unlike the males, they do not subsequently grow ant-lers. However, pedicle development and antler growth can be induced in ovariectomized females by administering injections of testosterone pro-pionate (Wislocki ej^al_., 1947). If estrogen is injected repeatedly into male fawns, pedicle development is prevented (Goss, 1970). From the foregoing i t appears that in female deer, ovarian secretion of estrogens prevents the occurrence of antlers, whereas in the male, testosterone promotes the development of the pedicles, which subsequently give rise to antler tissue, whether the testes are intact or not. 2. Sex hormones and the antler cycle An annual antler cycle is common to a l l species of deer although there are species differences in the pattern of events occurring within each cycle. The antler cycle can arbitrarily be divided into five phases: (i) the rest period, when there are no antlers and no growth occurs; (ii) the growth or regenerative phase; ( i i i ) the maturation phase, when secondary ossification, followed by shedding of the velvet occurs; (iv) the functional period when the antlers are maintained in a hard, polished condition; and 5 (v) the period when the antlers are cast, followed by healing of the pedicles. The anulers may be cast at different times of the year in different species, and younger males usually drop their antlers later than older ones (Goss, 1963) but in nearly a l l temperate and northern species the functional period starts in the f a l l and the growth period starts in the spring. In caribou, where both sexes carry antlers, the pregnant females commonly retain their antlers u n t i l shortly after parturition, whereas barren females usually drop their antlers earlier in the season (Espmark, 1971). Goss (1963) suggested that in female caribou, higher circulating levels of the sex hormones during pregnancy might delay antler shedding. In male deer the influence of sex hormones on the antler cycle has been examined by observing the effects of castration and hormone replacement therapy. The results of these experiments, reviewed by Goss (1963, 1968a, 1970), can be summarized as follows: If pedicle development has occurred and castration is performed dur-ing the antler growth phase the antlers do not mature, i.e. complete ossi-fication, followed by shedding of the velvet, does not take place. These antlers remain permanently in velvet, and renewed growth occurs during the appropriate season each year thereafter. I f castration i s performed after the antlers have matured, i.e. the velvet has been shed, the antlers are cast prematurely. If injections of either testosterone or estrogen are administered following castration, the antlers are retained for as long as the therapy is continued. In both intact and castrate bucks, testosterone and estrogen injections w i l l also inhibit the regeneration of new antlers, 6 or result i n a cessation of growth i f i t has already begun. In the latter instance, the partly grown antlers become ossified and the velvet is shed. In contrast to the effects of testosterone and estrogen, progesterone ad-ministered to normal deer (Waldo and Wislocki, 1951) and castrate deer (Waldo and Wislocki, 1951; Goss, 1963) had no effect on antler growth and did not induce shedding of the velvet. The results of the castration experiments led Wislocki (1943) to pos-tulate that in male deer testosterone promotes the maturation of antlers, shedding of the velvet, and i s responsible for the maintenance of antlers in the hard, functional condition. The results of the hormone injection experiments reported by Wislocki et_ al_. (1947) and Goss (1963) appear to support this hypothesis. As an additional test of Wislocki's hypothesis, I decided to measure the serum testosterone levels of captive black-tailed deer, to find out whether the serum testosterone level increased at the time of velvet shedding, and declined prior to casting of the antlers. Seasonal variation in the secretion of the gonadal hormones might be sufficient to explain the occurrence of certain events within the antler cycle. Those factors which are responsible for the i n i t i a t i o n of new growth, however, have yet to be determined. In this respect, no endocrine organ has been completely ruled out as a possible source of the antler growth stimulating hormone. 3. The thyroid gland and antler growth i In view of i t s role in the differentiation of bone and the growth of teeth in various mammalian species, one might expect the thyroid gland to influence antler growth. There i s , however, some confusion and disagreement 7 regarding the specific role that the thyroid gland might play in relation to antler growth. Grafflin (1942), was imable to detect any significant seasonal changes in the histological appearance of the thyroid gland in white-tailed deer (Odocoileus hemionus borealis). Histological examina-tion of the thyroids of red deer (Freundova, 1955; Pantic and Stosic, 1966) and roe deer (Capreolus capreolus) (Pantic and Stosic, 1966), showed that an increase in activity occurs during the spring and summer, and a decrease in activity occurs in the f a l l and winter. Yousef and Luick (1971) re-ported no significant changes i n thyroxin secretion rate at different times of the year in reindeer, but their sampling was inadequate. Gist (1971) recorded a higher radioiodine uptake by the mule deer (Odocoileus hemionus  hemionus) thyroid in winter than in the spring and summer, but he suggested that the total (hot and cold) iodine uptake was probably less in winter be-cause of a reduced food intake. Silver et a l . (1969) found the metabolism of white-tailed deer to be lowest in winter and highest during the summer. Changes in thyroid weights showed a similar trend (Hoffman and Robinson, 1966) although the thyroid weights began to drop several months earlier than the metabolic rate. Enhanced antler growth was noted in roe bucks that received thyroxin injections as fawns (Lebidnisky, 1939) and as adults (Bubenik, 1966). Curiously however, these effects did not become apparent until the following year. Bruhin (1953) found that oral administration of 1 mg thyroxin per day for one month during the antler growing period had no effect on antler growth in roe deer, but in yearling reindeer and fallow deer (Dama dama) thyroxin injections induced strikingly greater antler growth than in controls (Goss, 1963). (In the case of the roe deer, the oral dose may have been inadequate). According to Goss (1963) , Wislocki 8 thyroidectomized a two-month old white-tailed fawn, and i t subsequently showed normal antler growth. Thus, xt appears that the thyroid gland can influence antler growth but i t s hormones may act synergistically with the antler growth stimulat-ing hormone to promote, rather than to i n i t i a t e antler growth. 4. The parathyroids and antler shedding. Owing to the action of parathormone in the resorbtion and deminerali-zation of bone, one might expect the parathyroids to be involved i n the antler shedding process. However, there is no evidence to indicate that antler shedding is under the direct control of the parathyroids. Grafflin (1942) and Urschel (1967) were unable to detect any significant seasonal changes in the histological appearance of parathyroids in either white-tailed deer or mule deer. Goss (1963) was unable to induce the premature shedding of antlers in three adult sika bucks (Cervus nippon) by injecting 2 x 1 gm/wk of powdered ox parathyroid glands for 8 weeks in the autumn. 5. The adrenal glands and the antler cycle Since the adrenal glands produce steroid hormones closely related to the sex hormones and their precursors, i t has been suggested that the adrenals might be involved in regulating the antler growth cycle. Bubenik (1966) proposed that i n phylogenetically younger genera such as reindeer and caribou, the adrenals have an important influence on antler growth, whereas in older genera e.g., Cervus and Odocoileus, they are less impor-tant, and in the oldest genera, e.g., Capreolus and Muntiacus, they have no effect. It i s well known that the antler cycles of reindeer and caribou are less dependent on gonadal hormones than are other species of deer. Thus, castrated reindeer of both sexes continue to show an annual cycle of antler replacement, although the antlers are less c a l c i f i e d , the velvet is not shed, and the antlers are dropped later than normal (Tandler, 1910; Tandler and Grosz, 1913; Hadwen and Paliner, 1922; Fisher, 1939; Goss, 1963). Although i t is possible that the adrenal cortex re-leases a sufficient quantity of sex steroids to influence the antler cycle, the exact nature of this influence was not explained by Bubenik (1966), and there is l i t t l e evidence to support his proposal. Occasional-l y , antler-bearing female deer of the genus Odocoileus have been found, and in one such case this was attributed to a masculinizing adrenal tumor (Doutt and Donaldson, 1959). Wislocki (1943) reported that there were no apparent seasonal variations in the histological appearance of the adrenals of male white-tailed deer. . Hucin (1957) however, noted an increase in size and cellular activity of the zona glomerulosa during the period of antler growth in red deer, and a decrease in activity after the antlers matured and the velvet was shed. Bubenik (1966) proposed that a direct connection exists between the mineralocorticoids and the ossification of antlers, because he and Tachezy were able to stop the growth of antlers through the administration of cortisone (the implications of this are un-certain, because cortisone is a glucocorticoid). Taft et^al_. (1956) how-ever, found that ACTH injections had no effect on the antlers of white-tailed deer, and Goss (1963) was unable to induce shedding of the velvet in castrated sika deer by injections of cortisone (500 mg/wk for 8 weeks). 10 In addition, cortisone administered (250 mg 2 x weekly) to a sika buck castrated a^ter the velvet had been shed failed to prevent the antlers from being cast prematurely (Goss, 1963). Any possible role that the adrenal gland might play in regulating the antler cycle is obscured by the other functions i t performs, which may also be affected by season. 6. The pituitary glartd and antler growth It i s reasonable to expect that the overall hormonal control of re-production and the antler cycle is regulated by the hypothalamus and the pituitary gland. Wislocki (1943) suggested that the pituitary gland ex-erts a direct influence on antler growth by producing a hormone which s t i -mulates antler growth. Wislocki et^ al_. (1947) postulated further that the male counterpart of the lactogenic hormone (prolactin, LTH) might be the antler growth stimulating hormone. Bubenik (1966) proposed that growth hormone (STH) and not prolactin, stimulates antler growth. However, STH appears to be species specific, and so Bubenik (1966) argued that the nega-tive effect of bovine STH on antler growth did not disprove his hypothesis. In the winter of 1965, Hall, Ganong and Taft (1966) hypophysectomized eleven 6-month old male white-tailed fawns, only one of which survived. This animal was maintained on 25 mg of cortisone daily. Antler growth did not occur in this animal during the next summer. At this time 2 gm of bovine growth hormone administered daily in 100 mg intramuscular doses failed to induce antler growth. The above authors also reported that: 11 1) 50 mg of testosterone propionate administered three times within a month had no appreciable effect on antler growth, (it i s not clear what the investigators expected to find in this case, except perhaps that a low dose of testosterone might stimulate antler growth). 2) thyroxin did not affect the re-growth of antlers (no informa-tion i s given as to how this treatment was applied). 131 3) I had no effect on antler growth (unpublished observa-tions) . 4) the effects of hypophysectomy on the antlers may have been due to the absence of pituitary hormones other than ACTH, TSH and the gonadotrophins ( i t is not clear why the gonadotrophins were ruled out in this case). Blauel (1935) reported that injections of whole pituitary powder did not influence the growing antlers of roe deer, but the hormonal properties and effects of this type of preparation are uncertain. Taft et a l . (1956) found that large doses of growth hormone had practically no effect on ant-ler growth. Goss (1963) reported that no effects were observed on the mature antlers of two sika deer injected with 1 gm of lyophilized, defatted beef pituitary twice a week for two months. Although i t appears that the pituitary gland is involved i n stimulating antler growth, the exact nature of this influence has yet to be established. 7. The gonadotrophins and antler growth Although shedding of the old antler i s not a necessary prerequisite to the regeneration of a new one (Goss, 1963; personal observations), Tachezy 12 (1956) thought that antler shedding and regrowth might be controlled by a common stimulus, possibly the gonadotrophins. To test this hypothesis, Goss (1963) castrated seven sika bucks in the autumn, and then injected these deer with the contraceptive Enovid, and also a purified preparation of i t s main ingredient norethynodrel, in an attempt to inhibit gonadotro-phin secretion by negative feedback. This treatment effectively delayed antler shedding until after the injections were discontinued. However, Goss also noted during the course of these and other experiments, that norethynodrel was as effective as testosterone or estrogen in inducing shedding of the velvet in castrated sika deer. Therefore, norethynodrel could have exerted a direct steroidal effect on the antlers, which would also inhibit antler shedding. On the other hand, i t is not clear why nor-ethynodrel, which could be expected to mimic the effects of progesterone, was effective i n inducing shedding of the velvet, whereas progesterone was incapable of producing this effect. Goss (1963) also attempted to induce antler shedding by injecting 2 x 5000 I.U. HCG weekly for four months into four normal antler-bearing sika deer. The failure of this treatment to i n -duce antler shedding may be attributed to the following: 1) HCG, which i n mammals has effects similar to LH or ICSH, pro-bably stimulated the testes to produce additional amounts of testosterone, which in turn would inhibit antler shedding. 2) Even without the HCG injections the testosterone level should normally be high enough from November to January to prevent antler shedding. In fact, Jaczewski, and Galka (1970) and Jaczewski and Topinski (1970) found 13 that HCG injections started just prior to the presumed date of antler shedding delayed loss of the antlers in red deer and in fallow deer. Thus i t seems unlikely that antler shedding is actively induced by gonadotro-p i n secretion. Perhaps Wislocki's (1943) suggestion that antler shedding simply occurs as a result of the withdrawal of testosterone is sufficient to explain this phenomenon. The gonadotrophins however, might s t i l l be involved in stimulating new antler growth. Wislocki (1943) suggested that gonadotrophin secretion in white-tailed deer does not occur until two months after i n i t i a l antler growth. Until recently, there was no reason to question this, because i t is in keeping with the idea that the main role of the gonadotrophic hormones i s to stimulate testosterone secretion and spermatogenesis as the breeding season approaches. Markwald (1968) however, found a slight increase i n the testicular activity (both tubular and i n t e r s t i t i a l ) of mule deer in A p r i l , which coincided with the i n i t i a -tion of antler growth. This brief activation of the gonads raises the possi b i l i t y that the gonadotrophins are released at this time. It is i n -teresting to note that tropical species such as the hog deer (Hyelaphus  pOrcinus) and axis deer (Cervus axis) are capable of f e r t i l e mating while the antlers are in velvet, and the Venezuelan deer (Odocoileus gymnotis), has mature sperm in abundance regardless of whether or not the velvet has been shed (Goss, 1963). Apparently the gonadotrophins are secreted during the antler growing period in these deer. When the seasonal patterns of FSH and ICSH secretion are known, this information might provide some clues to their respective roles in regulating reproduction and the antler cycle. The influence of a third (gonadotrophic?) hormone, prolactin, also needs to be studied. Perhaps only one of these hormones is required to i n i t i a t e 14 antler growth, whereas a l l three are involved in regulating reproduction and the angler cycle. Earlier investigators made some progress in determining the hormonal control of the antler cycle through castration experiments and hormone re-placement therapy. One objective of my study was to apply this same prin-ciple to an investigation of pituitary function in deer by chemically i n -hibiting gonadotrophin secretion with methallibure ( I . e . I . 33828), and follow this by injecting various hormones. This approach avoids the drastic consequences of complete hypophysectomy, and since methallibure i s non-steroidal, i t also avoids the problems encountered by Goss in his experiments with Enovid and norethynodrel. 15 II HORMONAL REGULATION OF REPRODUCTION AND THE ANTLER CYCLE A. Introduction In A p r i l , 1966, Cowan and Nordan i n i t i a t e d an experiment to study the effect of constant photoperiod on the antler and gonadal cycles of Columbian. black-tailed deer. Two males were subjected to a photo-period of 12 hours light and 12 hours dark (12L/12D), while two others were exposed to continuous illumination (24L). Temperature was held constant at 25°C. For two years the deer kept under 12L/12D did not shed their antlers. During the second year some antler growth occurred around the base of the old antlers. This new growth was abbreviated and evidently hindered by the presence of the old antlers (which remained firmly attached to the pedicles), but the fact that i t occurred at a l l posed a problem in relation to the hormonal control of antler cycles. In view of earlier statements about the relationship of testosterone to the antler cycle one might expect that i f the level of testosterone is s u f f i -cient to maintain the antlers in a hard, functional, condition i t should also inhibit new growth. This obviously was not the case in these deer and I decided to find out what the circulating testosterone level might be under these unusual circumstances. Thus, the i n i t i a l objective of this study was to measure the serum testosterone levels of deer kept under na-tural and a r t i f i c i a l photoperiods, and to relate this to the antler cycle. In July, 1969, two adult males were added to serve as indoor controls. They were maintained under a natural photoperiod provided by a r t i f i c i a l illumination (room temperature held constant at 25°C). Two others were ut i l i z e d as outdoor controls (natural photoperiod; variable temperature). ( 16 A t the same time, a series of tests were started to determine whether a new technique u t i l i z i n g competitive-protein-binding would be suitable for measuring testosterone in deer serum. These experiments were in progress when Goss (1969) published the results of his work on the effect of light on antler cycles in deer. He showed that deer native to temper-ate zones grow antlers in relation to the phase and frequency of the an-nual light cycle. Goss suggested that deer have an endogenous circennian rhythm which may express i t s e l f in the absence of these cues, but this rhythm is normally entrained by annual variations in the photoperiodic cy-cle. In view of these findings the experiments pertaining to photoperiod in the present study were discontinued, and greater emphasis was placed on the endocrine aspects of reproduction and antler growth. The sexual cycles of various species of deer have been studied by many investigators, only some of whom w i l l be mentioned here. The histology and histochemistry of the testes and accessory sex glands has been studied in white-tailed deer (Wislocki, 1943, 1949; Robinson et_ al_., 1965), sika deer (Wislocki, 1949; Morii, 1956), roe deer (Stieve, 1950; Short and Mann, 1966), mule deer (Markwald, 1968), red deer (Aughey, 1969; Lincoln, 1971), and fallow deer (Chapman and Chapman, 1970). A seasonal cycle of testicular and accessory sex gland activity was found in a l l the above species. The period of maximal activity occurred during the breeding sea-son, and the organs were least active during the sexually quiescent period following the rut. Many investigators (including some of those mentioned above) have traditionally considered the testes to be functionally inactive during the spring and summer months. However, Robinson et^al_. (1965) noted that the seminiferous tubules of white tailed deer were slig h t l y more active during the antler growing period, and Markwald (1968) detected a temporary rise in the testicular activity of mule deer which coincided with the i n i t i a t i o n of antler growth in Apr i l . In the present study the testes and accessory sex glands of Columbian black-tailed deer were examined histologically to determine whether a similar period of reduced activity could be found. If so, this could re-flec t hormonal changes which might also be involved in i n i t i a t i n g antler growth. In addition, the reproductive organs of wild deer were studied so they could be compared to the normal and methallibure-treated captive deer. Of the many studies describing the sexual cycles of deer, few have included an investigation of the hormonal changes that accompany or regu-late these cycles. Short and Mann (1966) measured the concentration of testosterone i n the testes of roe deer and related this to the sexual cycle. Testosterone was low in January, increased gradually during the spring, and reached a maximum during the rut. A dramatic f a l l occurred at the end of the; rut and after that the concentration remained low until January. Lincoln (1971) measured the concentration of testosterone and andros-tenedione i n the testes of red deer stags and related these to the physio-logical changes associated with puberty. The testes of immature males showed an increase in testosterone concentration and content associated with development of the pedicles. Testosterone content was highest during the rut and then declined to a low level u n t i l the following summer. In the two-year-old males, testosterone increased gradually during the summer months, reached i t s highest level during the rut, and then declined to a 1 8 low level after the breeding season. The competitive-protein-binding technique u t i l i z e d in the present study-was also used to measure plasma testosterone in captive caribou and rein-deer for Whitehead and McEwan in 1970-71 (published in 1973). In these species the testosterone level was low until the velvet was shed in August, and reached a peak during the rut. At the end of the breeding season the plasma testosterone f e l l rapidly to very low levels (less than 1 ng/ml), and remained low un t i l the onset of the next reproductive season. The antlers were cast 2 to 8 weeks after the decline in testosterone secretion. In the present study, serum testosterone was measured in wild deer and in normal and experimentally-treated captive deer to test the hypothesis that testosterone induces the maturation of antlers and shedding of the velvet, and is responsible for the maintenance of antlers in the hard, functional condition. In addition, this information would verify that the testosterone levels were highest during the breeding season, and lowest dur-ing the non-reproductive, antler-growing period. Semen collected by electroejaculation from normal and dieldrin-treated white-tailed deer during the last month of the rutting season was examined by Bierschwal et^ al_. (1970) . Semen has also been collected from reindeer during the breeding season (Dott and Utsi, 1971), but no information on the seasonal pattern of sperm production was obtained in either of the above studies. During the last two years of the present study semen from normal and experimentally-treated captive deer was collected by electroejaculation and examined to determine the seasonal pattern of sperm production, and 19 also to study1 the effects of methallibure and hormone therapy on sperma-togenesis. Estimation of testicular volume with an orchidometer was also incorporated as a routine procedure during this part of the study. In 1961, Paget et_al_. reported that certain derivatives of dithiocar-bamoylhydrazine (e.g. methallibure, I.C.I, compound 33828), produced ef-fects i n rats and some other animals indicative of selective inhibition of pituitary gonadotrophin function (Parkes, 1963; Walpole, 1965). The overt effects were suppression of the estrous cycle and f o l l i c u l a r maturation in the female; atrophy of the testes, failure of spermatogenesis, and re-duction in the size of the prostate and seminal vesicles in the male. These effects could be prevented by the simultaneous administration of exoge-nous gonadotrophin, so they were almost certainly caused by the inhibition of pituitary gonadotrophin rather than by direct desensitization of the gonads. Studies on the effects of methallibure in the pregnant mare indi-cate that the secretion of gonadotrophins of non-hypophysial origin is not suppressed (Schmidt-Elmendorff, et_ al_. , 1962). Malven (1971) found that methallibure inhibits pituitary gonadotrophin secretion i n the guinea pig by acting on the hypothalamus. Numerous literature citations on the ef-fects and actions of methallibure in laboratory and domestic animals are available in an extensive bibliography compiled by Imperial Chemical Indus-tries Limited. In the present study, methallibure was administered to captive deer to determine i t s effect on reproduction, and to test the hypothesis that the gonadotrophins are responsible for i n i t i a t i n g antler growth. Antler growth, sperm production and serum testosterone were measured routinely in normal and methallibure-treated deer, before and after the administration of 20 exogenous gonadotrophins and several androgenic steroids. Since methallibure can also inhibit other aspects of pituitary f u n c -tion (e.g., oxytocin release; Benson and Zagni, 1965; Garbers and Fi r s t , 1968), and has effects that may or may not be mediated via the pituitary Ce.g. appetite suppression), various tests of physiological function were performed on methallibure-treated deer. These diagnostic procedures in -volved a quantitative analysis of the enzymes, metabolic wastes, cholester-o l , phosphorus, calcium and proteins in the serum, and the determination of sodium, potassium and chloride in the urine. The extent to which methalli-bure interferes with thyroid function was investigated by measuring serum thyroxine and resin T^ uptake. The thyroids, adrenals, and pituitary glands of wild deer were also examined histologically and'compared to those of methallibure-treated deer. In the fi n a l stages of this investigation, an attempt was also made to measure serum FSH and ICSH in normal and methalli-bure--treated deer by radioimmunoassay (using commercially available kits designed to measure human FSH and LH), but this was unsuccessful. 21 B. Materials and Methods 1, Serum collection From July, 1969, to August, 1973, blood samples were collected at 3 to 4 week intervals from 6 to 16 male deer kept i n captivity at the U.B.C. wildlif e research unit. To f a c i l i t a t e blood collection, the deer were immobilized by intramuscular injections of succinyl choline chloride (Anec-tine; Burroughs-Wellcome) at a dose level of .05 to .06 mg/kg body weight. Deer have a.very narrow range of tolerance to this drug, and when respira-tory arrest occurred a r t i f i c i a l respiration was applied manually until nor-mal breathing was restored. Serum samples from the captive deer were col-lected between 1 and 5:00 p.m. Starting in November, 1971, blood samples were also obtained from wild deer shot at 2 month intervals on northern Vancouver Island. A l l serum samples were frozen and.stored at -20°C until the day of analysis. 2. Determination of serum testosterone by competitive-protein-binding The competitive-protein-binding (CPB) technique was f i r s t developed for thyroxin by Ekins (1960) and later applied to steroids by Murphy and co-workers (Murphy, 1967). Since then a number of CPB methods have been described for the assay of testosterone in human plasma (Vermeulen and Verdonck, 1970), and many investigators are currently using this technique. c The method used in this study is similar to that of Maeda et_ al_. (1969), but i t u t i l i z e s thin-layer chromatography and includes some modifications in procedure that were incorporated to accommodate the wide fluctuations in testosterone levels of deer serum. This technique is specific, sensitive, 22 and faster than most other methods because i t u t i l i z e s a single thin-layer-chromatography step. A complete description of this method, includ-ing a s t a t i s t i c a l analysis and a discussion of the results is presented in Appendix A. 3. Estimation of testicular volume Starting in July, 1971, testicular volumes were estimated at the time of blood collection by comparative palpation, using testicular models of known volume. An orchidometer comprised of 15 e l l i p t i c a l models with vol-umes of .5, 1, 2, 3, 4, 5, 6, 8, 10, 12, 15, 20, 25, 30 and 40 cm3 was fashioned from modelling clay (Figure 1). The testicles were palpated with one hand while model testicles were palpated with the other, and their sizes compared. The scrotum and epididymis interfere with measurement so that the exact size cannot be determined, but this technique is simpler, faster, and just as accurate as in situ measurement of the longitudinal and transverse axis of the testes with calipers (Prader, 1965). 4. Semen collection and evaluation From October, 1971, to August, 1973, semen was collected at monthly intervals from 12 to 15 captive deer. This was usually done immediately after blood collection, while the animals were s t i l l immobilized. An a-c pow-ered electroejaculator (A.H. Northcott, Guelph, Ont) with a porcine-ovine i. rectal probe was u t i l i z e d in a manner similar to that described by Bierschwal et^ al_. (1970) . Semen was collected in a 15 ml graduated centrifuge tube f i t t e d with a rubber stopper and seated into a 125 ml flask which served as a water bath to protect the sperm from temperature shock. During the col-23 FIGURE 1. An orchidometer; used f o r measuring t e s t i c u l a r volume i n s i t u , by com-p a r a t i v e p a l p a t i o n . 24 25 l e c t i o n procedure the probe was i n s e r t e d i n t o the deer's rectum and the c o l l e c t i o n tube was held over the penis. An assistant operating the e l e c t r o -e j a c u l a t o r applied i n t e r m i t t e n t impulses (2 seconds on, 5 seconds o f f ) of gradually i n c r e a s i n g voltage u n t i l a maximum of 3 v o l t s was reached. The s t i m u l i were repeated u n t i l e j a c u l a t i o n r e s u l t e d . Ten to 20 s t i m u l i were us u a l l y required f o r each e j a c u l a t i o n , the number depending more on repro-ductive condition than the degree o f immobilization. Erection of the penis and complete e j a c u l a t i o n was always achieved by deer i n f u l l reproductive condition. Since immobilization with the proper dose of s u c c i n y l - c h o l i n e chloride did not prevent erection or i n h i b i t the ejaculatory response, i t was the p r e f e r r e d method of handling, because i t avoided much of the trauma and i n j u r y associated with manual r e s t r a i n t . Immediately a f t e r c o l l e c t i o n the volume and appearance of the ejaculate was recorded, and the tube containing the semen was put i n t o a l a r g e r water bath kept at 37°C. A drop of semen was placed on a pre-warmed s l i d e and the gross m o t i l i t y of spermatozoa was observed under low power (100 X) of a microscope. M o t i l i t y was graded i n t o f i v e categories: " E x c e l l e n t " (very vigorous forward motion with extremely rapid waves and eddies), "Very Good" (vigorous progression with r a p i d l y forming waves and eddies), "Good" (pro-gressive movement of spermatozoa with slow-moving waves or eddies), " F a i r " ( o s c i l l a t o r y or rotatory movement with no waves or eddies), and "Poor" (only occasional labored o s c i l l a t o r y or weak rotatory movement of i n d i v i d u a l spermatozoa). The percentage o f l i v e spermatozoa was determined by making two smears (as a blood smear i s made) from a few drops of a 1:1 mixture of undiluted semen and eosin-nigrosin s t a i n kept at 37°C f o r 5 minutes 26 (Hancock, 1951). The smears were quickly air-dried, and later a count was made of the dead spermatozoa, which stained pink in relation to the un-stained " l i v e " cells outlined against the dark background of the nigrosin. These and other smears stained with giemsa were also used for the examina-tion of morphological abnormalities. Sperm concentration was estimated by diluting a small amount of semen with water and counting the spermatozoa on a hemocytometer. 5. Methallibure treatment arid hormone injections Methallibure (I.C.I. 33828), obtained as a pre-mix (AIMAX) from Ayerst Laboratories, Montreal, was mixed and pelleted with formulated feed (U.B.C. deer ration No. 36-57) to give a concentration of 88 mg methallibure per kg of feed. Starting in June, 1971, two adult deer were administered methalli-bure-treated feed at a dose level of 1.25 mg/kg body weight per day while two fawns received methallibure in their milk (one at a dose level of 1 mg/ kg per day and the other at a dose level of 1.25 mg/kg per day). These dose levels were arrived at by evaluating the results of experiments on domestic animals (mostly swine) and were considered sufficient to suppress gonado-trophs secretion, while keeping side-effects to a minimum. From June, 1971 to September, 1973, methallibure was administered for varying periods of time to a total of 14 deer during the course of a series of experiments which included the injection of gonadotrophic hormones, prolactin, and androgenic steroids. The experimental treatment for each deer is given in Tables vA to XXA of Appendix B. Commercially prepared gonadotrophic hormones admini-stered to methallibure-treated deer were: PMS (Gestyl, Organon, Inc., Montreal), HCG (Antuitrin - "S", Parke-Davis, Brockville, Ont.), and LTH (Prolactin, Ferring, Malmo, Sweden). The activity of PMS and HCG is ex-pressed in units of FSH and LH respectively, although PMS contains some LH-like activity. Purified preparations of FSH (NIH-P1) and LH (NIH-B 8) were donated by the National Institute of Arthritis and Metabolic Diseases, N.I.H., Bethesda, Maryland. The gonadotrophic hormones were reconstituted in s t e r i l e saline just prior to injection to ensure freshness. Solutions of testosterone (4-androsten-17 beta-ol-3-one), androstenedione (4-androsten-3-17-dione), dehydroepiandrosterone (5-androsten-3-beta-ol-17-one), and 19-nortestosterone (4-estren-17 beta-ol-3-one) (Schwarz/Mann, Orangeburg, N.Y.) were prepared by dissolving the powdered hormone in st e r i l e sesame o i l containing 10% benzyl alcohol. A l l hormones were administered by intra-muscular injection. The dose levels for a l l hormones except the purified preparations of FSH and LH were arrived at by consulting various publications and veterinary manuals (e.g. Merck) containing information about the normal physiological levels, secretion rates, and effective therapeutic doses for domestic animals. Insufficient amounts of pure FSH and LH were obtained to permit the administration of therapeutic doses. 6. Routine measurements and tests of physiological function Observations on the antler cycle were recorded routinely for a l l the captive deer. Antler growth was measured linearly with a ruler and expressed as the mean of the aggregate length of the two antlers. Body weight was recorded every 3 to 4 weeks, and daily records of food and water consumption were also kept. Twenty-four-hour urine samples were collected for a period 28 o f 7 weeks from a normal and a methallibure-treated deer kept i n metabolism cages equipped with u r i n e - c o l l e c t i n g devices. The volume and s p e c i f i c g r a v i t y of urine was recorded, and routine laboratory procedures were em-ployed to measure sodium, potassium, chloride and the osmolarity of urine. A Technicon autoanalyzer (SMA 12/60) was used to measure serum glutamic oxaloacetic transaminase (SGOT), l a c t i c dehydrogenase (LDH), a l k a l i n e phos-phatase, b i l i r u b i n , c r e a t i n i n e , blood urea nitrogen (BUN),cholesterol, i n -organic phosphorous, calcium, and albumin of normal and methallibure-treated deer. Total thyroxin (TT^) was determined by the method o f Murphy et a l . (1966) , and r e s i n t r i i o d o t h y r o n i n e uptake (Resin T^U) was measured by the methods of Hamolsky et_ a l . (1959) and Clark (1963). The " f r e e " thyroxin index (FT^) i s the product o f the TT^ and Resin T^U values. Two commercial-l y a v a i l a b l e radioimmunoassay k i t s (Bio-RIA, Montreal, P.Q.) used f o r mea-suring human FSH and LH were t r i e d on deer serum, using bovine LH (NIH-B8) as an a d d i t i o n a l standard i n the LH system. Eight serum samples assayed f o r FSH and 45 samples assayed f o r LH did not contain s i g n i f i c a n t amounts of FSH or LH a c t i v i t y . When the standard curves f o r bovine LH and human LH were compared i t was evident that the antibodies to human LH did not cross-react extensively with bovine LH i n t h i s system. It was concluded that these k i t s were unsuitable f o r measuring FSH and LH i n deer. The time, e f f o r t and costs involved i n developing a s u i t a b l e radioimmunoassay system p r o h i b i t e d further e f f o r t s i n t h i s d i r e c t i o n . 7. Histology of the adrenals, t h y r o i d , and reproductive organs S t a r t i n g i n November, 1971, the adrenals, t h y r o i d , testes and accessory sex glands of 21 adult males and f i v e male fawns were c o l l e c t e d at 2 9 approximately two month i n t e r v a l s from wild deer shot on northern Vancouver Island. A f t e r estimating t e s t i c u l a r volume with the orchidometer, the testes and e n t i r e reproductive t r a c t were dissected out and f i x e d i n 10% buffered formalin f o r 48 hours. The adrenals and thyroid glands were also d i s s e c t e d out and f i x e d i n the same manner. A se c t i o n o f f i x e d t i s s u e from the middle o f each gland was embedded i n p a r a f f i n , sectioned at 8 u and stained with haematoxylin and eosin. Stained sections from each specimen were examined under the l i g h t microscope and the seasonal changes i n h i s t o -l o g i c a l appearance were noted and recorded. Stained sections from the endocrine organs of three methallibure-treated deer were also examined and compared to the normal deer. Photomicrographs showing the h i s t o l o g i c a l appearance of these glands were taken with a Zeiss photomicroscope. 30 C. RESULTS 1. Seasonal variation in serum testosterone and testis volume of  wild deer Seasonal variations in serum testosterone and testis volume of wild deer are presented in Table I. The serum testosterone levels in these deer were minimal (i.e., less than 1 ng/ml) in March, May and July. Higher levels of serum testosterone were recorded in September, and by November the circulating levels reached 10 ng/ml in the adult males. By late January a decline in testosterone secretion was evident. In January, the lowest concentration (0.74 ng/ml) was recorded in a buck that had recently dropped his antlers. The serum of two male fawns sampled in November and one sampled in January contained between 1.5 and 3.5 ng testosterone per ml. The serum testosterone levels of two adult females sampled in November and two others sampled in March were below 1 n.g/ml. Variations in testicular volume in the adult males also reflected a seasonal pattern of testicular activity. There was a three to five-fold 3 increase in testis volume (6 to 25 cm ) from May to November, followed by a comparable reduction in size from January to March. 2. Seasonal variation in the histology of the testes of wild deer Seasonal variation in the histological appearance of the tubular and i n t e r s t i t i a l tissue of wild deer is illustrated in Figures 2, 3, 5, 6, 7 and 8 (Plates I and II). On March 28 the seminiferous tubules were small and the epithelium consisted of a basal layer of Sertoli cells and type A resting spermatogonia. 31 TABLE I SEASONAL VARIATION IN SERUM TESTOSTERONE CONCENTRATION AND TESTIS VOLUME OF WILD BLACK-TAILED DEER ON VANCOUVER ISLAND Animal Sex Approx. Testis Testosterone Number Age Volume (ng/ml) (yrs.) (cc) March 28-29 8 M 1 •8 8 .41 (no antler growth) 9 M 2 .8 7 10 M 1 .8 8 .39 11 M 5 .8 6 34 F 1 .8 .63 38 p* 2 .8 .74 May 24-25 26 M 2 .0 6 .28 (antlers budding) 27 M 2 .0 5 .29 28 M 3 .0 6 .62 July 23 12 M 3 .2 9 .28 (antlers in 13 M 2 .2 7 .35 velvet) 14 M 1 .2 6 .34 15 M 6 .2 15 September 11-12 18 M 4 .3 25 2 .48 (start of rut; 19 M 15 .55 velvet shedding) 1 .3 1 20 M 5 .3 18 .73 16 M .3 3 .34 November 16-30 2 M 1 .5 25 6 .15 (mid-rut; antlers polished) 4 M 5 .5 25 10 .82 5 M 3 .5 20 4 .26 6 M 6 .5 25 10 .44 1 M .5 8 3 .18 7 M •5 7 1 .57 12 F 3, .5 .51 15 F 4, .5 .14 32 Table I (cont'd) Animal Sex Approx. Testis Testosterone Number Age Volume (ng/ml) (yrs.) (cc) January 18-21 21 M 2.7 8 (end of rut; antlers dropping) 22 M 3.7 10 .74 25 M 2.7 12 4.80 23 M • .7 3 3.25 24 M .7 3 * Pregnant 33 PLATE I FIGURE 2. Low power photomicrograph of the testis of an adult wild deer (no. 9) in March. Spermatogenetic activity is minimal, and the i n t e r s t i t i a l cells (lower center) also appear inactive at this time of year. H. and E. X 240. FIGURE 3. Photomicrograph of the testis of an adult wild deer (no. 27) in May. Spermatogenetic activity is greater than in March. Many tubules contain primary spermato-cytes and spermatids. H. and E. x 240. FIGURE 4. Medium-power photomicrograph of an epididymal tubule of an adult wild deer (No. 27) in May, to show that sperma-tids are being transported from the seminiferous tubules into the epididymis. This activity was not observed i n March. H. and E. x 600. FIGURE 5, Photomicrograph of the testis of an adult wild deer (No. 12) in July. Spermatogenetic activity in most tu-bules i s less advanced than i n May. Many of the sper-matocytes and spermatids in these tubules are degenerat-ing (arrows). H. and E. x 240. 54 35 PLATE II FIGURE 6. Photomicrograph of the testis of an adult wild deer (No. 18) in September. Man> tubules show a l l stages of spermatogenesis, but ful l production has not yet been reached. The activity of the interstit ial cells (right centre) is also approaching a maximum, as evi-denced by their enlarged nuclei and expanded cytoplasm, which stains more intensely than in March, May and July. H. and E. x 240. FIGURE 7. Photomicrograph of the testis of an adult wild deer (No. 4) in November. The seminiferous tubules show a l l stages of spermatogenesis. Fully formed spermatozoa are being released into the lumen of the tubules, and resi-dual bodies are common (arrows). The interstit ial cells (upper right and left centre) are expanded, and many nu-clei are larger and stain less intensely than in March, May, July and September. The cytoplasm of some intersti-t ia l cells contains a vacuole where l ipid droplets have been dissolved away during preparation of the section. This represents the period of maximal activity. H. and E. x 240. FIGURE 8. Photomicrograph of the testis of an adult wild deer (No. 22) in January. Tubular involution is in progress. De-generating germinal cells are being extruded into the lumen of the tubules (arrows). H. and E. x 240. FIGURE 9. Photomicrograph of the testis of a methallibure-treated captive deer in November. Spermatogenesis is almost com-pletely suppressed, and the interstit ial tissue is also inactive. Many of the interstit ial cells have a fibroblas-t ic appearance; their nuclei are elongate or flattened. H . and E. x 240. 36 37 There was no evidence of spermatogenetic activity at this time (Figure 2). No sperm were present in the ductus deferens, although a few spermatids in the cap or acrosomal stage were found in the epididymis. The volume of i n t e r s t i t i a l tissue in March was minimal,and the cells were small and flattened. The small, elongated, nuclei were f i l l e d with dark-staining chromatin. On May 25 spermatogenetic activity was observed in nearly a l l tubules. This was characterized by the appearance of many type B spermatogonia and primary spermatocytes, and some secondary spermatocytes and spermatids (Figure 3). Some spermatids in the cap or acrosomal phase were observed in the epididymis (Figure 4), but no sperm were present in the ductus de-ferens. The volume of i n t e r s t i t i a l tissue had not increased significantly in May. The Leydig cells were small and compressed, with dense, oblong nuclei. On July 23 the spermatogenetic activity in three out of four specimens was less advanced than i t had been in May. No spermatids were observed in the seminiferous tubules of these deer (Figure 5). However, in specimen No. 15 (a six-year old) the tubules were enlarged, and some showed a l l stages of spermatogenesis. Spermatids in the cap or acrosomal stage were present, but no spermatids or sperm were found in the epididymis or ductus deferens. The i n t e r s t i t i a l tissue in July had changed l i t t l e from March and May. The Leydig cells were s t i l l small, with oblong, dark-staining nuclei. By September 11, a l l the seminiferous tubules were noticeably larger (approximately 1.3 X) and more active. A l l stages of spermatogenesis were observed, and spermiogenesis was often complete (Figure 7). Residual bodies were commonly seen in the lumen of most tubules. However, the epididymis 38 contained many (50-80%) immature spermatids, and no sperm were present in the ductus deferens of No. 19 and No. 20. No signs of spermatogenetic activity were observed in No. 16, a male fawn. In the adult males, the volume of i n t e r s t i t i a l tissue had doubled by September,owing to an increase i n cytoplasmic area of the Leydig c e l l s . The nuclei also were larger, more rounded, and stained less intensely. During the last two weeks of November the germinal epithelium was s t i l l active, but spermiogenesis was the dominant feature of the seminiferous tubules (Figure 7). The epididymal tubules and ductus deferens were us-ually f i l l e d with sperm. Some spermatogenetic activity (spermatid forma-tion) was evident in the seminiferous tubules of male fawns in November, but no mature sperm were found. The i n t e r s t i t i a l cells of the adult males reached their maximal size in November. They were two times larger than in March, and the cytoplasm stained more intensely. The large , spherical nuclei were usually eccentric in the c e l l . By mid-January, 40-60% of the tubules had ceased their spermatogenetic activity, and the tubular epithelium was in a state of regression (Figure 8). Many displaced germ cells were found free in the lumen, and the cyto-plasm of the Sertoli cells was diffuse and vacuolated. Sperm were s t i l l present in some of the epididymal tubules but the ductus deferens was empty. The i n t e r s t i t i a l tissue also appeared to be regressing at this time. A higher proportion of cells had oval or elongated nuclei and a reduced cy-toplasm. No sperm were found in the tubules of male fawns in January, and the Leydig cells were small and flattened. 39 3. Histology of the testes of methallibure-treated captive deer. In striking contrast to the testes of normal deer, the testes of a methallibure-treated deer in November were small and inactive. The semini-ferous tubules were reduced in size and the germinal epithelium consisted of a basal layer of Sertoli cells and type A resting spermatogonia (Figure 9). Only 5 to 10% of the tubules contained type B spermatogonia and p r i -mary spermatocytes. No spermatids or sperm were present. The i n t e r s t i t i a l tissue was shrunken and atrophic. The Leydig cells were small and flattened, with dense, elongate, nuclei. The testes of another methallibure-treated deer sacrificed i n mid-December contained spermatogonia, spermatocytes and some spermatids, but the maturation of spermatids did not proceed beyond the cap or acrosomal phase. Degenerating spermatids were common in the tubules and in the epi-didymis. There were no sperm in the epididymal tubules or ductus deferens. The i n t e r s t i t i a l tissue of this deer was also shrunken and appeared inactive. In mid-July the germinal epithelium of a methallibure-treated deer con-sisted of Sertoli cells and type A resting spermatogonia, and the i n t e r s t i -t i a l tissue appeared inactive. 4. Seasonal variation in the histology of the accessory sex glands  of wild deer and methallibure-treated captive deer. a. Seminal vesicles and ampullae. The paired seminal vesicles consist of two long, folded tubes which have the appearance of a compound gland because they are divided into lobes or lobules by septae of connective tissue and smooth muscle. They are situated on either side near the base of the bladder, dorsal and lateral to the ampullae. The ampullae are the enlarged terminal portions of the ductus 40 deferens. Seasonal changes in the histological appearance of the seminal vesicles of wild deer are shown in Figures 10 to 15 (Plates III and IV). In March the seminal vesicles were small and compact. The glandular epithelium was low columnar (average height, 20 u), and the cytoplasm stained faintly owing to the scarcity of secretory granules. The dark-staining nuclei were tightly packed together along the basal portion of the c e l l s . No secretory material was present in the lumen of the tubules and ducts (Figure 10). In May and July the glands were larger and showed signs of redevelop-ment (Figures 11 and 12). About 65% of the nuclei in May, and a l l of the nuclei in July were larger, stained less intensely, and were less crowded together, owing to tubular expansion and the extension of cytoplasm into the lumen. The glandular epithelium appeared to be undergoing a reorganiza-tion process which often resulted in the extrusion of cells with small dark-staining nuclei into the lumen of the tubule. In July a few tubules contained light-pink-staining, secretory material. By September, the glandular cells had become high columnar, and they were actively producing secretory material that was. extruded into the lumen (Figure 13). In November the seminal vesicles reached their maximal size, which was two times greater than in March. The glandular cells were high columnar (average height, 40 u), and the cytoplasm stained intensely, owing to an abundance of secretory granules (Figure 14). The expanded lumina of the tubules and ducts were f i l l e d with pink-staining secretory material. Some secretory activity was evident in the seminal vesicles of male fawns during November, although the glands were small and compact. PLATE III Low-power photomicrograph of the seminal vesicle of an adult wild deer (No. 9) i n March. Secretory droplets are absent. The glandular epithelium is low columnar, and there i s l i t t l e evidence of secretory activity. The tubules are shrunken, and the glandular cells are crowded together. H. and E. X 240. Photomicrograph of the seminal vesicle of an adult wild deer (No. 27) i n May. The tubules are slig h t l y expanded, and the glandular epithelium appears to be undergoing a process of reorganization. Many nuclei are larger, and stain less intensely than i n March. H. and E. X 240. Photomicrograph of the seminal vesicle of an adult wild deer (No. 15) in July. The tubules are slig h t l y expanded, and the height of the glandular epithelium is increasing, but there i s s t i l l not much evidence of secretory activity. H. and E. X 240. Photomicrograph of the seminal vesicle of an adult wild deer (No. 18) i n September. The tubules are expanded, and the glandular epithelium is high columnar. Some cells have actively begun to secrete droplets into the lumen of the tubules. H. and E. X 240. 42 43 PLATE IV FIGURE 14. Photomicrograph of the seminal vesicle of an adult wild deer (No. 4) in November. Expansion of the tubular tissue i s maximal, and nearly a l l of the glandular cells are actively secreting droplets which f i l l the lumen of the tubules. H. and E. X 240. FIGURE 15. Photomicrograph of the seminal vesicle of an adult wild deer (No. 22) in January, during the regressive phase. Secretory droplets can s t i l l be seen i n the lumen of the tubules, but the secretory activity of the glandular cells is less than in November. H. and E. X 240. FIGURE 16. Photomicrograph of the seminal vesicle of a methallibure-treated captive deer in November. The tubules are shrunken and there i s l i t t l e evidence of secretory activity. H. and E. X 240. FIGURE 17. Photomicrograph of the ampulla of an adult wild deer (No. 4) in November. The lumen of the ductus deferens contains spermatozoa (arrows). The glandular tissue (bottom) resem-bles the secretory tubules of the seminal vesicles. H. and E. X 240. 44 4 5 In January the seminal vesicles s t i l l appeared to be active, but signs of regression were evident (Figure 15). Epithelial c e l l heights were diminished (average height 30 u), and the cytoplasm stained less intensely. Less secretory material was present in the lumina of the tu-bules and ducts. The mucosal lining of the ampullae i s folded and gland-like (Figure 17). The smooth muscle surrounding the lumen also contained glandular tissue which resembled the seminal vesicles and showed similar seasonal variations in secretory activity. Sperm were present in the ampullae from September to January. No sperm were found in the ampullae of male fawns. The seminal vesicles of a methallibure-treated deer i n July were small and compressed. The cytoplasm of the glandular cells was clear and often vacuolated, and the nuclei stained intensely. No secretory material was found in the lumen of the tubules. The seminal vesicles of methallibure-treated deer sacrificed in November (Figure 16) and December were also small and inactive. The cytoplasm of the glandular cells stained faintly, and the nuclei were small and dense. There was no evidence of secretory a c t i v i -ty. The ampullae of a methallibure-treated deer i n November also appeared inactive. No secretory droplets were found in the tubules, and the ductus deferens was empty. b. Prostate The prostate gland in deer i s disseminate. The glandular tissue con-sists of simple branched tubules organized into lobules embedded within a stroma of smooth muscle and fibrous connective tissue surrounding the pelvic 46 urethra. The tubules open separately into the urethra along 5 to 6 cm of i t s length (Figure 18). Sections taken from the middle of the prostate gland at different times of the year showed only slight variations in histological appearance (Figures 20 to 24, Plates V and VI). The glandular epithelium was expanded and the cytoplasm stained more intensely in September and November than in March, May and July. The nuclei was larger and stained less intensely in the f a l l . In January the prostate showed some signs of regression. Most cells had clear cytoplasm, and the chromatin of many nuclei was becoming condensed. The prostate glands of methallibure-treated deer sacrificed in July, November, (Figure 25) and December were similar in histological appearance to the glands of normal deer in March. c. Bulbo-urethral glands The paired bulbo-urethral glands are ovoid, compound-tubular glands that are p a r t i a l l y embedded i n , and extent laterally from, the urethral muscle as i t curves around the pelvic outlet, A single duct from each gland enters the pelvic urethra. A seasonal pattern in the activity of the bulbo-urethral glands was not readily discernible because the size and histological appearance of glands collected at the same time of year was different in some individuals (Figures 26 to 31), Plates VII and VIII. In March the glands were compressed laterally and the tubules were small. Some pink-staining material was usually present in the lumen. The cytoplasm of the glandular epithelium was clear, and the tightly-packed nuclei stained intensely (Figure 26). PLATE V ' Low-power photomicrograph showing the glandular tissue of the prostate (bottom) and a duct which enters the pelvic urethra (top). H. and E. X 240. Medium-power photomicrograph of the prostate gland of an adult wild deer (No. 9) i n March. The nuclei are slightly smaller and stain more intensely than in May, July, Sep-tember, November and January. There i s l i t t l e evidence of secretory activity. H. and E. X 600. Photomicrograph of the prostate of an adult wild deer (No. 26) in May. Many nuclei are slightly larger and stain less intensely than in March, but the cytoplasm is s t i l l faint. H. and E. X 600. Photomicrograph of the prostate of an adult wild deer (No. 15) in July. The histological appearance i s similar to that of a prostate gland in May. H. and E. X 600. PLATE VI Photomicrograph of the prostate of an adult wild deer (No. 18) in September. The nuclear chromatin i s less condensed, and the cytoplasm stains more intensely than in March, May and July. H. and E. X 600. Photomicrograph of the prostate of an adult wild deer (No. 6) in November. The glandular cells are expanded, and appear most active at this time. Many of the nuclei are enlarged, and the cytoplasm stains more intensely than in March, May, July and January. H. and E. X 600. Photomicrograph of the prostate of an adult wild deer (No. 22) in January. The glandular cells appear less active than in September and November, but the chromatin i s not as condensed as in March. H. and E. X 600. Photomicrograph of the prostate of a methallibure-treated captive deer in November. The glandular cells are small and appear inactive compared to those of a normal deer at the same time of year. H. and E. X 600. 50 • <•* IT? if* -.rv • 51 PLATE VII FIGURE 26. Low-power photomicrograph of the bulbo-urethral gland of an adult wild deer (No. 9) i n March. At this time of year the glandular tissue i s shrunken, and the tightly-packed nuclei stain intensely. H. and E. X 240. FIGURE 27. Photomicrograph of the bulbo-urethral of an adult wild deer (No. 28) in May. Some of the nuclei are larger, and stain less intensely than in March, but there is not much evidence of secretory activity. H. and E. X 240. Photomicrograph of the bulbo-urethral of a six year-old wild deer (No. 15) in July. The glandular cells appear slightly active. The nuclei are larger and stain less intensely than in March and May, and in three younger specimens also collected in July. H. and E. X 240. Photomicrograph of the bulbo-urethral of an adult wild deer (No. 18) in September. The glandular epithelium is high cuboidal, and the cytoplasm stains intensely, but the nuclei are smaller than in Figure 28. Secretory material is present in the lumen of some tubules (arrows), while others are devoid of secretory material. These variations in histological appearance made i t d i f f i c u l t to discern a seasonal pattern. H. and E. X 240. FIGURE 28. FIGURE 29. 53 PLATE VIII FIGURE 30. Photomicrograph of the bulbo-urethral of an adult wild deer (No. 4) in November. The cytoplasm of the glandular c e l l s stains intensely, but very l i t t l e secretory material accumulates i n the lumen of the tubules, which suggests that this material may be exuded or ejected into the urethra. H. and E. X 240. FIGURE 31. Photomicrograph of the bulbo-urethral of an adult wild deer (No. 25) in January. Some of the glandular cells s t i l l appear active, but many tubules show signs of regres-sion, and cellular material i s being extruded into the lumen of some tubules (arrows). H. and E. X 240. FIGURE 32. Photomicrograph of the bulbo-urethral of a methallibure-treated captive deer in July. The glandular cells appear inactive, as evidenced by their condensed chromatin and reduced cytoplasm. H. and E. X 240. FIGURE 33. Photomicrograph of the bulbo-urethral of a methallibure-treated captive deer in November. The glandular tissue appears atrophic and disorganized. Cellular material is being sloughed o f f into the lumen of the tubules (arrows). There i s l i t t l e evidence of secretory activity. H. and E. X 240. 55 In May the glands of two deer were expanded and more rounded. In these specimens the nuclei were larger and the cytoplasm stained more i n -tensely thai; i n March. Small amounts of secretory material were present in the lumina (Figure 27). In the third specimen the glands were similar in appearance to those collected in March. In July the glands of three younger deer appeared inactive, whereas the glands of a six-year-old were enlarged, and appeared to be producing secretory material (Figure 28). The nuclei also were larger, and stained less intensely than in the other specimens. In September the glandular epithelium was expanded, and the cytoplasm stained more intensely than in March, May and July (Figure 29). Secretory material was present in the lumen of some tubules. In November the cytoplasm also stained intensely, but very l i t t l e se-cretory material accumulated in the lumen of the tubules in the adult males, whereas the tubules of male fawns were usually f i l l e d with secretory ma-t e r i a l . This suggests that the glands of the adult males may exude or eject their secretory material during the reproductive season. In January the glands were s t i l l active, although some tubules were regressing. The cytoplasm stained less intensely than in November,and cel-lular material was being sloughed off into the lumen of some tubules (Figure 30) . The bulbo-urethral glands of a methallibure-treated deer in July were compact and flattened. The low cuboidal cells had pink cytoplasm and small, dark-staining nuclei. No secretory material was present i n the lumen of the tubules (Figure 31). In November, the glands of a methallibure-treated deer were small and atrophic. The cytoplasm of most cells was clear, and there was no secretory material in the lumen of the tubules. 56 The glandular t i s s u e appeared disorganized and c e l l u l a r material was being sloughed o f f i n t o the lumina (Figure 33). In the December specimen, the glandular t i s s u e also appeared i n a c t i v e , although some tubules con-tained secretory material. 5. Histology of the t h y r o i d gland of wild deer and methallibure-treated  captive deer The h i s t o l o g i c a l appearance of the t h y r o i d gland of w i l d deer at d i f -ferent times o f the year i s i l l u s t r a t e d i n Figures 34 to 39 (Plates IX and X). In November, January, and March the f o l l i c u l a r c e l l s were low cuboidal with f l a t t e n e d , dark-staining n u c l e i . There was l i t t l e evidence of "vacuo-l i z a t i o n " or pinocytosis of c o l l o i d droplets from the c e n t r a l lumen of the t h y r o i d f o l l i c l e s . Thus, c o l l o i d storage was a c h a r a c t e r i s t i c feature of the gland i n winter. In May, J u l y and September the f o l l i c u l a r c e l l s were high cuboidal and the n u c l e i were l a r g e r , more round, and stained less i n t e n s e l y . C o l l o i d droplets were being a c t i v e l y "vacuolized" by cytoplasmic psuedopodia from the a p i c a l region. The pinocytosis of c o l l o i d was most prominent i n J u l y . This suggests that there i s an increase i n the breakdown of thyroglobulin with the consequent release of thyroid hormones during the summer. The th y r o i d gland of younger animals usually had a more uniform appear-ance than that of older deer. In an old buck the f o l l i c l e s v a r i e d consider-ably i n s i z e and appearance (Figure 36). Thus, the average diameter of the f o l l i c l e s i s not always a r e l i a b l e i n d i c a t o r of thyroid a c t i v i t y . These va r i a t i o n s i n s i z e and h i s t o l o g i c a l appearance of the t h y r o i d f o l l i c l e s might be r e l a t e d to s t r u c t u r a l changes that occurred during the l i f e t i m e of 57 PLATE IX FIGURE 34. Low-power photomicrograph of the thyroid of an adult wild deer (No. 11) in March. The f o l l i c u l a r cells are flattened-cuboidal. The nuclei are small and elongate, and the chromatin is condensed. There is l i t t l e evidence of colloid uptake from the lumen of the f o l l i c l e s . The gland appears least active at this time of year. H. and E. X 240. FIGURE 35. Photomicrograph of the thyroid of an adult wild deer (No, 20) in May, The gland appears much more active than in March, The f o l l i c u l a r cells are high cuboidal, and their nuclei are expanded and more rounded. Colloid i s actively being taken up by the f o l l i c u l a r c e l l s . H. and E. X 240. FIGURE 36. Photomicrograph of the thyroid of a six-year-old wild deer (No. 15) in July, showing extreme variations in f o l -l i c u l a r activity. Many f o l l i c u l a r cells are hyperactive, as evidenced by vacuolization or pinocytosis of colloid drop-lets from the lumen of the f o l l i c l e s , but other f o l l i c l e s appear inactive or dormant (bottom and upper l e f t ) . H. and E. X 240. FIGURE 37. Photomicrograph of the thyroid of an adult wild deer (No. 18) in September. The gland i s s t i l l active, but there is some evidence of a decline i n activity. The f o l l i c u l a r cells are decreasing in height, and there i s less vacuoli-zation of colloid, H. and E. X 240. 58 59 PLATE X FIGURE 38. Photomicrograph of the thyroid of an adult wild deer (No. 4) in November. The f o l l i c u l a r cells appear less active-than in September, but they are s t i l l more active than in January, H. and E, X 240. FIGURE 39. Photomicrograph of the thyroid of an adult wild deer (No. 22) in January. The f o l l i c u l a r cells are low cub-oidal, and most of the nuclei are oval or elongate. This suggests that the gland i s less active i n winter than in summer. H. and E. X 240. FIGURE 40. Photomicrograph of the thyroid of a methallibure-treated captive deer in July. Vacuolization of colloid by the f o l l i c u l a r cells indicates that methallibure did not have a pronounced depressing effect on thyroid activity in this deer. H. and E. X 240. FIGURE 41. Photomicrograph of the thyroid of a methallibure-treated captive deer i n November. The histological appearance was similar to that of a normal deer in late f a l l or winter. H. and E. X 240. 61 an individual. The thyroid glands of methallibure-treated deer sacrificed in July (Figure 40), November (Figure 41) and December were similar in histologi-cal appearance to those of normal deer collected at the same time of year. Seasonal variation in the histological appearance of the p a r a f o l l i -cular cells was not apparent in any of these sections. 6. Histology of the adrenal cortex of wild deer and methallibure-treated  captive deer. As in some other mammals, the adrenal cortex of the Columbian black-tailed deer i s composed of three zones: the zona glomerulosa, the zona fasciculata, and the zona reticularis. The boundary between the zona glomerulosa and zona fasciculata is f a i r l y distinct, whereas the border be-tween the zona fasciculata and zona reticularis is not. The approximate ratio of the zona glomerulosa to the zona fasciculata and the zona reticu-l a r i s combined, at different times of the year was: ZG : ZF + ZR January 20-30% : 70-80% March 20-30% : 70-80% May 25-35% : 65-75% July 30-40% : 60-70% September 25-35% : 65-75% November 25-40% : 60-75% The cytoplasm of the zona fasciculata cells stained more intensely in March, 62 and there was also a greater degree of vascularization at this time. The adrenals of methallibure-treated deer were similar in appearance to those of normal deer at the same time of year. 7. Seasonal variations in testis volume, serum testosterone, sperm  production and the antler cycle of captive deer exposed to natural  photoperiod and to continuous light. Seasonal changes in the serum testosterone levels of untreated cap-tive deer were similar to those of deer in the wild, although much higher concentrations (20 to 30 ng/ml) were found in some individuals, and the testosterone did not reach a minimal level until late February or March (Figures 42, 48 and 49, and Tables XA, XIA and XIIA, Appendix B). Some captive deer were extremely aggressive during the rut. They remained in rut longer and dropped their antlers two months later than deer in the f i e l d . In spite of this, antler growth in these deer was initi a t e d at the normal time (late April or early May), Throughout the antler growing period the testosterone levels in the normal captive deer were below 1 ng/ml. An increase in serum testosterone occurred just prior to or at the time of velvet shedding, in late August or early September. The timing of this event was remarkably precise, and i t frequently occurred during the same week each year. However, the length of time that the antlers were retained in the polished condition was much more variable. The antlers were usually cast several weeks after the testosterone level had dropped below 1 ng/ml. Compared to the control deer, the two bucks kept under continuous light showed an irregular pattern of reproductive activity and antler re-placement. Their cycles varied from 6 months to 14 months, and they were 63 KEY TO ABBREVIATIONS AND SYMBOLS USED IN FIGURES 42 to 57: 1) Antler Cycle: ad: antlers dropped; as: antlers stopped growing; av: antlers in velvet; ng: new growth pd: pedicles developing; ps: pedicles stopped growing; vs: velvet shedding 2) Experimental Treatments: 24/L: continuous illumination; A : androstenedione; DHEA: dehydroepiandrosterone; FSH : follicle-stimulating hormone; HCG : human chorionic gonadotrophin; LH : luteinizing hormone (ICSH); LTH : prolactin; MT : methallibure treatment; nT : 19-nortestosterone; PMS : pregnant mares' serum gonadotrophin; T : testosterone 64 FIGURE 42. Seasonal variation in serum testosterone, testis volume, sperm concentration and the antler cycle of No. U-26, a captive deer exposed to outdoor conditions. (For de-t a i l s see Table VA, Appendix B). Key to abbreviations and symbols used in Figures 42 to 57 are shown on page 63. TESTOSTERONE (ng/ml) Xf. i ^ j_ _ IO K> K> . LLILJJI \v • tlt'.l 111111^  lit"i|m<..i« I l l 1 - * iS T I I if' S i p T E S T I S V O L U M E T o o (cm3) "V " ' " 'T" "•r'",i,t"ijl',ilf 2 > S >l ll[ > 5 "riivrr'''^"i ,! SPERM (X lOyml) 66 FIGURE 43a. Seasonal variation in serum testosterone and the antler cycle of captive deer No. W-l during and after exposure to continuous light; temperature held constant at 25°C (for details see Table VIA, Ap-pendix B). a-68 FIGURE 43b. Seasonal v a r i a t i o n i n serum testosterone, t e s t i s volume, sperm concentration and the a n t l e r cycle of captive deer No. W-l before, during and a f t e r treatment with m e t h a l l i -bure, p r o l a c t i n , HCG and LH (for d e t a i l s see Table VIA, Appendix B). TESTOSTERONE (ng/ml) oo aba o j Z ! D i o i l l ! =P=?=fB=sTs: TESTIS VOLUME » (cn.3) =rrrfW!I 111 > 5 o o SPERM (X 106/ml) O z o 70 FIGURE 44a. Seasonal variation in serum testosterone and the antler cycle of captive deer No. W-3 during and after exposure to continuous light; temperature held constant at 25°C (for details see Table VIIA, Appendix B). TESTOSTERONE (ng/ml) •'3 f. 10 10 O 10 to 72 FIGURE 44b. Seasonal v a r i a t i o n i n serum testosterone, t e s t i s volume, sperm concentration and the a n t l e r cycle of captive deer No. W-3 before and during methallibure treatment (for d e t a i l s see Table VITA, Appendix B.). o TESTOSTERONE • . OB . 3 '..,..„! ... „.,„„ ,1,,.,^.,,., (ng/ml) •HAHII.I )V J nuk to to f i r 1 T I 2 >i o r if ~ P i -ta J • fit -*6 of A* 1 w 1 * T TESTIS VOLUME T (cm3) > r o § SPERM (X 106/ml) 74 FIGURE 45. Seasonal variation in serum testosterone, testis volume, sperm concentration and the antler cycle of captive deer No. Y-9 before, during and after methallibure treatment (for details see Table VIIIA, Appendix B). 76 FIGURE 46. Seasonal variation in serum testosterone, testis volume, sperm concentration and the antler cycle of captive deer No. Y-16 before, during and after treatment with methalli-bure, PMS and HCG (for details see Table XIA, Appendix B). 78 •9 FIGURE 47. Seasonal variation in serum testosterone and the antler cycle of No. Y-23, a captive deer exposed to natural photoperiod with a r t i f i c i a l light; temperature held con-stant at 25°C (for details see Table XA, Appendix B). 80 FIGURE 48. Seasonal variation in serum testosterone, testis volume, sperm concentration and the antler cycle of No. Z-l, a captive deer exposed to outdoor conditions (for details see Table XIA, Appendix B). TESTOSTERONE (ng/ml) •o SPERM (X 104/ml) o 82 FIGURE 49: Seasonal variation in serum testosterone, testis volume, sperm concentration and the antler cycle of No. Z-7, a captive deer exposed to outdoor conditions. (for details see Table XIIA, Appendix B). TESTOSTERONE (ng/ml) •° SPERM (X 106/ml) o Oo 84 F I G U R E 5 0 . Seasonal variation in serum testosterone, testis volume, sperm concentration and the antler cycle of captive deer No. B-1 before, during and after treatment with methalli-bure, prolactin and HCG (for details see Table X I T I A , Appendix B). 86 FIGURE 51. Seasonal variation in serum testosterone, testis volume, sperm concentration and the antler cycle of captive deer No. B-2 before, during and after methallibure treatment (for details see Table XIVA, Appendix B). 88 FIGURE 52. Seasonal variation in serum testosterone, testis volume, sperm concentration and the antler cycle of captive deer No. B-3 before, during and after treatment with methalli-bure, PMS and HCG (for details see Table XVA, Appendix B) TESTOSTERONE (ng/ml) q 00 safes M 1 I "TJ £ Is; > s flzl l i t = r r r f r r r « 222; l | , t n , ^ , . p.i „ ¥ u^j r T T E S T I S V O L U M E T ( c m 3 ) SPERM (X 104/ml) > j -O p i l t 90 FIGURE 53. Seasonal variation in serum testosterone, testis volume, sperm concentration and the antler cycle of captive deer No. C-l before, during and after treatment with methalli-bure, HCG and dehydroepiandrosterone (for details see Table XVIA, Appendix B). TESTOSTERONE (ng/ml) 92 FIGURE 54. Seasonal variation in serum testosterone, testis volume, sperm concentration and the antler cycle of captive deer No. C-2 before, during and after treatment with methalli-bure, HCG and androstenedione (for details see Table XVIIA Appendix B). 94 FIGURE 55. Seasonal variation in serum testosterone, testis volume, sperm concentration and the antler cycle of captive deer No. C-3 before, during and after treatment with methalli-bure, HCG,i9-nortestosterone, and testosterone (for de-t a i l s see Table XVIIIA, Appendix B). 95 24 22 20-fi 184 16 ~ 14 0 ) c w 12 10 o-i 30 i O X S 100 so -0 u.;,u. l|j Ji»u, SONDjJFMAMJ J ASONDI 96 FIGURE 56. Seasonal variation in serum testosterone, testis volume, sperm concentration and the antler cycle of captive deer No. C-5 before, during and after treatment with methalli bure and HCG (for details see Table XIXA, Appendix B). TESTOSTERONE 0 . 0 3 , 0 (ng/ml) " to o S 3 4k o I t 8-S It o .3 S • TESTIS o o VOLUME T Ui o O o (tn.3) mi rinriv i r t r ' r n " l l i " " O w — W " i " <-p"'TM"T A. • „ , , , , . r.-, j.-,,,., y.,- ,|n,., -r—-r""",iBB!i CO o S o O o O o O o •° SPERM (X l0 4 /ml) o o Jr 2 op 98 FIGURE 57. Seasonal v a r i a t i o n i n serum t e s t o s t e r o n e , t e s t i s volume, sperm co n c e n t r a t i o n and the a n t l e r c y c l e of c a p t i v e deer No. C-6 b e f o r e , during and a f t e r treatment w i t h m e t h a l l i -bure, PMS, HCG, FSH and t e s t o s t e r o n e ( f o r d e t a i l s see Table XXA, Appendix B). 100 out of phase with the seasons and each other (Figures 43 and 44a, and Tables VIA aid VIIA, Appendix B) The indoor controls that were kept for two years under a r t i f i c i a l light with a natural photoperiod cycled in a normal fashion (Figures 46 and 47, Tables IXA and XA, Appendix B). Seasonal variation in testicular volume of captive deer was similar to deer in the f i e l d , although much greater volumes (40 to 50 cm ) were recorded in some animals (Figures 48 and 49). Testicular size was maximal in October-November and minimal from March to July. Sperm production of the control deer was greatest during the breeding season (Figures 42, 48 and 49, Tables VA, XIA and XIIA, Appendix B). Sperm concentration varied between 100 x 10^/ml and 700 x 10^/ml from October to January, and then declined to a low level (0 to 30 x 10^/ml). A significant increase in the concentration of sperm was recorded in the spring, when the serum testosterone levels and testis volumes were decreasing or had already reached a low point. After this brief period of activity, the sperm con-centration declined and remained low throughout the summer months, although some captive deer produced sperm a l l year round. 8. The effect of methallibure treatment arid hormone injections on repro- duction and the antler cycle of captive deer. In the f i r s t series of these experiments (starting on June 1, 1971) methallibure was administered to two adult males at a dose level of 1.25 mg/kg per day. As a result, antler growth was retarded, the antlers did not mature, and the velvet was not shed (Tables VIIIA and IXA, Appendix B). Testicular redevelopment was also prevented in these deer. Both spermato-101 genesis and testosterone production were inhibited (Figures 45 and 46, and Tables VIIIA, VIIIB, IXA and IXB, Appendix B). Starting on November 2, 1971, the administration of 3 X 667 I.U. PMS per week for six weeks to one of these methallibure-treated deer pro-duced a rise in serum testosterone from .28 ng/ml to 20.98 ng/ml, and re-sulted in shedding of the velvet (Figure 46 and Table IXA, Appendix B). The PMS injections may also have initi a t e d the production of sperm, which appeared in the semen two months later (Table IXB, Appendix B) . The sub-sequent administration of 3 x 667 I.U. HCS/wk for six weeks did not appear to sustain testosterone production in this deer, because the serum testos-terone declined to a low level (.16 ng/ml), and the antlers were cast prior to the termination of the HCG injections. During the f i r s t year of experimentation methallibure was also admini-stered to two male fawns. Starting on August 10, 1971, a dose level of 1.25 mg/kg per day suppressed testicular development and growth of the pedi-cles, but at a lower dose level, complete suppression of pedicle growth was not achieved (Figures 53 and 55, and Tables XVIA, XVIB, and XVIIIA, Appen-dix B) . In the second series of experiments (1972) , methallibure treatment was i n i t i a t e d in early Ap r i l . As a result, antler growth was either com-pletely prevented or suppressed (Figure 58, Plate XI; Tables VIA and XIIIA to XXA, Appendix B). Of two methallibure-treated deer that received PMS injections (starting on June 1, 1972; 3 X 933 I.U./wk for 8 weeks) one (a mature adult) showed a temporary rise in serum testosterone and polished his antler buds (Figure 52 and Table XVA, Appendix B), while a yearling 102 animal on the same injection schedule did not show this response (Figure 57 and Table XXA, Appendix B). PMS did not stimulate antler growth in either of these deer. The subsequent administration of HCG (3.x 933 I.U./ wk for 3 weeks) caused a temporary increase i n the serum testosterone of both deer. Several months later the semen of these deer contained sperm of low quality and concentration (Figures 52 and 57, and Tables XVB and XXB, Appendix B). Both deer shed their partly grown antlers prematurely; one on September 19, 1972, and the other on December 5, 1972. Of two methallibure-treated deer that received prolactin (LTH) injec-tions (starting on June 1, 1972; 3 X 933 I.U./wk for 8 weeks), both showed an increase in serum testosterone (from .37 ng/ml to 2.20 ng/ml and .18 ng/ml to 3.01 ng/ml; Figures 43b and 50 and Tables VIA and XIIIA, Appendix B). Prolactin did not stimulate antler growth in either of these deer. The subsequent administration of HCG (3 x 933 I.U./wk for 3 weeks) to both deer resulted in shedding of the velvet 10 days later, but the testosterone levels were low by the end of the three week period. The partly grown antlers of these deer were also shed prematurely (on September 8, 1972, and October 4, 1972). Methallibure treatment was subsequently discontinued in these deer (on September 11, 1972, and October 5, 1972), and within three weeks their testosterone levels had increased sufficiently to induce them to rub their pedicles bare. For the remainder of the breeding season their testosterone levels followed a normal pattern, but high sperm counts were not recorded until February and March of 1973 (Tables VIB and XIIIB, Appendix B). The thin bony caps representing the polished extremities of their pedicles were shed several weeks after the serum testosterone had 103 declined to a low level (on January 26, 1973 and March 23, 1973). Of two methallibure-treated deer that received HCG injections (start-ing on June 1, 1972; 3 x 933 I.U./wk for 12 weeks), one showed a signi-ficant increase in antler growth whereas the other did not. Pedicle de-velopment in the latter animal had been retarded by methallibure treatment at an early age (Table XVIIIA, Appendix B). The HCG injections appeared to rejuvenate some development in this deer, but the amount of antler growth was not pronounced. However, the other deer grew antlers that reached a length of 25 cm (Figure 59, Plate XI; Table XIXA, Appendix B). These antlers were larger than those normally grown by yearling bucks of this species. HCG did not appear to stimulate testosterone secretion or sperm production in either of these deer (Figures 45 and 46, and Tables XVIIIA, XVIIIB, XIXA and XIXB, Appendix B). One of them (C-3) showed a slight increase in serum testosterone and polished his partly grown antlers in September. Mature sperm were found in the semen from October to January, indicating that the suppression of gonadotrophins was incomplete. Evidently a dose level of 1.25 mg/kg per day was insufficient in this instance, and so the amount of methallibure for this animal was subsequently increased to 1,50 mg/kg per day. By the end of the second year of experimentation i t became apparent that 1.25 mg/kg per day was the minimum effective dose of methallibure for most deer. At 1 mg/kg per day, only partial suppression of testicular func-tion was achieved (Figures 44b and 51, and Tables VILA, VIIB, XIVA and XIVB, Appendix B), 104 In April of 1973, a third series of experiments was started with five methallibure-treated deer to investigate further the effect of the gonadotrophins and also some androgenic steroids on antler growth. Start-ing on June 1, 1973, HCG was administered to two methallibure-treated deer (3 x 933 I.U./wk for 4 weeks), but neither one responded by growing antlers; nor did they show an increase in testosterone production (Tables XVIA and XVIIA, Appendix B). The subsequent administration of androstenedione (3 x 17 mg/wk for 3 weeks, and 3 x 33 mg/wk for an additional 2 weeks) to one deer, and dehydroepiandrosterone (3 x 33 mg/wk for 3 weeks, followed by 3 x 66 mg/wk for 2 weeks) to the other, did not produce a significant amount of antler growth in either deer. Starting on June 1, 1973, a low dose of LH (3 x 8 U./wk) was adminis-tered for 3 weeks to a methallibure-treated deer,but this did not stimulate antler growth (Table VIA, Appendix B); nor did i t produce an increase in serum testosterone. On July 4, 1973, methallibure treatment was discontin-ued in this deer. Eight weeks later, renewed antler growth was observed, and by the end of September, the antlers had added 12 cm to their length (Table VIA, Appendix B). On October 21, 1973, the velvet was shed, and the antlers were maintained in the hard, polished condition un t i l they were cast on February 22, 1974. The extensions on these antlers were thinner and quite distinct from the i n i t i a l parts (Figure 60, Plate XI). These antlers were also much smaller than those of a normal deer (Figure 61, Plate XI). Starting on June 1, 1973, a low dose of FSH ( 3 x 6 U./wk) was adminis-tered for 3 weeks to a methallibure-treated deer, but this did not stimulate 105 antler growth (Table XXA, Appendix B). The subsequent administration of low doses of testosterone to this animal ( 3 x 8 mg/wk for 3 weeks f o l -lowed by 3 x 16 mg/wk for 2 weeks) had no effect on antler growth, but the latter dose was sufficient to raise the serum testosterone level from .39 ng/ml to 1.03 ng/ml, and velvet shedding occurred one week later. Starting on June 1, 1973, 19-nortestosterone (a weakly androgenic, synthetic steroid with anabolic properties) was administered to a methalli-bure-treated deer ( 3 x 17 mg/wk for 3 weeks), but this treatment did not stimulate antler growth (Table XVIIIA, Appendix B). The subsequent ad-ministration of a moderate dose of testosterone to this animal (3 x 33 mg/wk for 2 weeks) raised the serum testosterone level from .19 ng/ml to 1.76 ng/ml, and resulted i n velvet shedding at the end of the 2 week per-iod. 9. The effect of methallibure treatment and hormone injections on other  aspects of physiological function. The immediate effects of methallibure treatment were loss of appetite, increased t h i r s t , and increased urine production. A temporary reduction in daily feed intake resulted in loss of body weight, but even when feed con-sumption returned to i t s former level, body weight gains were reduced. Caution had to be exercised when administering methallibure early in the spring, because even captive deer fed ad l i b . are in poor condition f o l -lowing the rut, and a reduction in feed intake at this time could result in death. The daily water intake and urine output of methallibure-treated deer increased two to three-fold, and did not return to normal until after 106 PLATE XI FIGURE 58. FIGURE 59. FIGURE 60. FIGURE 61. Photograph of a methallibure-treated captive deer (No. W-l on August 23, 1972), shewing the inhibitory effect that methallibure had on antler growth. Photograph of a methallibure-treated captive deer (No. C-5 on August 23, 1972) showing the amount of antler growth that occurred during the administration of HCG in June and July. Photograph showing the antlers of methallibure-treated captive deer No. W-l after they were cast on February 22, 1974. I n i t i a l l y , antler growth was disrupted by ad-ministering methallibure from April 9, 1973 to July 4, 1973. The terminal extensions represent the renewed growth that occurred in September, 1974, two months after the termination of methallibure treatment. Photograph showing the antlers of an untreated captive deer (No. C-l in December, 1974). The antlers grown by this animal during the previous year were only 6 cm long, owing to the suppressing effect of methallibure. Methal-libure treatment was discontinued on August 24, 1973, and in 1974, antler growth returned to normal. 107 1 0 8 treatment was discontinued. This diabetes insipidus-like condition was characterized by a significant decrease in the osmolarity Of urine, owing to a decline in the concentration of sodium, potassium, and chloride (Tables VD and VID, Appendix B). Since the ionic balance was not upset, i t appears that this condition may have been caused by the suppression of ADH. The blood profiles of normal and methallibure-treated captive deer are presented in the "C" series of Tables in Appendix B, and those of wild deer are presented in Table XXI, Appendix B. Some constituents in the serum of normal deer showed a seasonal pattern, although most did not. In the wild deer and in the untreated captive deer, alkaline phos-phatase was high throughout the antler growing period, and low from Sep-tember to March. Serum thyroxin was also higher during the summer months, although there was considerable variation between individuals. Some para-meters (e.g., calcium and phosphorus) were relatively constant throughout the year, while others (e.g., SGOT and LDH) varied considerably in d i f f e r -ent individuals at the same time of year. Thus, a seasonal pattern in these constituents was not discernible. Methallibure treatment did not cause a significant change in the circulating levels of any constituents other than alkaline phos-phatase. The concentration of this enzyme in the serum of a methallibure-treated deer was lower than normal during the antler growing season. An increase in alkaline phosphatase was recorded in the HCG-injected, methal-libure-treated deer that grew antlers (Table XIIC, Appendix B). Total thyroxin decreased slightly in some methallibure-treated deer (Tables VIIC § VTTIC ) but the free thyroxin index was s t i l l 109 within the range of normal values (Tables VC, IXC, XC and XXI, Appendix B). 110 D. DISCUSSION 1. The Sexual Cycle As part of an investigation concerned with the hormonal control of reproduction and antler growth, seasonal changes in the histology of the reproductive organs, testicular volume, serum testosterone, sperm concen-tration, and the antler cycle were studied in wild deer and in captive deer, some of which were treated with methallibure and various hormones. In the wild deer, testicular size and serum testosterone were maximal i n November and minimal in March, May and July. An increase i n serum tes-tosterone, accompanied by velvet shedding in late August or early September marked the beginning of the rutting season, and a decrease in testis size and testosterone production in January indicated i t s decline. The peak of the breeding season is in November, when most of the adult females be-come pregnant (Thomas, 1970). A microscopic examination of the reproductive organs, using cytologi-cal and histological c r i t e r i a of functional activity (e.g. epithelial c e l l height, number of secretory droplets, cytoplasmic and nuclear staining and morphology), revealed that a seasonal pattern of cyclic activity occurs in these organs. The activity of the testes and accessory sex glands was high in September and reached a maximum in November. By January, a decline in activity was evident, and in March, the least amount of activity was noted. The activity of the testes and seminal vesicles was greater i n May and July than in March. Similar changes in the activity of the reproductive organs have been found in other seasonally breeding cervids. However, few studies have included a description of the histological appearance of a l l the accessory I l l glands. Wislocki (1943, 1949) reported that in white-tailed deer, sea-sonal changes were not apparent in the glrndular tissue surrounding the urethra, which he regarded as representing the prostate and bulbo-urethral glands. Aughey (1969) described a disseminate prostate in the red deer stag. On a histochemical basis, she divided the prostate gland into cranial and caudal portions. The cranial part, which produced mucopro-tein and fat, showed only slight seasonal variation, whereas the caudal part, which produced glycolipid, was most active during the rut. Neither Wislocki nor Aughey were able to discern separate bulbo-urethral glands in white-tailed deer and red deer. The present investigation revealed that these glands are distinct i n the Columbian black-tailed deer. Per-haps the caudal prostatic tissue described by Aughey (1969) is homologous to the bulbo-urethral glands of black-tailed deer. Seasonal changes in the bulbo-urethral and prostate glands of black-tailed deer were less pronounced than in the testes and seminal vesicles., but they showed a similar cyclic pattern. Although a l l the reproductive organs may be androgen dependent, and their activity generally reflects the pattern of testosterone production, some appear to be affected more than others. The testes and seminal vesi-cles showed the most pronounced seasonal changes whereas the prostate changed the least. It may be that large amount of testosterone are not required for the function of some organs. The gonadotrophins and prolactin may play an important role in stimulating the activity of these glands. In the untreated captive deer testicular size, sperm production, and serum testosterone were maximal from October to December, and minimal during the summer months. Seasonal variations in 112 serum testosterone were similar to those of caribou and reindeer (Whitehead and McEwan, 1973), and reflected a pattern of testosterone production that i s comparable to the seasonal changes in testosterone content found in the testes of roe deer- (Short and Mann, 1966) and red deer (Lincoln, 1971). Sperm production during the breeding season was similar to that of white-tailed deer electroejaculated by Bierschwal et_ al_. (1970) and Lambiase (1972). The concentration of sperm and quality of the semen, when compared to that of domestic animals, was judged adequate to enable successful breed-ing between September and January (and much later in some individuals). The higher testosterone levels and extended period of rut observed in some deer was probably related to the captive condition. Among those fac-tors that may have contributed to this situation are a high plane of nutri-tion and abnormal social conditions. In regard to the latter, only limited social contact was permitted between certain males, and only one male used as a breeder was in direct contact with the females. The recurrence of estrus in those females that were not mated, and the presence of visual and olfactory stimuli from both sexes probably added to the sexual and social frustrations of the confined males. By preventing these males from esta-blishing their positions in a social hierarchy, each might consider himself to be dominant, because none suffered defeat in an aggressive encounter. This situation could have resulted in the high level of aggressiveness ob-served in some captive deer. This idea is supported by the observation that the aggressiveness of a docile male increased when he was moved from an isolated area and placed i n a pen adjacent to an extremely aggressive buck. 113 Lincoln e_t al_. (1972) showed that testosterone has a direct inductive effect on social aggressive behavior i n the red deer stag, but no studies on the effects of social stimuli or social interactions on testosterone production in deer have been reported. In addition to the effects of social stimuli provided by other males, the sexual stimuli released by seasonally polyestrous females coming into heat every 22 to 25 days from November until February or March (Cowan, 1956; West, 1968) might have induced some males to remain in rut longer than nor-mal. The breeding buck U-26, which was penned with several females from October to January, did not experience an extended period of rut. 2~*. The activity of the reproductive organs in the spring and i t s re- 1atioriship to antler growth. A histological examination of the reproductive organs of wild deer re-vealed that an increase in spermatogenetic activity of the testes and an increase in the cellular activity of the seminal vesicles occurred in May. In July the reproductive organs also showed signs of activity, but sperma-togenesis was less advanced than i t had been in May. This suggests that the testicular cycle of the Columbian black-tailed deer also has four phases, similar to those recognized by Markwald (1968) in mule deer. These phases are: (i) a period of maximal activity in the f a l l , ( i i ) regression in the winter, ( i i i ) a period of reduced activity in the spring, and (iv) redevelop-ment in late summer. This seasonal pattern was easily discernible in the wild deer, and i t was quite pronounced i n the captive deer. The captive deer showed a dramatic increase in sperm production in the spring, while the 114 testes were diminishing in size and the serum testosterone was declining or had already reached a low level. This brief period of spermatogenetic activity usually coincided with the i n i t i a t i o n of antler growth, and de-clined several months before the onset of gonadal redevelopment in late summer. Of the many investigators who studied the sexual cycles of various temperate species of deer, most considered the testes to be active only during the breeding season. Consequently, the activity of the testes at other times of the year has not always been thoroughly investigated. For example, Lambiase (1972) electroejaculated captive white-tailed deer b i -weekly from August through March (and found a high concentration of sperm 1 from October to February), but no sperm samples were collected during the spring, and the testes of wild deer were examined only during the breeding season. There are however, some reports indicating that increased testicu-lar activity in white-tailed deer does occur in the spring. Robinson, et_ al_. (1965) noted that the semmiferous tubules of white-tailed deer*were slightly more active during the antler growing period. Even Wislocki's (1943) pioneer-ing study on white-tailed deer contains some information which f i t s this pattern, but too few animals were sampled for him to reach this conclusion. Stieve's (1950) voluminous account of the seasonal changes in histological appearance of testicular tissue in roe deer contains some photomicrographs that clearly show an increase in spermatogenetic activity at the beginning of the antler growing period. A study of male reindeer (Meschaks and Nordkvist, 1962) revealed that renewed germinal activity in the testes, accompanied by a significant increase in urinary excretion of 17-keto-steroids occurs during the spring. The mule deer, which is a very close 115 relative of the Columbian black-tailed deer, also shows an increase in testicular activity in the spring (Markwald, 1968). On the basis of his-tological evidence, Markwald (1968) inferred that the activity of both che Leydig cells and the seminiferous tubules of mule deer increases in the spring. In the present study, there was no evidence to indicate that the Leydig cells of black-tailed deer secrete significant amounts of testos-terone at this time. In fact, i t seems lik e l y that f u l l reproductive func-tion and rutting behavior do not occur in the spring because not enough testosterone is produced. However, this does not preclude the poss i b i l i t y that small amounts of testosterone, or some other steroid, may be secreted at this time. The results of Meschaks and Nordkvist (1962) appear to support this, but their data makes no distinction between steroid metabo-li t e s of testicular and adrenal origin. The studies of Short and Mann (1966) and Lincoln (1971) revealed that the content of testosterone in the testes of roe deer and red deer increases gradually during the antler growing period. In addition, Lincoln's (1971) study showed that the concentration of testosterone rose sharply at the beginning of the antler growing period, and then declined to a lower level. The proposal that testicular stimulation occurs i n the spring is sup-ported further by the observation that captive male deer sometimes exhibit two periods of rut in a single year (e.g., reindeer; Whitehead and McEwan, 1973). In the instance cited above, the spring rut was characterized by a brief, but pronounced increase in plasma testosterone, accompanied by shed-ding of the velvet from the partly grown antlers, and rutting behavior. The antlers were cast in May, several weeks after a decline i n the testosterone 116 level, and new ones were subsequently grown during the remainder of the antler growing period. Although no attempt was made to explain this anomaly, i t is evident that gonadotrophin secretion must have occurred in the spring, and that in this particular instance the testes responded by producing enough testosterone to stop antler growth, and to induce velvet shedding and rutting behavior. The role that photoperiod might have played in stimulating gonadotrophin secretion in this instance bears some consi-deration . Increased spermatogenetic activity in the testes of deer during the spring suggests gonadotrophin release. In view of the respective roles of FSH and ICSH in stimulating spermatogenesis and testosterone production, i t i s tempting to speculate that FSH is secreted at this time. If both hormones are produced, then for some reason the testes do not respond f u l l y . The possible involvement of prolactin in this instance w i l l be considered later. Gonadotrophin secretion i s controlled by exogenous and endogenous factors, which stimulate or inhibit the pituitary and the appropriate areas of the hypothalamus (Ellendorf and Smidt, 1970). Testicular regression in late winter would "free" the hypothalamus and the pituitary from the nega-tive feedback effects of testosterone and possibly other steroids (Dillon, 1973, Ezrin, 1973) on FSH and ICSH secretion. If, at the same time, the appropriate stimuli are present, then gonadotrophin secretion should occur. In the f a l l , the photoperiod provides this stimulus (Goss, 1968b, 1969). However, the ratio of light to dark is the same in March as i t is in Septem-ber, and so some stimulation might also occur in the spring. Among the spring breeders, some species of birds respond b r i e f l y to photoperiodic stimulation in the f a l l (Van-Tienhoven, 1968), but^reproduction does not "\ % v» Si 117 usually occur at this time because gonadal activation is incomplete. It is inferred that the same situation could occur in f a l l breeders, only in reverse. At present, the photoperiodic control mechanism in f a l l breeders i s not well understood, so further speculation in this direction would be capricious. Nevertheless, the available information indicates that at least one of the gonadotrophins is released in the spring. It is suggested that this initiates antler growth, and in the process, the gonads are also reactivated. A gonadotrophin might stimulate antler growth directly, as suggested by Tachezy (1956) , or i t could stimulate the gonads to produce an antler growth stimulating hormone. Since castrates grow antlers, the latter alternative is unlikely, unless the adrenal cortex takes over this function when the testes have been removed. Bubenik (1966) has suggested that the adrenals induce secondary ossification of the antlers in reindeer and caribou, but there is l i t t l e evidence to support this. Additional evi-dence supporting the hypothesis that the gonadotrophins stimulate antler growth w i l l be considered in Section 5. 3. The effect of methallibure on the reproductive Organs. A histological examination of the reproductive organs of methallibure-treated deer revealed that methallibure prevents the increase in activity that normally occurs in the f a l l . The testes of methallibure-treated deer were small and inactive. Both spermatogenesis and the secretory activity of the Leydig cells were suppressed. In addition, the glandular activity of the accessory sex glands was reduced. These effects are not caused by a direct desensitization of the reproductive organs, but rather by hypothala-mic inhibition of pituitary gonadotrophin secretion. By suppressing the 118 production and release of FSH and ICSH, adequate stimulation of spermato-genesis and testosterone production is prevented. In the absence of the gonadotrophins and testosterone, normal function of the testes and acces-sory sex glands is impaired. In addition, rutting behavior and sexual activity do not occur. 4. Testosterone and the antler cycle Goss (1963, 1968a, 1970) reviewed this subject extensively, and a summary of this information was presented on Pages 14-16. The serum testosterone data from the present study support Wislocki's (1943) hypothesis that testosterone is responsible for the maturation and maintenance of antlers in the hard,functional condition. Low levels of testosterone were found throughout the antler growing period. An increase in serum testosterone occurred at the time of velvet shedding, and the antlers were maintained in the hard, functional condition until several weeks after the serum testosterone had declined to a low level. This re-lationship was substantiated further by observations on the deer whose seasonal cycles had been affected by continuous illumination or by methal-libure treatment. Aseasonal increases in the testosterone level of the deer under 24L resulted in a cessation of antler growth and premature shed-ding of the velvet (Figures 43a and 44a). On the other hand, when gonado-trophic stimulation of testosterone production was prevented by methallibure, the antlers did not mature and the velvet was not shed (Figure 45). Velvet shedding occurs when a deer is motivated to polish his antlers, and i t is accompanied by an increase in aggressive behavior. The level of aggressiveness, in turn, probably depends on the amount of testosterone 119 present (Lincoln, eJTal_., 1972). A serum testosterone level of 1.5 ng/ml was usually sufficient to i n i t i a t e velvet shedding, but the antlers were cleaned much more rapidly when the testosterone level was higher. Occasion-a l l y , testosterone production rose so sharply that the velvet was stripped before i t s blood supply had been completely shut off. When the testosterone level increased gradually the velvet dried up and the shedding process took longer. In the methallibure-treated deer, the velvet appeared to lose i t s v i a b i l i t y after several months, but i t was not shed unless an increase in serum testosterone, produced by hormonal replacement therapy (Figures 46, 55 and 57), or resulting from an insufficient dose of methallibure (Figures 44b and 51) was recorded. From these and other observations on intact and castrated deer (Wislocki et^ al_., 1947; Goss, 1963, 1970), i t appears that testosterone exerts several important effects on the antler cycle; (i) i t induces secon-dary ossification of the antler, and by accelerating the maturation process i t terminates antler growth; ( i i ) i t induces behavioral changes that result in velvet shedding; ( i i i ) the continued production of testosterone aids in the maintenance of osteoblasts and osteocytes, to ensure that the antlers are retained i n the hard, functional, condition throughout the breeding season; and (iv ) , the withdrawal of testosterone at the end of the breeding season permits the resorbtion of bone at the base of the antlers, and they are cast off. After velvet shedding, the antler gradually 'dies' back to-ward i t s base, but the presence of testosterone may be required to maintain a viable connection between the living tissue of the pedicle and the osteo-cytes in the base of the antler. 120 5. The hormonal regulation of antler growth. This study showed that methallibure, a non-steroidal inhibitor of gonadotrophin secretion, effectively suppresses antler growth. This find-ing supports Tachezy's (1956) suggestion that the antler growth stimulating hormone is a gonadotrophin. Instances of the gonadotrophins exerting a direct influence upon male secondary sex characteristics are rare. The effect of ICSH (LH) on the breeding plumage of the male weaver finch is the best known example of this. The pos s i b i l i t y that ICSH may be involved in stimulating antler growth w i l l be considered shortly. When this investigation was initiated, i t was uncertain whether methal-libure would have an effect on prolactin secretion. However, i t has been reported that methallibure does not suppress prolactin secretion in sows (Garbers and F i r s t , 1968) and rats (Ben-David, et_ a l . , 1971; Deis and Vermouth, 1973/74). These findings, and the fact that additional amounts of prolactin administered exogenously to methallibure-treated deer did not rejuvenate antler growth, appears to disprove Wislocki's (1947) hypothesis that the 'male counterpart of the lactogenic hormone* is responsible for stimulating antler growth. This leaves FSH and ICSH as possible candidates. The synthesis and release of both FSH and ICSH is suppressed by methal-libure (Parkes, 1963; Walpole, 1965; Brown and Fawke, 1972; Hemsworth, et^ a l . , 1968; Aoki and Massa, 1972; and Deis and Vermouth, 1973/74). Methalli-bure acts at the hypothalamic level (Malven, 1971), and appears to exert i t s effect by increasing the activity of enzymes (peptidases), which i n -activate the gonadotrophin releasing factors (Griffiths and Hooper, 1973). The renewal of spermatogenetic activity in the spring suggests that FSH may be involved in the i n i t i a t i o n of antler growth. Unfortunately, the 121 specific effects of FSH and ICSH on antler growth in deer could not be f u l l y investigated because adequate amounts of the purified preparations of these horr..ones were not obtained. Instead, HCG, which has ICSH-like activity, and PMS, which has both FSH- and ICSH-like activity were used. The i n a b i l i t y of PMS and HCG to exactly mimic the effects of pure FSH and ICSH probably limited the success of these experiments. A gradual de-crease in v i a b i l i t y of the velvet in methallibure-treated deer may also account in part for the poor growth response that was achieved with some hormone injections. Even when gonadotrophin production was allowed to return to normal by terminating methallibure treatment, antler growth did not resume until 8 weeks later (Figure 43b and Table VIA, Appendix B). Prolonged hormone therapy would not solve this problem because the effect-iveness of some hormones is reduced by antibody formation after one or two months. These d i f f i c u l t i e s might have been avoided by starting the hormone injections soon after the i n i t i a t i o n of methallibure treatment. However, at that time i t was considered essential to ensure that methallibure com-pletely suppressed antler growth in a l l cases. The apparent a b i l i t y of HCG to stimulate antler growth in one instance (Table XIXA, Appendix B) was not substantiated during subsequent tests, and no other hormones that were tried restored antler growth. Nevertheless, the results obtained with HCG in even this one instance might be significant, because every precaution was taken to ensure that antler growth was being suppressed by methallibure prior to the administration of HCG. However, HCG has LH-or ICSH-like activity, and i t was suggested earl i e r that other evidence appears to favor FSH as the antler growth stimulating hormone. 122 This apparent discrepancy w i l l be resolved when the seasonal patterns of FSH and ICSH secretion are known. With this knowledge, appropriate amounts of the pure hormones could be administered exogenously to methallibure-treated deer ,to duplicate the normal physiological levels. Information about the seasonal patterns of FSH and ICSH secretion would also be an im-portant adjunct to further experimentation with exogenous hormones on antler growth, because ICSH can affect the antlers indirectly via the testes. Jaczewski and Topinski (1970) found that HCG administered at the appropriate time could delay antler shedding, presumably by prolonging the production of testosterone. Thus, the gonadotrophins, when administered alone or in combination, may produce quite different effects at different times of the year, in accordance with the normal cyclic pattern of hormonal stimulation and response. The effect of gonadotrophic hormones on the testes of methallibure-treated deer provides the basis for additional dis-cussion on this subject in the next section. 6. The hormonal regulation of testicular function. Under certain conditions, and during the antler growing period in particular, the testes of methallibure-treated deer did not respond to the exogenous administration of HCG by secreting testosterone, whereas both PMS and prolactin were effective in this respect. PMS was capable of pro-ducing a sudden and dramatic increase i n testosterone secretion (Figure 52 and Table XVA, Appendix B), while the effect of prolactin was more gradual and less pronounced (Figures 43b and 50, and Tables VIA and XVA, Appendix B). After the administration of PMS, HCG was capable of stimulating tes-123 tosterone production (Figures 52 and 57, Tables XVA and XXA, Appendix B), whereas i t did not have this effect after treatment with prolactin (Fig-ures 43b ana 50, and Tables VIA and XIIIA^ Appendix B). Neither PMS nor HCG administered alone restored spermatogenesis, although some activity-was evident after sequential treatment (Tables IXB and XXB, Appendix B). When the inhibition of gonadotrophin secretion was removed in the f a l l by terminating methallibure treatment, testosterone production and sperma-togenesis resumed (Figures 51 and 52), but the deer that had previously received prolactin did not produce mature sperm until the following spring (Figures 43 and 50). The significance of some of these results i s not clear. However, i t appears that PMS, HCG and prolactin are a l l capable of stimulating the testes to secrete testosterone, although HCG may only be effective at cer-tain times of the year, and/or only after the testes have been primed with PMS (FSH). These results also showed that the testes do not always respond to gonadotrophic stimulation, particularly after treatment with prolactin. In mammals, the spermatogenetic process appears to be under the control of testosterone and FSH. ICSH is indirectly involved by virtue of i t s ca-pacity to stimulate testosterone production. Testosterone is required p r i -marily for the reduction-division process that results in the formation of secondary spermatocytes from primary spermatocytes. It may also promote the early steps of spermatid formation, but the late stages of spermatid maturation require FSH (Steinberger, 1971). The exact role of FSH is s t i l l uncertain, but i t i s believed to regulate Sertoli c e l l function (Lostroh, 1971; Steinberger, 1971). Although the effects of FSH on the testes are distinct from those of ICSH, each hormone enhances the effect of the other when administered together. This may explain why PMS, which has both FSH-124 and ICSH-like activity, was more effective in stimulating the testes of methallibure-treated deer than HCG, which only has ICSH-like activity. Nevertheless, HCG by i t s e l f was capable of producing a rise in testoster-one secretion after treatment with PMS. This raises the question as to whether the gonadotrophic hormones act in a consecutive manner or in syner-g i s t i c fashion. In the methallibure-treated deer, PMS might have primed the testes to respond to HCG, whereas prolactin appeared to have the re-verse effect. Prolactin also appeared to delay the recovery of spermato-genesis after methallibure treatment was discontinued. These findings suggest that the testicular response depends on the temporal sequence as well as the type of hormonal stimulation. In regard to spermatogenesis, this could mean that ICSH exerts i t s effect before FSH, because testoster-one is required for the completion of meiotic divisions, while FSH is necessary for the fi n a l stages of spermatid maturation. Without testoster-one there would be no spermatids for FSH to act on. In the deer however, i t is apparent that a high serum testosterone level is not required for sperm production. Thus, a small localized amount of testosterone and the presence of FSH might be sufficient to produce mature sperm. This idea is supported by the observation that some tropical species of deer produce mature sperm in abundance regardless of whether or not the velvet has been shed (Goss, 1963). In view of earlier statements, one could expect to find a low testosterone level in these deer when the antlers are in velvet. In hypophysectomized mammals, neither FSH, ICSH nor testosterone, alone or in combination, are capable of maintaining quantitatively the spermatogenetic process (Steinberger, 1971). Prolactin and growth hormone, at doses having no direct effect on the reproductive system, have been 125 shown to augment the effects of ICSH alone or a combination of FSH and ICSH (Hafiez, ejt al_., 1971; Steinberger, 1971). The effects of prolactin on the reproductive organs of the mammalian male include: (i) synergism with androgens on the accessory sex glands, ( i i ) increased androgen binding in the human prostate, ( i i i ) elevated cholesterol levels in the mouse test i s , and (iv) the stimulation of enzyme activity associated with andro-gen biosynthesis i n the rodent testis (Hafiez, et_ al_., 1970; Nicoll and Bern, 1971). On the other hand, prolactin is antigonadal (antigonadotrophic) in birds (Nicoll and Bern, 1971), and there is some evidence to suggest that i t has this effect i n mammals as well. Prolactin appears to antago-nize gonadotrophin secretion in rats (Ben-David, 1971), and i t inhibits copulatory activity in male rabbits (Nicoll and Bern, 1971). Although pro-lactin appears to stimulate increased androgen synthesis in methallibure-treated deer, this does not preclude the poss i b i l i t y that i t might also be antigonadotrophic. Although i t is not clear how prolactin might exert this effect, i t did appear to cause a refractory state in the deer t e s t i s . Fur-ther consideration of this and other p o s s i b i l i t i e s concerning the roles and interactions of FSH, ICSH and prolactin in regulating reproduction and the antler cycle of deer must await the results of additional experimentation in this f i e l d . At present, not enough information is available to permit the elaboration of a scheme that explains a l l aspects of the hormonal con-trol of -reproduction and antler growth, but an attempt w i l l be made to i n -tegrate and summarize this information at the end of Section V. 126 7. Seasonal v a r i a t i o n i n t h y r o i d a c t i v i t y . Seasonal changes i n the h i s t o l o g i c a l appearance of the th y r o i d gland suggest that i t i s more active i n summer than i n winter. A s i m i l a r h i s t o -l o g i c a l pattern of thyroid a c t i v i t y has been found i n roe deer (Pantic and S t o s i c , 1966) and i n red deer (Freundova, 1955; Pantic and S t o s i c , 1966). G r a f f l i n (1942) was unable to detect any s i g n i f i c a n t seasonal changes i n the thyroids of w h i t e - t a i l e d deer, but other i n v e s t i g a t o r s found evidence to suggest that the t h y r o i d glands of w h i t e - t a i l e d deer are also more ac-t i v e during the summer. Hoffman and Robinson (1966) reported that the thy-r o i d weights of w h i t e - t a i l e d deer were maximal i n J u l y and August and minimal i n January and February. Although t h y r o i d weight may not be a r e l i a b l e index of th y r o i d a c t i v i t y , S i l v e r et a l . (1969) found that the me-t a b o l i c rate of immature and adult w h i t e - t a i l e d deer was one and a h a l f times greater i n summer than i n winter. In the present study, v a r i a t i o n s i n the serum thyroxin and r e s i n uptake of wild deer revealed a seasonal cycle of thyro i d a c t i v i t y which corroborates the h i s t o l o g i c a l findings (Table XXI, Appendix B). Seasonal v a r i a t i o n s i n the serum thyroxin o f captive deer also showed t h i s trend (Table XC, Appendix B), but i n some i t was less evident than others (Tables VC and . IXC, Appendix B). Seasonal v a r i a t i o n i n th y r o i d a c t i v i t y was not apparent i n captive deer No. Y-16, an indoor control kept at constant temperature (Table V I I I C , Appendix B). Yoiisef and Luick (1971) found that the thyroxin s e c r e t i o n rate of reindeer d i d not increase i n response to cold during the winter period. Apparently the th y r o i d gland o f deer (and some other w i l d mammals as w e l l ) , does not respond to winter cold i n the same manner as that of some laboratory 127 and domestic animals (Yousef and Luick, 1971). Hoffman and Robinson (1966) postulated that the combination of cold and semi-starvation in winter leads to reduced thyroid activity in deer. These changes in thyroid ac-t i v i t y reflect a seasonal pattern of metabolism which could have evolved as an adaptation to store energy during the summer, and to u t i l i z e i t con-servatively i n winter. If this is true, then some deer (captive animals in particular) might not show a reduction i n thyroid activity in winter i f they are on a high plane of nutrition and protected from severe cold. Seal et_ al_. (1972) found that pregnant female white-tailed deer on a high plane of nutrition did not show a decrease in serum thyroxin in winter, whereas in the wild deer and in captive deer kept on a moderate diet, thy-roxin production was significantly lower during the winter. In the present study, a high plane of nutrition and protection from severe cold could ac-count for the relatively constant thyroxin levels of some captive deer. However, since male deer in captivity often reduce their feed intake volun-t a r i l y during the rut, this might cause a reduction in thyroid activity. The cyclic pattern of thyroid activity was most pronounced in captive deer No. Z-7, which also showed the greatest reduction in feed intake while in rut. 8. The adrenal glands Of wild deer and methallibure-treated captive deer. Few studies on the adrenal glands of deer have been reported. Accord-ing to Goss (1963), Wislocki was unable to detect any seasonal variations in the histology of the adrenals in male white-tailed deer. Hucin (1957) reported that an increase in the size and cellular activity of the zona glomerulosa occurs during the summer in red deer. The present study 128 revealed that the ratio of the zona glomerulosa to the zona fasciculata and zona reticularis was greatest in July and lowest in January and March. This could ire an that the zona glomerulosa, which secretes the mineralo-corticoids, i s more active during the summer, whereas the zona fasciculata and zona re t i c u l a r i s , which produce the glucocorticoids and androgenic corticoids, are more active in winter. Hughes and Mall (1958) attempted to investigate Selyes' General Adap-tation Syndrome in deer, and found that the amount of adrenal cortical t i s -sue i n adult female Columbian black-tailed deer was related to their overall condition. Since some stress factors, such as cold and semi-starvation are greatest in the winter, one might expect to find an increase in the activity of the zona fasciculata cells at this time. However, Hughes and Mall (1958) found that the size of the adrenals and the condition of individual deer varied considerably within each sample. Thus, i t was not possible for them to make comparisons between populations. Yousef et a l . (1971) found that the hydrocortisone secretion rate of reindeer was greater in the winter than in summer, and concluded that the adrenal cortex was involved in cold acclimatization. In,view of the multifunctional nature of the adrenal gland, and i t s response to various types of stress, any possible role that i t may play in regulating the antler cycle remains obscure. Histological examination of the adrenals of methallibure-treated deer did not provide any clues re-garding this possibility. 9. The effect Of methallibure on physiological function. In addition to the inhibition of gonadotrophin secretion, methallibure has other effects that may or may not be mediated via the pituitary. It 129 caused a temporary reduction i n the daily feed intake, and reduced body weight gains i n deer. Similar effects have also been noted i n rats (Benson and Zagni, 1965), hamsters (Stratman et_ al_., 1969), guinea pigs (Das and Benson, 1967), and swine (Stratman and Fi r s t , 1969; Stratman et^ a l . , 1969). These effects result from appetite suppression, and since methallibure acts at the hypothalamic level (Malven, 1971), i t could af-fect the appetite control centre of the hypothalamus. However, methalli-bure has also been shown to lower thyroid function (Tullock, et_ al_., 1963; Walpole, 1965), and hypothyroidism is known to reduce appetite and nutri-ent absorbtion from the gut (Leatham, 1953). Bourke et_ al_. (1970) found that methallibure has three actions on the thyroid-pituitary system in rats: (i) inhibition of the release of TSH from the pituitary; ( i i ) i n h i b i -tion of TSH synthesis (evident only at higher doses); and ( i i i ) a thyroid-blocking action (which also was observed only at the higher dose levels), with consequent pituitary stimulation via the thyroid-pituitary feedback mechanism. The lowest dose administered by Bourke et_ al_. (1970) was 8-10 mg/kg per day, and the highest dose level was 40-50 mg/kg per day. This is from 6 to 50 times greater than the dose administered to deer in the present study. A dose level of 1.25 mg/kg per day caused a slight reduc-tion i n serum thyroxin of some deer (Tables VIIC and VIIIC, Appendix B) but the total thyroxin and the free thyroxin index of a l l the methallibure-treated.deer were s t i l l well within the range of normal values (Tables VC, IXC, XC and XXI, Appendix B). In addition, there were no apparent dif-ferences in the histological appearance of thyroid glands from normal and methallibure treated deer (Figures 34 to 41). Thus, at the dose level administered, methallibure did not significantly alter thyroid function. 130 These findings suggest that appetite suppression by methallibure was probably not due to an effect on thyroid function, but rather to i t s effect on the appetite control centre of the hypothalamus. According to Stratman and Fi r s t (1969) this is also what occurs in swine. Concerning the relationship of the thyroid gland to antler growth, i t i s evident that methallibure does not inhibit antler growth by sup-pressing thyroid function. This finding supports Goss' (1963) suggestion that thyroxin (or T^) might influence antler growth in a permissive way, by acting as a synergist of the antler growth stimulating hormone, rather than by stimulating antler growth i t s e l f . Another effect of methallibure on deer was increased water consumption and increased urine output. Increased t h i r s t has been noted in other ani-mals (Call eJTal_., 1959) but the diuretic effect of methallibure has not been reported. An analysis of the urine from normal and methallibure-treated deer (Tables IVD and VD, Appendix B) suggests that this diabetes insipidus-like condition was probably caused by the suppression of ADH. Methallibure is known to inhibit oxytocin release (Benson and Zagni, 1965; Garbers and F i r s t , 1968), and since both oxytocin and ADH are produced in the hypothalamus, and their release sometimes occurs simultaneously in re-sponse to a common stimulus (Gorbman and Bern, 1962), i t is interesting to note that methallibure exerts an inhibitory effect on ADH as well. An investigation of the blood chemistry of normal and methallibure-treated deer showed that methallibure did not significantly alter the c i r -culating levels of any constituents other than alkaline phosphatase. This enzyme catalyzes the reaction that produces the inorganic phosphate neces-sary for the calcification of bone, and as such i t probably plays an 131 important role in the ossification of antlers. Molello et_ al_. (1963) found that the rapidly proliferating antler tissue of mule deer contains high alkaline phosphatase activity, and Graham et_ al_. (1962) reported a five-fold increase in the activity of this enzyme in the serum of male white-tailed deer during the antler growing period. Graham et^ al_. (1962) also found that the metabolic demands of antler growth did not s i g n i f i -cantly alter the blood levels of calcium, phosphorus, albumin and globulin. In the present investigation, the blood profiles of normal black-tailed deer substantiated these findings (Tables VC, VIIIC, IXC and XXI, Appendix B). Alkaline phosphatase was 2 to 10 times higher during the antler grow-ing period, whereas there were only slight seasonal variations in serum calcium, inorganic phosphate, albumin and total protein. Other studies have shown that deer are able to maintain relatively con-stant levels of calcium and phosphorus during the antler growing period be-cause the skeleton serves as a reservoir for these minerals. Meister (1956) Banks, ejt al_. (1963), and Hillman, et a l . (1973) found that increased re-sorbtion of the skeleton occurs during the period of antler growth. Taft e_t al_. (1956) reported that radiocalcium i n i t i a l l y deposited in the skeleton was later resorbed and u t i l i z e d in the ossification of antlers. The relationship of alkaline phosphatase to the antler growth stimula-ting hormone has not been established, but i t appears that methallibure, by suppressing the secretion of the antler growth stimulating hormone, also prevents the normal rise in alkaline phosphatase that occurs during the antler growing period. It i s interesting to note that the level of alkaline phosphatase in the methallibure-treated, HCG-injected deer that grew antlers (Table XIC, Appendix B) declined when methallibure treatment was applied, 1 3 2 and then increased during the administration of HCG in June. Thus, i f the antler growth stimulating hormone is z gonadotrophin, one of i t s effects might be to increase the production of alkaline phosphatase in the growing antler. How this i s brought about while the skeleton under-goes marked osteoporosis is not clear. Evidently the hormones that i n f l u -ence antler growth do not affect the tissues of the growing antler and the skeleton in the same manner. 133 III CYTOLOGY OF THE ANTERIOR PITUITARY A. Introduction Very l i t t l e is known about the cytology of the pars d i s t a l i s in deer. Purves and Bassett (1963) stained some deer pituitaries along with a number of other mammalian pituitaries, but no specific information on deer was reported. Pantic (1965) and Stosic and Pantic (1966) attempted to relate histological changes in the pituitaries of red deer to the antler cycle. Their interpretation was that the acidophils (which produce STH and LTH) and the thyrotrophic basophils probably stimulate antler growth, because these cells were more abundant and appeared to be more active during the antler growing period. The gonadotrophs were also reported to be active during the spring and summer, although their peak in activity was surmised to occur just prior to the breeding season. The pituitary showed the least signs of activity in winter, when small, tightly-packed chromophobes were predominant. Nicolls (1971) attempted to relate seasonal changes in the histology of the pituitary acidophils to the antler cycle of mule deer. He found that nuclear size and cytoplasmic area were maximal in the f a l l , which he believed indicated a high rate of hormone synthesis and a relatively low rate of secretion. The acidophil nuclear area was also significantly grea-ter in the spring than in winter and summer, but the cytoplasmic area was minimal at this time. Nicolls interpreted this to mean an increase in both the synthesis and release of hormones produced by the acidophils in spring. From these data, Nicolls implied that hormones secreted by the 134 acidophils induce antler shedding and new growth in the spring. He pos-tulated further that in f a l l , the acidophils respond to testosterone se-cretion by producing hormones which induce the resorption of minerals from the skeleton. These minerals were subsequently u t i l i z e d for the ossifica-tion of antlers. Freund (1955) found that the number of basophils and eosinophils (acidophils) i n the pituitaries of red deer also varied with the seasons. The percentage of acidophils was lowest in January (25%). A gradual in-crease occurred during the spring and summer (from 30% in April to 50% in August). By the end of September the acidophil population had declined to 35%. T h i s was followed by a second increase that reached 50% in November. It then declined steadily until January. The basophilic cells also showed two main periods of abundance; one in late July and the other in late Sep-tember. A third peak in the number of basophils was noted in the interval between late January and mid-April. . The interpretation of these data is complicated by the fact that there are two types of acidophils; one that secretes STH and the other LTH. Thus, each c e l l type may have i t s own seasonal cycle, and each could play a d i f -ferent role in relation to reproduction and the antler cycle. Both Freund's (1955) and Nicoll's (1971) data could be interpreted in the following manner: the acidophilic cells which increase in number during the spring and summer are somatotrophs, whereas the more abundant cells of the f a l l are lacto-trophs. Since growth in immature deer and body weight gains in a l l deer increase during the spring and summer, this might reflect an increase in the secretory activity of the somatotrophs. The lactotrophs however, might not increase their activity until f a l l , when the accessory sex glands are 135 most active. With regard to the basophilic c e l l s , the thyrotrophs, and perhaps the FSH cells as well, could be active in late July, whereas the ICSH cells might not become f u l l y active u n t i l September. The basophilic cells which increase in number during the spring might be responsible for an increase in gonadotrophic activity at this time. To examine these p o s s i b i l i t i e s , i t would be expedient to differentiate a l l the c e l l types of the pars d i s t a l i s . This was the main objective of the present study. In addition, i t was hoped that an examination of the pituitary glands from normal and methallibure-treated deer would reveal some clues about the activity of the gonadotrophic c e l l s . There are a few modern staining techniques that can differentiate be-tween the various kinds of acidophilic and basophilic c e l l s . However, the results achieved with these techniques have not been the same for a l l spe-cies. Their success also depends on the kinds of fixation, oxidation, and staining procedures u t i l i z e d . Variations in staining procedure have resulted in much confusion in the past. Nevertheless, the application of several techniques has led to the identification of four to six c e l l types in the adenohypophyses of a number of vertebrate species. The main c r i t e r i a used are: morphological characteristics, t i n c t o r i a l a f f i n i t i e s , and the histo-chemical reactions of each c e l l type. In recent years the greatest success has been achieved with two histochemical techniques: the acid permanganate-Alcian blue - PAS - orange G (OX-AB-PAS-OG) technique of Herlant (1960), and the acid permanganate-aldehyde thionin- PAS - orange G (OX-AT-PAS-OG) technique of Paget (1959) and Paget and Eccleston (1960). In the present study, these and many other staining techniques were tried. Tentative identification of the various c e l l types was made by 136 comparing them with the c e l l types in other mammals, including a few ungulate species. An attempt was made to identify a l l the c e l l types of the anterior pituitary, and to consider each c e l l type in relation to the physiological changes associated with i t s function. In addition, the pituitaries of methallibure-treated deer were examined and compared to those of normal deer. 137 B. Materials and Methods Starting i n November, 1971, a total of 25 pituitaries were collected at approximately two month intervals from wild deer shot on Vancouver Island (see Table I, p.31 ). The glands were removed within 20 minutes post mortem and halved in the mid-sagittal plane. One half was fixed in freshly prepared Helly's f l u i d (Zenker's stock with 10% formalin), and the other half was fixed in 10% buffered formation. After 20 hours fixation, the-glands were washed for 20 hours in d i s t i l l e d water and then stored in 70% ethanol. Prior to embedding they were put into a saturated solution of iodine i n 70% ethanol for 5 hours to remove mercury deposits. Embed-ding in paraffin was performed in the usual manner. The embedded tissue was sectioned s e r i a l l y in the horizontal plane at 6 y, and every 40th section was mounted on albuminized slides. The mounted slides were a i r -dried for a month or more, and the day before staining they were.placed in a formalin-fumed oven at 56°C for 1 hour, then air-dried again at 37°C overnight. This procedure was found to improve the adherence of sections to their slides during the staining procedure (Heath, 1965). The staining procedures for AB-PAS-OG, modified after Herlant (1960) and AT-PAS-OG, modified after Paget (1959) and Paget and Eccleston (1960) are presented in Appendix A. Other staining procedures that were tried, with variable success, were: Cleveland and Wolfe's stain (1932), Herlant's tetrachrome (1960), Paget and Eccleston's OX-AT-Luxol Fast Blue-PAS (1960): Kerr's Luxol Fast Blue-PAS-OG (1965), PAS-Haem.-OG (Herlant, 1956), 138 Adams and Swettenham's Performic Acid-AB-PAS-OG (1958), Racadot's (1962) modification of Herlant's tetrac.I-.rome, Dawson and Friedgood's . Azan (1938), Crossmon's (1937) modification of Mallory's trichrome, Azo-carmine, PAS-OG, and Erythrosin-OG (Pearse, 1950). The OX-AB-PAS-OG technique was tried with various Schiffs reagents, but L i l l i e s ' (1965) cold Schiffs gave the best results. The OX-AB-PAS-OG procedure was also tried at different pH's ranging from 0.2 to 3.0. Differentiation at pH 3.0 was the best. The histochemical reactions of the AB-PAS-OG tecnique, and theoretical aspects of many of the above staining procedures have been reviewed by el-Hakeem (1971). He found that AB also gave better results at pH 3.0. The OX-AB-PAS-OG and OX-AT-PAS-OG techniques, as applied in the pre-sent study, gave similar results. Since the greatest success was achieved with these two histochemical procedures, the descriptions which follow are based on them. 139 C. Results Cytology of the Pars Anterior at Different Times of the Year in  Normal and Methallibure-treated Deer The morphology of the deer hypophysis (Figure 62) is similar to the bovine hypophysis described by Heath (1970). The zonation of the pars anterior into serous (acidophilic) and mucoid (basophilic) zones i s based on the histochemical reactions of the c e l l types which pre-dominate these areas. The AB-PAS-OG and AT-PAS-OG techniques demonstrated six c e l l types in the pars di s t a l i s and one ce l l type in the pars inter-media. The functional system of nomenclature recommended by Purves 1961, 1966; Herlant, 1964; and van Oordt, 1965 has been applied to avoid the confusion created by earlier systems, but i t must be emphasized that the identification of each c e l l type is only tentative. Type I: (somatotroph; STH cell) The secretory granules of these serous or acidophilic cells stained yellow with orange G (Figures 63 and 64). The staining reaction with various components in the combined histochemical procedures was: AB-ve, AT-ve, PAS-ve, and OG + ve. These cells are round or oval with distinct boundaries. They are medium-sized (10-12 y in diameter), and have a dis-tinct central or sli g h t l y eccentric nucleus. The nucleus usually contains one prominent nucleolus and several smaller bodies of chromatin along the inner surface of the nuclear membrane. The cytoplasm appears homogeneous owing to the even distribution of very fine granules. These cells are found mainly in the serous zone which extends laterally and ventrally 140 FIGURE 62. Diagram of a median section of the deer hypophysis i l l u s t r a t i n g the zonation of acidophilic and basophilic c e l l s , and the plane of sectioning (dotted li n e ) . ME, median eminence; NS, neural stalk; PD, pars d i s t a l i s ; PI, pars intermedia; PN, pars nervosa; PT, pars tuber-a l i s ; RL, residual lumen; AMZ, anterior mucoid (baso-philic) zone; CMZ, central mucoid zone; SZ, serous (acidophilic) zone. 141 142 FIGURE 63. Photomicrograph of the pars d i s t a l i s of an adult male deer (No. 26) i n May, showing five of six c e l l types; Type I yellow, Type II orange, Type III red-orange-f o l l i c u l a r , Type V purple and Type VI pink or magenta AT-PAS-OG. Kodacolor II. X 480. FIGURE 64. The pars d i s t a l i s of a male fawn (No. 1) in November, showing five of six c e l l types: Type I yellow, Type II orange, Type IV blue, Type V purple and Type VI pink or magenta. AT-PAS-OG. Kodacolor II. X 1200 ( o i l ) . 143 144 FIGURE 65. The pars di s t a l i s of an adult male (No. 9) i n March, 'showing chromophobic cells and Type II orange ce l l s . Some of the chromophobes are Type III red-orange cells which contain dark red-orange deposits. AB-PAS-OG. Agfachrome. X 1200 ( o i l ) . FIGURE 66. The pars d i s t a l i s of an adult male (No. 9) in March, showing chromophobes and partly granulated Type VI magenta c e l l s , one with a large crennelated nucleus. AB-PAS-OG. Agfachrome. X 1200 ( o i l ) . 146 FIGURE 67. The pars di s t a l i s of a methallibure-treated deer i n December, showing Type I yellow c e l l s , Type II orange c e l l s , and many chromophobic basophils. AT-PAS-OG. Kodacolor II. X 480. FIGURE 68. The pars dist a l i s of an adult male (No. 26) in May, showing five of six c e l l types, and the f o l l i c u l a r arrangement of the Type III red-orange cells (lower l e f t ) . AT-PAS-OG. Kodacolor II. X 480. 147 148 FIGURE 69. A f o l l i c l e of Type III red-orange cells from the pars dis t a l i s of an adult male (No. 26) in May, showing the characteristic red-orange basal staining and the pur-plish-grey apical staining of these c e l l s . AT-PAS-OG. Kodacolor II. X 1200 ( o i l ) . FIGURE 70. The pars d i s t a l i s of a mehtallibure-treated deer in July, showing f o l l i c l e s of Type III red-orange ce l l s containing purple colloid. Note also the chromophobic nature of most of the basophilic c e l l s . AT-PAS-OG. Kodacolor II. X 480. 1 5 0 FIGURE 71. The pars d i s t a l i s of an adult male (No. 18) in September, showing many f u l l y granulated Type V purple c e l l s ; some f u l l y granulated, partly granulated and degranulated Type VI pink or magenta c e l l s ; and a partly granulated Type IV blue c e l l . AT-PAS-OG. Kodacolor II. X 480. FIGURE 72. The pars di s t a l i s of an adult male (No. 18) in September, showing the distinctive t i n c t o r i a l properties of the Type IV blue c e l l , compared to the Type V purple c e l l s . AT-PAS-OG. Kodacolor II. X 1200 ( o i l ) . 152 FIGURE 73. The pars dist a l i s of an adult male (No. 6) i n November, showing partly granulated Type V purple c e l l s . Note also the partly granulated Type I yellow cells and a partly granulated Type IV blue c e l l . AB-PAS-OG. Agfachrome. X 1200 ( o i l ) . FIGURE 74. The pars dist a l i s of a male fawn (No. 1) in November, showing partly granulated Type V purple c e l l s , a partly granulated Type IV blue c e l l , a partly granulated Type I yellow c e l l , and three f u l l y granulated Type II orange cel l s . AT-PAS-OG. Kodacolor II. X 1200 ( o i l ) . 1 154 FIGURE 75. The pars d i s t a l i s of an adult male (No. 26) in May, showing f u l l y granulated (F), partly granulated (P), and degranulated (D) Type VI pink cel l s . Note also Type I yellow c e l l s , Type III red-orange c e l l s , and a partly granulated Type V purple c e l l . AT-PAS-OG. Kodacolor II. X 1200 ( o i l ) . FIGURE 76. The pars d i s t a l i s of an adult male (No. 12) in July, showing Type I yellow c e l l s , Type III red-orange c e l l s , f u l l y andepartly granulated Type V purple c e l l s , and a f u l l y granulated Type VI pink or magenta c e l l . AB-PAS-OG. Kodachrome II. X 1200 ( o i l ) . 155 156 FIGURE 77. The pars d i s t a l i s of a methallibure-treated deer in July, showing a cluster of mostly chromophobic Type V purple c e l l s . AB-PAS-OG. Kodachrome II. X 1200 ( o i l ) . FIGURE 78. The pars dist a l i s of a methallibure-treated deer in July, showing a f o l l i c l e of Type III red-orange c e l l s , Type I acidophils, and several degranulated Type V purple c e l l s . AB-PAS-OG. Kodachrome II. X 1200 ( o i l ) . NOTE: The blue-staining deposits in Figures 71 and 72 are artifacts (mercury residue). 158 FIGURE 79. The pars d i s t a l i s of an adult male (No. 26) in May, showing a f u l l y granulated Type VI pink or magenta c e l l . Also shown are Type I yellow, Type II orange and Type III red-orange cel l s . AB-PAS-OG Agfachrome. X 1200 ( o i l ) . FIGURE 80. The pars di s t a l i s of an adult male (No. 18) in September. showing f u l l y granulated (F), partly granulated (P), and degranulated (D) Type VI pink or magenta c e l l s . Also shown are Type I yellow c e l l s , Type III red-orange cells and f u l l y granulated Type V purple c e l l s . AT-PAS-OG. Agfachrome. X 1200 ( o i l ) . vi D 160 FIGURE 81. Adjacent areas of the neurohypophysis and pars intermedia of an adult male (No. 18) i n September, showing the blue-staining neurosecretory material of the pars nervosa and the Type VII red-purple (MSH) cells of the pars intermedia. AT-PAS-OG. Kodacolor II. X 1200 ( o i l ) . FIGURE 82. The pars tuberalis of a methallibure-treated deer in De-cember, showing part of a large f o l l i c l e f i l l e d with purple colloid. Similar structures were also found in normal deer. AB-PAS-OG. Agfachrome. X 1200 ( o i l ) . 162 throughout the pars anterior, but they were also found scattered throughout the central jasophilic zone. They were rarely seen i n the antero-medial area of the basophilic zone. In the adult male, the type I cells constitu-ted about 20-30% of a l l c e l l s . They were most common (about 30-40%) in the pituitaries of male fawns. Seasonal variations in the histological appearance of these cells were not as pronounced as in some other ce l l types. The cytoplasm was f a i r l y uniform in i t s staining characteristics at different times of the year, but the nuclei were smaller and the chro-matin stained more intensely in November, January and March. The histolo-gical appearance of these cells in the methallibure-treated deer was no different from that of normal deer (Figures 67, 77 and 78). Type II: (lactotroph; LTH cell) The secretory granules of these cells stained orange or light brown (Figures 63 and 64). They were AB-ve, AT-ve, weakly PAS + ve, and OG + ve in the combined staining procedures. When stained with PAS-OG or erythro-sin-OG they are easily distinguishable from the type I acidophilic c e l l s , which do not react with PAS or erythrosin. These cells are also round or oval shaped, but they were slightly more irregular than type I c e l l s . The c e l l boundaries are distinct. These cells are slightly larger in size (12-14 u in diameter) than type I c e l l s , but clusters of small specimens sometimes occurred. The nucleus is prominent and often eccentric. In male fawns the nuclei of these cells were usually more irregular and stained dark orange or red-orange owing to an aggregation of nuclear material (Figure 64). The cytoplasm of these cells appeared homogeneous because of 163 the evenly distributed fine granules. In the adult male the type II cells constituted about 10-15% of a l l c e l l s . In March and May these cells were relatively small and irregular in shape. The nuclei stained intensely. In July their appearance was similar, except that some nuclei stained less intensely. In September and November these cells were larger and more rounded. The nuclei also were larger and stained less intensely, so that several distinct nucleoli were usually v i s i b l e . In January, smaller cells with dark-staining nuclei became more common. In the methallibure-treated deer these cells were similar i n appearance to those of normal deer (Figure 67). Type III: (corticotropin; ACTH cell) These cells stained red-orange or brick-red because they were AB-ve, AT-ve, PAS + ve and weakly OG +ve (Figure 69). They have an oblong or irregular shape, and were frequently found i n f o l l i c l e - l i k e clusters, with long cytoplasmic processes extending toward the centre of each f o l l i c l e . In the winter, some f o l l i c l e s contained purple colloid. The f o l l i c u l a r cells are large cells (12 x 18 u) with round or oval, sl i g h t l y eccentric nuclei. There was usually a prominent nucleolus and several smaller bodies of chromatin. The cytoplasm characteristically stained red-orange along the basal portion of the c e l l , while the apical region was blue-grey or chromophobic. The cytoplasmic granules are very fine. The type III cells were found throughout the pars anterior, but they were more common in the ventral and lateral regions of the acidophilic zone, and scarce i n the antero-medial area. In the adult male they comprised about 20-30% of a l l c e l l s . The red-orange f o l l i c u l a r cells stained most intensely in May and July. In September they stained less intensely, and by November they were 164 almost chromophobic. In January and March, a l l the type III cells were chromophobic, but they could s t i l l be identified by their morphology and characteristic grouping. The accumulation of purple colloid at the centre of each f o l l i c l e - l i k e cluster was also a distinguishing feature of these cells in November, January and March. In March, dark red-orange deposits were sometimes seen in the periphery of these cells (Figure 65). These cells appeared normal in the methallibure-treated deer (Figure 78), although some f o l l i c l e s contained purple colloid in July (Figure 70). Type IV: (thyrotroph; TSH cell) These are basophilic, mucoid cells that stain blue with AB-PAS-OG and AT-PAS-OG. They are AB + ve, AT + ve, PAS - ve and OG-ve i n the com-bined histochemical reaction (Figures 64, 71, 72 and 74). They have a triangular or polygonal shape, and the c e l l boundaries are usually distinct. These are medium-sized (12 x 15 y) cells with a distinct eccentric nucleus containing one or two large nucleoli. The cytoplasm is clear, with medium-sized granules or droplets distributed throughout. The type IV cells were sparsely distributed, but they were found mainly in the mid-central and lateral regions of the .basophilic zone. In the adult male they comprised about 1-2% of a l l c e l l s , and in the male fawns they were almost as scarce. Owing to their scarcity, no estimates of seasonal activity were made on these c e l l s . The type IV cells of a methallibure-treated deer were similar in appearance to those of normal deer. In fact, they were sometimes easier to locate i n the methallibure-treated deer because there were fewer of the purple basophils (gonadotrophs) with which they might be confused. 165 Type V: (gonadotroph; ICSH dell) These mucoid basophils were AB + ve, AT + ve, PAS + ve and OG -ve. Because their a f f i n i t y for AB, AT, and PAS i n the combined histochemical reactions was about equal, they stained a deep purple color (Figures 71, 72, 74 and 76). These cells are oval or angular with distinct boundaries. They are medium-sized to small (8-12 u), with a distinct large, round nucleus. The nucleus is eccentric or peripheral and usually contains a single large, central nucleolus. The cytoplasmic granules were often aggregated into large masses which sometimes f i l l e d the entire c e l l . The purple cells were concentrated in the antero-medial region (where they are often associated with blood vessels), but they were also found scattered throughout the pars.distalis. In the adult male they constituted about 10-20% of a l l c e l l s . They were less abundant in the pituitaries of male fawns, although partly granulated purple cells were common in the pars di s t a l i s of fawns in November and January. In May, most of the purple cells i n the pars d i s t a l i s of the adult males were partly granulated. In July they stained more intensely, owing to an accumulation of secretory granules. By September they were the most conspicuous c e l l type in the pars anterior, owing to their heavy granulation and dark purple color (Fi-gures 71 and 72). In November, nearly a l l these cells were partly granu-lated or degranulated, and many had become chromophobic (Figure 73). In January, some partly granulated purple cells were s t i l l v i s i b l e , but most were chromophobic. By March, a l l the type V cells were chromophobic. In striking contrast to the normal deer, the type V cells of a methallibure-treated deer were degranulated or chromophobic (Figures 77 and 78). The 166 seasonal cycle of cellular activity which was so obvious in the normal deer was not apparent in the methallibure-treated deer. Type VI: (gonadotroph; FSH cell) These mucoid cells stained magenta or pink after the combined histo-chemical reactions because they had a slig h t l y greater a f f i n i t y for PAS than AB or AT, and were OG-ve (Figures 75 , 76, 79 and 80 ) . They have an irregular oval or ellipsoid shape, and are the largest cells (15-20 y) found in the pars d i s t a l i s . The round or oval nucleus i s eccentric or peripheral and usually contains a large prominent nucleolus. The secretory granules, which are variable in size, are homogeneously distributed through-out the cytoplasm. The magenta or pink cells were most abundant in the antero-medial region, often occurring in groups or clusters near blood ves-sels. In the adult male they constituted about 5-10% of a l l c e l l s . Fully and partly granulated type VI cells were common in May, July and September. In November most of these cells were degranulated or chromophobic, but by January many were f u l l y granulated again. At the end of March, partly granu-lated cells with enlarged nuclei were common (Figure 6 6 ) . The magenta cells of the methallibure-treated deer were mostly degranulated or chromophobic. Although there was very l i t t l e cytoplasmic staining, these cells could usually be distinguished from other chromophobes by their large size and morphology. Type VII: (melanotroph; MSH cell ) These cells appear to be restricted to the pars intermedia. If they also occur in the pars d i s t a l i s , they might be confused with the type VI 167 magenta or pink c e l l s . They are AB + ve, AT + ve, PAS + ve, and OG - ve, and so they stain red-purple (Figure 81). The boundaries are indistinct, but these cells appear to have an irregular shape. The nuclei are round or oval, and the chromatin was usually dense and homogeneous. In some cells a distinct nucleolus was v i s i b l e . After staining, the entire pars intermedia took on a uniform red-purple color because of the medium to heavy granulation, and homogeneous distribution of granules in the cytoplasm of these c e l l s . No seasonal changes were noted in the histological appear-ance of these c e l l s . The pars intermedia of methallibure-treated deer was no different from that of normal deer. Large f o l l i c u l a r structures containing purple colloid were frequently found in or near the pars tuberalis, and in an area of the pars di s t a l i s adjacent to the pars intermedia (Figure 82). Purple colloid from these f o l l i c l e s was often found in the residual lumen between the pars intermedia and pars d i s t a l i s . The functional significance of these structures and the purple colloid they appear to secrete is not clear. ' 168 D. Discussion The tentative assignment of a functional identity to each of six c e l l types in the pars dist a l i s of deer was made by analogous comparison with the c e l l types described for other animal species where similar his-tochemical techniques have been applied. In most of these studies the identification of hypophysial cells was i n i t i a l l y based on their morpholo-gical and tin c t o r i a l or histochemical properties, and subsequently con-firmed with physiological evidence. Some of the research papers and re-views that were consulted for the purpose of making analogous comparisons were: fish and amphibians mouse rat Caviomorpha rabbit fur seal dog ferret bat Suiformes sheep cow human being (Van Oordt, 1968) (Barnes, 1962, 1963) (Costoff, 1973) (el Hakeem, 1971) (Allanson et_ al_., 1966) (Craig-Aman, 1972) (Purves and Griesbach, 1957) (Holmes, 1960; 1963) (Herlant, 1956; 1964) (Herlant and Ectors, 1969; Allanson, 1971; and Anderson et a l . , 1972) (Tassell and Kennedy, 1972) (Jubb and McEntee, 1955; Heath, 1965, 1970; and Maracek and Arendarcik, 1971) (Ezrin and Murray, 1963; Purves, 1961, 1966; Herlant and Pasteels, 1967; and Von Lawzewitsch, 1972). 169 Although there i s general agreement on.the identity of some c e l l types (e.g. the STH and LTH cells) there i s much confusion regarding the specific identity of others. An attempt was made to sort out pertinent information from the above sources and to incorporate i t into a table. This proved to be an invaluable tool for making comparisons and drawing analogies, but i t was much too large and detailed for inclusion. The descriptions which follow were derived from this table. In most species the somatotroph is a yellow (OG + ve), round to oval-shaped acidophilic c e l l with small (150-400 my in the rat, 350-500 my in the cow) secretory granules. It is one of the most abundant c e l l types, . comprising 50-60% of a l l cells in the rat, 20-30% in the Suiformes, and 37% in the female fur seal. Somatotrophs are found throughout the pars dis-t a l i s , but they are concentrated in the posterior and lateral areas, es-pecially near the periphery. The type I c e l l in the pars dist a l i s of the Columbian black-tailed deer has the same tinct o r i a l and morphological pro-perties as the somatotroph of other species, and was tentatively identified as such. Since these cells have smaller nuclei with condensed chromatin in November, January and March, they are probably less active during the winter months. Thus, the acidophils of Columbian black-tailed deer appear to undergo seasonal changes in secretory activity that correspond in part to the cyclic changes noted in the aggregate acidophil population of mule deer (Nicolls, 1971) and red deer (Stosic and Pantic, 1966). Since methallibure treatment did not appear to have an inhibitory i n -fluence On these c e l l s , i t i s unlikely that they are responsible for secret-ing the antler growth stimulating hormone. Instead, their apparent increase in activity during the spring and summer months could be related to 170 somatic growth and an increase in body weight during this period. In most species the lactotroph is very similar to the somatotroph in i t s morphology and distribution, but i t s secretory granules are larger (up to 800 my in the rat and 550-750 my in the cow). Tinctorially i t has been distinguished from the somatotroph in the rat, sheep and human being with erythrosin-OG, and with PAS-OG in the rat, ferret, seal, sheep, Suiformes, cow and human being, owing to i t s slight a f f i n i t y for erythrosin and PAS. The type II orange acidophils found in the pars d i s t a l i s of Columbian black-tailed deer have these same characteristics and were ten-tatively identified as lactotrophs. The relative abundance of this c e l l depends on sex and reproductive condition. In the male rat i t constitutes 10% of a l l c e l l s , and in the female, 40%. It is more frequently seen in the lactating cow, pregnant hiefer, and mid-cycle hiefer (Heath, 1970). In the male warthog, lacto-trophs constituted 20% of the c e l l population (Allanson, 1971). In the male Columbian black-tailed deer the lactotroph was less abundant than the somatotroph, and i t s seasonal cycle of activity was also different. These cells appeared to be most active in September and Novem-ber, whereas the somatotrophs appeared to be more active in May and July. Thus, the biphasic pattern (spring and f a l l ) of acidophil activity that was noted by Nicolls (1971) in mule deer could indicate seasonal changes in the secretory activity of the somatotrophs and lactotrophs. Increased prolactin secretion in the f a l l might f a c i l i t a t e androgen production and stimulate the accessory sex glands in deer, as i t does in some other mammals (Nicoll and Bern, 1971), but there is also a p o s s i b i l i t y that i t may act to antagonize gonadotrophin secretion (see p. 123 ), 171 Methallibure had no noticeable effect on the lactotrophs of black-tai l e d deer, and since i t does not interfere with prolactin secretion in other animals (Gerrits et_ al_., 1965; Garbers and F i r s t , 1968; Ben David, et a l . , 1971; Deis and Vermouth, 1973/74) i t is unlikely that the lactotroph produces the antler growth stimulating hormone. In many species the corticotroph has not been positively identified. Owing to i t s weak reaction with most stains i t has frequently been termed a chromophobe. However, i t has tentatively been identified as a large red-orange or brick-red c e l l in the Suiformes (Herlant and Ectors, 1969; Allanson, 1971), cow (Maracek and Arendarcik, 1971), and human being (Herlant and Pasteels, 1967) with the AB-PAS-OG and AT-PAS-OG technique. This c e l l has an oval or irregular shape and long cytoplasmic processes. The nucleus is eccentric and contains a distinct nucleolus. The cytoplasmic granules are very small (e.g., 50-180 my in the rat). The corticotroph is most common in the posteromedial and ventral part of the pars d i s t a l i s . Heath (1970) described a chromophobic f o l l i c u l a r c e l l in the bovine pituitary which f i t s the above description. Although this c e l l was mostly chromopho-bic, PAS + ve droplets were sometimes present in the cytoplasm. These cells were frequently arranged in groups (mostly in the basophilic zone) with microvillous processes extending into a matrix of PAS + Ve colloid. A l -though Heath did not identify this c e l l as the corticotroph, i t resembles the c e l l which Maracek and Arendarcik (1971) believe produces ACTH in the cow, and i t corresponds closely to the type III c e l l described in the pre-sent study. This c e l l type was tentatively identified as the corticotroph of the deer pituitary. 172 It appears that the type III cells vary with the seasons, but i t i s not clear what some of these changes mean. The cytoplasm stained more intensely in summer, whereas in winter i t ^ as chromophobic; but at this time large purple colloid deposits were observed within many of the f o l l i -cle-like structures. Perhaps this indicates an increase in the production of ACTH during the winter, when the stresses of cold and starvation are more pronounced. Similar seasonal changes were also noted in the p i t u i -taries of methallibure-treated deer, so i t i s unlikely that methallibure has a pronounced effect on the activity of these cel l s . The thyrotroph in the rat, mouse, fur seal, dog, ferret, Suiformes, cow, and human being is a medium-sized triangular or polygonal mucoid c e l l with very fine cytoplasmic, granules (e.g., 40-150 my in the rat) which often condense to form larger droplets in the periphery of the cytoplasm. These granules are selectively AB + ve with the AB-PAS-OG technique, and they stain blue-black with AT-PAS-OG. In certain applications of these tech-niques the cells may react slightly with PAS (Heath, 1970), and they are strongly PAS + ve when stained with aniline blue- PAS or PAS-OG. Herlant and Pasteels (1967) found that in the human being, these cells sometimes contain large PAS + ve droplets. These variations in staining reactions make identification based on ti n c t o r i a l properties unreliable. Maracek and Arendarcik (1971) identified the thyrotroph in the cow as an AB +_ve, AT + ve, slightly PAS + ve, blue-purple c e l l ; whereas the gonadotrophic ICSH c e l l was identified as a selectively AB + ve turquoise-blue c e l l . No physiological evidence was cited to support this. Heath (1970) was unable to make a clear-cut distinction between the blue and purple cells in the cow. However, some authorities have reported that the thyrotroph i s the 173 least abundant c e l l type. It i s scarce in the Amphibia (Van Oordt, 1968). In the rat i t constitutes only 1-2% of a l l cells (Costoff, 1973), and in the human being i t represents 2-7% of a l l the granular cells (Herlant and Pasteels, 1967). If the thyrotroph i s in fact the least abundant c e l l type, then many of the purple or blue-purple cells reported by other investigators are probably gonadotrophs. Unfortunately, many of the investigators who called these cells thyrotrophs had l i t t l e or no direct physiological evidence to support their interpretations. With these same limitations in mind, the fourth c e l l type in the deer, which was the least abundant c e l l , and had tin c t o r i a l and morphological properties that were similar to the thyrotrophs identified in some other mammals, was tentatively identified as the thyrotroph. Physiological evidence w i l l be presented later to support the suggestion that the more abundant type V purple cells are in fact gonadotrophs, and not thyrotrophs. The histolo-gical appearance of the type IV cells was similar in both normal and methal-libure-treated deer. The ICSH c e l l in some mammalian species has been described as a small to medium-sized oval or polygonal basophil with a relatively large eccentric nucleus. The cytoplasmic granules are medium-sized (75 - 235 my) in the rat (Costoff, 1973) and small in the human being (Herlant and Pasteels, 1967). These cells stain purple in the Suiformes (Allanson, 1971) and human being (Herlant and Pasteels, 1967) with the AB-PAS-OG and AT-PAS-OG techniques, but according to Herlant and Pasteels (1967) i t is not the se-cretory granules that stain with AB or AT, but the cytoplasmic inclusions. 174 The secretory granules are slightly PAS + ve, and so the ICSH c e l l has been described as a pink or red c e l l in the rat (Costoff, 1973), human being (Herlant and Pasteels, 1967) and fur seal (Craig-Aman, 1972). In the amphibian i t is a red-violet c e l l (Van Oordt, 1968). The situation becomes even more confusing when one considers Herlant's (1960) report that with the AB-PAS-OG technique, the ICSH c e l l in the rat stains red at pH 0.2, and light green at pH 3.0. However, the size and morphology of this c e l l is similar in many species, and some comparisons can be made on this basis. The ICSH cells are found scattered throughout the basophilic zone, but they are heavily concentrated in the antero-medial area of the pars d i s t a l i s , where they frequently associate with blood vessels. They constitute 5-10% of a l l cells in the pars di s t a l i s of the male rat, and 20% in the female (Costoff, 1973). In the male deer, the type V cells stained purple at both pH 0.2 and pH 3.0, but the intensity of staining was greater at pH 3.0. These c e l l s , which were f i l l e d with purple staining material in September, and contained l i t t l e or none of this material in November, were tentatively identified as ICSH ce l l s . This suggests that during the summer and early in the breed-ing season they are synthesizing material faster than they are releasing i t , whereas at the peak of the breeding season the rate of release exceeds the rate of synthesis. From January to March a decline in both the syn-thesis and release of hormone results in a smaller, chromophobic type of c e l l . These seasonal alterations in cellular activity could be expected to produce the pattern of testosterone secretion observed in.the adult male deer. Because methallibure had a pronounced inhibitory effect on the ac-t i v i t y of these cells,.they are almost certainly gonadotrophs, and not thyrotrophs. 175 The FSH c e l l in many species has been described as a large (often the largest), round or oval mucoid c e l l with a large peripheral nucleus, and small to medium-sized cytoplasmic granules. The weakly AB + ve, AT + ve and PAS + ve granules are evenly distributed in the cytoplasm. Some-times the cytoplasm of these cells contains a vacuole (e.g., Suiformes), or a large PAS + ve inclusion (e.g., rat and human being). The FSH c e l l commonly stains pink or red-purple with the AB-PAS-OG and AT-PAS-OG tech-niques. This c e l l type is found throughout the basophilic zone, although i t i s more concentrated in the antero-medial area. It constitutes 30% of a l l cells in the pars d i s t a l i s of the male rat , and 20% in the female (Costoff, 1973). In the deer, the type VI c e l l had similar morphological and ti n c t o r i a l properties, and was tentatively identified as the FSH c e l l . Although these cells did not show a marked increase in the accumula-tion of secretory material from May to September (as the type V cells did) , many became degranulated in November. This suggests that the rate of hor-mone release also exceeded the rate of synthesis in the f a l l . The occurrence of partly granulated cells with enlarged nuclei late in March suggests that an increase in synthetic activity may begin at this time. However, the possibility that the activity of these cells i s related to the i n i t i a t i o n of antler growth should be checked out by examining them in April as well as in March and May, because this is when the new antlers start to grow. In addition, the temporal pattern of FSH secretion and i t s relationship to spermatogenetic activity in March, Apr i l , May and June needs to be investi-gated and considered in relation to the activity of these c e l l s . 176 IV. SUMMARY AND CONCLUSIONS The hormonal regulation of reproduction and the antler cycle in the male Columbian black-tailed deer was investigated by measuring serum testosterone, testis volume, sperm production and antler growth of wild deer and of captive deer treated with methallibure and various hormones. A histological examination of the pituitary gland, adrenals, thyroids, testes and accessory sex glands of normal and methallibure-treated deer was also performed to study the functional relationships of these organs to reproduction and antler growth. Various constituents in the serum and urine of normal and methallibure-treated deer were measured to investigate more f u l l y the effects of methallibure on physiological function. Serum testosterone, testis volume and sperm production were maximal in November, at the height of the reproductive season. The activity of the accessory sex glands was also greatest at this time. When the peak of the breeding season had passed, spermatogenesis, testis volume, testosterone production, and the secretory activity of the accessory sex glands declined, until a minimum was reached in February or March. The antlers were also cast during this period, usually within several weeks after the serum testoster-one had dropped to a low level. In the spring, a significant increase in spermatogenetic activity was noted. This brief period of activity coincided with the i n i t i a t i o n of antler growth. Sperm production declined in June and July, but the seminiferous tubules were s t i l l more active than in Febru-ary or March. The.accessory sex glands also appeared more active during the spring and summer than in late winter. The serum testosterone level however, remained low (below 1 ng/ml) throughout the antler growing period. In the latter half of the summer, the reproductive organs began to show signs of redevelopment, and by late August or early September there was a no-ticeable increase in testosterone production. As the serum testosterone level rose, rutting behavior and velvet shedding ensued. Sperm production and the secretory activity of the seminal vesicles increased markedly as the peak of the breeding season approached. The concentration and v i a b i l i -ty of sperm was greatest between October and December, but some captive deer produced sperm a l l year round. These deer also had a higher serum testosterone level, and remained in rut longer than deer in the f i e l d . Spermatogenetic activity in the spring was also much more pronounced in the captive deer. Methallibure (ICI 33828), a non-steroidal inhibitor of gonadotrophin secretion, effectively suppressed reproduction and arrested antler develop-ment in deer. A dose level of 1.25 mg/kg per day was sufficient to prevent gonadotrophic stimulation of spermatogenesis and testosterone production. In the absence of sufficient quantities of testosterone, the partly grown antlers of methallibure-treated deer failed to mature, and the velvet was not shed. PMS and HCG, at dose levels of 3 x 667 and 3 x 933 I.U./wk were capable of stimulating the testes of methallibure-treated deer to secrete enough testosterone to induce velvet shedding, but neither hormone by i t s e l f was effective in completely restoring reproductive function. When methalli-bure was applied early in the spring, antler growth was completely suppressed. The subsequent administration of prolactin, PMS and some androgenic ster-oids during June and July did not produce a significant amount of antler growth. However, HCG did rejuvenate antler growth in one instance. 178 Antibody formation, and a gradual loss of v i a b i l i t y of the velvet during methallibure treatment may account in part for the poor growth response. that was achieved with some of the hormone injections. During the normal period of antler growth, the testes of methallibure-treated deer did not respond to the exogenous administration of HCG by se-creting testosterone, whereas both PMS and prolactin were effective in this respect. PMS was capable of producing a sudden and dramatic increase in testosterone production, whereas the effect of prolactin was more gradual and less pronounced. After the administration of PMS, HCG was capable of stimulating testosterone production, but i t was ineffective in the deer previously treated with prolactin. When the inhibition of gonadotrophin secretion was removed in the f a l l by terminating methallibure treatment, testosterone production and spermatogenesis soon recovered, but the deer that had previously received prolactin did not produce mature sperm until the following spring. These results indicate that PMS, HCG and prolactin are a l l capable of stimulating testosterone production, although HCG may only be effective at certain times of the year (e.g., in August but not in June), or under certain conditions (e.g., after the testes have been primed with FSH [PMS]). They also indicate that the testes do not always show a complete response to gonadotrophic stimulation, particularly after treatment with prolactin. From these and other observations i t appears that the testicular response may depend on the temporal sequence as well as the type of hormonal stimulation. A histological examination of the adrenal cortex and the thyroid gland revealed that the activity of these organs varies with the seasons, but the 179 extent to which they might influence the antler cycle was not apparent. Methallibure did not significantly depress thyroid function at the dose level that was administered, so i t i s unlikely that thyroxin provides the primary stimulus for antler growth. Methallibure has other effects in addition to the inhibition of gona-dotrophin secretion. It caused a reduction in feed intake, probably by affecting the appetite control centre of the hypothalamus. It also pro-duced a diabetes insipidus-like condition, which appeared to result from the suppression of ADH. An analysis of the blood chemistry revealed that methallibure did not significantly alter the circulating levels of any con-stituents other than alkaline phosphatase. The concentration of this en-zyme normally increased during the antler growing period (alkaline phospha-tase catalyzes the reaction which results in the calcification of antlers), whereas in the methallibure-treated deer i t did not. However, the methalli-bure-treated, HCG-injected deer that grew antlers showed a rise in alkaline phosphatase during the administration of HCG in June. Thus, one of the ef-fects of the gonadotrophins might be to stimulate the production of alkaline phosphatase in the growing antler. The application of histochemical staining techniques to the anterior pituitary revealed the presence of seven different c e l l types. Most c e l l types undergo cyclic changes that may be related to a seasonal pattern of functional activity. The c e l l type tentatively identified as the ICSH ce l l had the most pronounced seasonal cycle. Hormonal release by these c e l l s , and other cells tentatively identified as FSH c e l l s , was judged to be greatest in the f a l l , but some activity was also apparent during the spring and summer. The somatotrophs appeared most active in spring and 1 8 0 summer, whereas the lactotrophs appeared most active in the f a l l . With the possible exception of the corticotrophs, a l l the cells of the anterior pituitary showed the least signs of activity in late winter. At this .^ime, many cells were small and chromophobic. The pituitaries of methallibure-treated deer sacrificed in July and November contained ICSH cells and FSH cells that were small and chromophobi whereas other c e l l types appeared to be affected l i t t l e or not at a l l . Thi suggests that the main effect of methallibure on the anterior pituitary was to suppress the secretory activity of the gonadotrophic c e l l s . The serum testosterone data from the present study supports the hypo-thesis that testosterone i s responsible for the maturation and maintenance of antlers in the hard, functional, condition. It is concluded that tes-tosterone regulates this phase of the antler cycle. Experiments with me-thallibure support the hypothesis that a gonadotrophic hormone stimulates antler growth. In view of the fact that one gonadotrophin (ICSH) stimulate testosterone production, and either one or both (FSH and ICSH) stimulate antler growth, i t i s concluded further that the overall control of the ant-ler cycle i s regulated by the gonadotrophins. Increased spermatogenetic activity in the spring suggests that FSH is released at this time, and so i t could be responsible for i n i t i a t i n g antler growth. However, ICSH is also implicated, by virtue of the finding that HCG was capable of stimulat-ing antler growth in a methallibure-treated deer. If ICSH stimulates ant-ler growth, then for some reason the testes do not respond by secreting large amounts of testosterone i n the spring. There is some evidence to suggest that prolactin might be involved in preventing this response, 1 8 1 because i t appeared to cause a refractory state i n the deer testis. Thus, FSH, ICSH, and prolactin may a l l play a role in regulating the antler cycle. The gonadotrophins, prolactin, and testosterone also act i n various ways to regulate the reproductive cycle. When gonadotrophin secretion was inhibited by methallibure, testosterone production and spermatogenesis were suppressed. This occurs because ICSH is required to stimulate tes-tosterone production (which is necessary for spermatid formation), and FSH is required for the maturation of spermatids. The gonadotrophins and tes-tosterone are also needed to stimulate the activity of the accessory sex glands. Large amounts of testosterone are not required for this or for spermatogenesis, but a high circulating level of testosterone induces rut-ting behavior and provides maximal stimulation of the reproductive organs during the breeding season. The role of prolactin in regulating reproduction is s t i l l uncertain. It may augment the effects of the gonadotrophins and testosterone on the reproductive organs, and stimulate the testes to produce small amounts of testosterone, but this does not preclude the possibility that i t might also be antigonadal or antigonadotrophic. A simplified hypothetical scheme i l l u s t r a t i n g the hormonal regulation of reproduction and the antler cycle i s presented in Figure 83. The most quiescent period normally occurs in late winter, when the photoperiod is too short to induce hypothalamic stimulation of the gonado-trophic ce l l s . Since gonadotrophin secretion is minimal at this time, the testes are inactive, and almost no antler growth occurs. In the spring, an increase in the ratio of light to dark (ca. 12L/12D) induces hypothalamic 182 FIGURE 83. Hypothetical scheme i l l u s t r a t i n g how various hormones may act to regulate reproduction and the antler cycle. Details of the photoperiodic-hypothalamic-hypophysial axis and various feedback mechanisms are not known. The effects and interrelationships of certain hormones also require further study. The antler growth stimulating hormone (AGSH), a gonadotrophin, probably stimulates antler growth directly; the p o s s i b i l i t y that i t may act indirectly via the testes or adrenals is indicated by a broken line. 183 Exterocept ive factors ( e . g . , p h o t o p e r i o d , socia l st imuli ) 1 HyP< p o t h a l a m u s MM Releas ing h o r m o n e s a n d + Inhibit ing fac tors A n t e r i o r Pituitary G o n a d o t r o p h i c cells Lac to t rophs A G S H F S H ^ I C S H L t+ cor tex * t Stero id hormone T e s t e s Seminiferous Interstit ial tubules cells Spermatogenes is Testosterone Sertoli cells A c c e s s o r y sex glands N o growth G r o w t h J In v e l v e t Ant le rs M a t u r a t i o n J Velvet s h e d d i n g Pol ished ; (Cas t ing Funct ional 184 stimulation of the gonadotrophic c e l l s , and they respond by secreting a hormone which initiates antler growth. It appears that this hormone acts directly on the antler-growing tissue, but the pos s i b i l i t y that i t stimu-lates the testes or adrenals to secrete a steroid which promotes antler growth cannot be ruled out. The gonadotrophin responsible for i n i t i a t i n g antler growth also appears to reactivate the testes, but reproduction does not occur because the i n t e r s t i t i a l cells do not secrete large amounts of testosterone at this time. Perhaps the photoperiod i s approximately right, but the fact that i t i s increasing rather than decreasing makes a dif f e r -ence. Thus, the right combination of FSH and ICSH, and/or the proper tem-poral sequence necessary for a f u l l testicular response might not occur in the spring. There is some indication that prolactin may also play a role in preventing the testicular response, but this requires further study. In summer, long photoperiods continue to stimulate the production of the gona-dotrophic hormone responsible for antler growth; but by i t s e l f , this hormone is incapable of stimulating further increases in testicular activity. At the end of the summer the photoperiodic ratio again approaches 12L/12D, but this time i t is decreasing rather than increasing, and so hypothalamic s t i -mulation of the gonadotrophic cells results in increased production of the appropriate amounts of both FSH and ICSH. This stimulates redevelopment of the testes, and results in sufficient amounts of testosterone being produced to stimulate sperm production and to f a c i l i t a t e maturation of the antlers. In the f a l l , a decreasing photoperiod of approximately 12L/12D provides maximal hypothalamic stimulation of the ICSH-secreting c e l l s , and they re-lease a l l their stored secretory material. The i n t e r s t i t i a l cells of the testes respond by secreting large amounts of testosterone, which in turn 185 induces velvet shedding and rutting behavior. The continued production of ICSH, FSH, and a high concentration of testosterone ensure maximum sperm production. The gonadotrophins, prolactin, and testosterone may also act in synergistic fashion to stimulate maximal activity of the ac-cessory sex glands. Although the mechanisms and details are not known, a l l these hormones, and possibly an inhibiting factor released by the Sertoli cells as well, may provide feedback information for the hypothala-mo-pituitary axis. Negative feedback, and a continually decreasing photo-period i n winter, could lead to a reduction i n gonadotrophin secretion, and testicular regression. A decline i n testosterone production would hasten this process, and ultimately result in casting of the antlers. 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Univ. of British Columbia, Vancouver. 2 1 1 p. Tienhoven, A. van. 1 9 6 8 . Reproductive physiology of vertebrates. W.B. Saunders Co. 4 9 8 p. Tullock, M.I., J. Crooks and P.S. Brown. 1 9 6 3 . Inhibition of thyroid function of dithiocarbamoylhydrazine. Nature, Lond. 1 9 9 : 2 8 8 -2 8 9 . Urschel, F.M. 1 9 6 7 . Parathyroid glands of mule deer. M.Sc. Thesis, Colorado State Univ. Fort Collins. 8 2 p. Vermeulen, A. and L. Verdonck. 1 9 7 0 . Testosterone assays by competitive protein binding. Acta Endocrinol. 6 4 , suppl. 1 4 7 : 2 3 9 - 2 5 7 . Waldo, CM. and G.B. Wislocki. 1 9 5 1 . Observations on the shedding of the antlers of Virginia deer (Odocoileus hemionus borealis) Amer. J. Anat. 2 8 : 3 5 1 - 3 9 6 . Walpole, A.L. 1 9 6 5 . Non steroidal agents inhibiting pituitary gonadotro-phic function, P. 1 5 9 - 1 7 9 . In Austin and Perry [eds.]. Agents affecting f e r t i l i t y . Biol, council symp. on drug action, Churchill, London. West, N.O. 1 9 6 8 . The length of the estrous cycle in the Columbian black-tailed deer or coast deer (Odocoileus hemionus columbianus). B.Sc. Honours Thesis., Zool. Dept., Univ. of B.C., Vancouver, B.C. 3 1 p. Whitehead, P.E. and E.H. McEwan. 1 9 7 3 . Seasonal variation in the plasma testosterone concentration of reindeer and caribou. Can, J. Zool. 5 1 ( 6 ) : 6 5 1 - 6 5 8 . Wislocki, G.B. 1 9 4 3 . Studies on growth of deer antlers, p. 6 3 1 - 6 5 3 . In Essays in biology. Univ. Calif. Press. Wislocki, G.B., J.C Aub and CM. Waldo. 1 9 4 7 . The effects of gonadectomy and the administration of testosterone propionate on the growth of antlers in male and female deer. Endocrinol. 4 0 : 2 0 2 - 2 2 4 . Yousef, M.K. and J.R. Luick. 1 9 7 1 . Estimation of thyroxin secretion rate in reindeer (Rangifer tarandus): effects of sex, age and season. Comp. Biochem. Physiol. 4 0 A : 7 8 9 - 7 9 5 . Yousef, M.K., R.D. Cameron and J.R. Luick. 1 9 7 1 . Seasonal changes in hydrocortisone secretion rate of reindeer, Rangifer tarandus. Comp. Biochem. Physiol. 4 0 A : 4 9 5 - 5 0 1 . 196 VI APPENDIX A Techniques 197 A Competitive Protein Binding Technique for the Determination of Serum Testosterone Introduction The competitive protein binding assay for testosterone u t i l i z e s the principle of displacement of labelled testosterone from a specific binding protein by testosterone in the sample. It is basically the same as the radioimmunoassay, but i t uses a naturally occurring binding protein instead of an antibody. The binding protein is a 3-globulin found in high concen-trations in the blood of pregnant human females in their third trimester, and in human females taking the oral contraceptive Enovid E. High estrogen levels appear to stimulate the production of testosterone binding globulin (TBG). A review of competitive protein binding methods and their practical and theoretical aspects are discussed by various authors in Diczfalusy [ed.] 1970. Materials 3 3 Testosterone - 1, 2, 6, 7 - H (TH) specific activity 90 C/m mole (New England Nuclear Corp). A standard solution containing 3000 CPM 3 or 7000 DPM T H (0.01 ng) per microliter of ethanol was prepared from a 3 known amount of T H. Non-radioactive testosterone - (Mann Research) was made up into standard solutions containing 1 ng/ul and .01 ng/yl of t r i p l e d i s t i l l e d ethanol. Organic solvents - methylene chloride, ethyl acetate, chloroform and methanol (Fisher Scientific) were tri p l e d i s t i l l e d before use. F i l t e r paper - Whatman No. 7 qualitative, 12.5 cm. (Fisher Scientific) was 198 prewashed with 2 x 5 ml methylene chloride before f i l t e r i n g the extract. Thin layer chromatography (TLC) sheets - (Gelman 1TLC-SAF, s i l i c a gel impregnated glass fiber) were suspended between two strips of paper with paper clips and prewashed three times with methanol: methylene chloride (3:1) by descending chromatography in a paper chromatography chamber. Washed TLC sheets were activated for 15 minutes at 100°C and stored in a vacuum oven until use. Thin layer chromatography chamber - Eastman Chromatogram glass sandwich type (Fisher S c i e n t i f i c ) . Ammonium sulfate solution - was prepared by dissolving 100 grams of ammon-ium sulfate (Fisher Scientific) in 100 ml d i s t i l l e d water. TestOsterone-bindirtg globulin (TBG)-was obtained from the plasma of a hu-man female taking the oral contraceptive Enovid E. The plasma was divided into.5 ml aliquots and stored at -20°C unt i l use. Scin t i l l a t i o n f l u i d - was prepared by adding 4 grams Ommifluor (New England Nuclear) to 980 ml scintanalysed toluene (Fisher Scientific) and 20 ml methanol. Glassware - was routinely washed in Sparkleen detergent (Fisher S c i e n t i f i c ) , rinsed with tap water, then d i s t i l l e d water, and f i n a l l y in t r i p l e dis-t i l l e d ethanol. Methods Extraction The extraction procedure is the most c r i t i c a l part of this tech-nique because a relatively pure extract of testosterone must be obtained 199 to ensure that method interfering substances. (MIFS) are kept to a minimum. Some procedures that achieve this objective are limited in their applica-tion for routine use because they are long, tedious, and costly. The following procedure can be performed in less than 5 hours. MIFS are minimized by using t r i p l e d i s t i l l e d solvents and pre-washed TLC sheets. 3 Approximately 3000 CPM (7000 DPM) T H was added to each 40 ml extraction tube for recovery. Volumes of .125 to 4 ml (depending on the expected testosterone level) deer serum were added to each extraction tube. Serum volumes less than 4 ml were made up to 4 ml with d i s t i l l e d water. Extraction tubes containing 4 ml d i s t i l l e d water were also pro-cessed to determine blank values. Each sample was mixed b r i e f l y on a vortex mixer before extracting with 20 ml methylene chloride and 2 ml IN NaOH. Extraction was f a c i l i t a t e d by placing the extraction tubes on an automatic shaker for 20 minutes. The major portion of the top layer was aspirated and discarded before washing with 10 ml d i s t i l l e d water. The aqueous phase was similarly aspirated and discarded. Each sample was then f i l t e r e d through prewashed Whatman #7 f i l t e r paper to eliminate the remaining aqueous phase and the emulsified material that usually forms when deer plasma is extracted. The extraction tubes were rinsed with 5 ml methylene chloride and this was added to the f i l t r a t e . Evaporation of the methylene chloride was f a c i l i t a t e d by placing the tubes in a water bath at 40°C and passing a gentle stream of nitrogen over them. A simple evaporating apparatus was constructed by using the head of a pipet washer (Canlab) as a distributor and f i t t i n g i t with six pasteur pipettes. This apparatus ensures the delivery of a constant stream of nitrogen to each extraction tube (A similar evaporating apparatus was constructed to 200 accommodate nine of the 12 x 75 mm test tubes used in the assay procedure). After evaporation of the methylene chloride the sides of each tube were washed down with 400 ul ethyl acetate and again taken to dryness. The residue was redissolved in 40 ul ethyl acetate for spotting on TLC sheets. Thin Layer Chromatography Most researchers using the CPB technique for testosterone deter-mination have found i t necessary to use paper chromatography (PC) or one or more steps of TLC followed by PC in their purification procedure, in order to obtain low blank values. The PC system described by Maeda et a l . (1969; i.e. cyclohexane: p-dionane: methanal: water, 100: 25: 100 : 10), and several other PC systems were tried with variable success. Tests showed that 1, 4-di oxane can interfere with the CPB assay, possibly owing to impurities in the solvent or to i t s tendency to form peroxides when taken to dryness. Although Maeda et_ a l . (1969) did not indicate the mag-nitude of blanks obtained with this system, I found them to be higher than the blanks obtained with some TLC systems. Prewashed s i l i c a gel impreg-nated glass fiber sheets however, (see Materials section) yielded extracts that were sufficiently free from interfering substances to permit a single TLC step to be ut i l i z e d . Each sample and two samples of authentic testos-terone were spotted on a TLC sheet and developed for 45 minutes in chloro-form: ethyl acetate (80:20). The relative mobilities for various steroids in this system are given in Table II. Elution and Recovery The.spots of authentic testosterone were visualized by UV absorp-tion and sample areas 1.5 cm square corresponding to this zone were cut 201 TABLE II. Rf values of steroids in'the chloroform: ethyl acetate (80:20) Gelman 1TLC-SAF thin layer chromatography system. Running time 45 minutes; solvent front 15 cm from origin. Steroid Distance Travelled Rf (cm) 5 •* - Androstanediol 4.8 .320 5 oc - Androstenediol 5.2 .346 17 g-Testosterone 6.2 .413 Epitestosterone 6.3 .426 17 g-Estradiol 7.3 .486 Dehydroepiandrosterone 7.8 .520 Pregnenolone 8.1 .540 Dihydrotestosterone 8.3 .553 Androsterone 8.9 .593 Andros tenedione 9.5 .633 Estrone 11.0 1 .733 202 out and eluted with 1 ml methanol. The ease of handling s i l i c a gel impregnated glass fiber sheets permits the use of a simple but efficient elution apparatus originally designed for PC by Dominguez (1967). Six 10 ml glass syringes mounted in a small chamber were f i t t e d with 7/8 in. x 25 G needles bent into a hook at the t i p . The sections to be eluted were suspended from the needles. One ml of methanol was pipetted into each syringe and the eluent was collected in the 12 x 75 mm assay tubes. When methanol is used as a solvent, 95% of the testosterone is eluted within the f i r s t ten drops. Recovery The eluent was evaporated to dryness under a stream of nitrogen and redissolved in 2 ml ethanol. One half ml of this sample was pipetted into a s c i n t i l l a t i o n v i a l , evaporated to dryness and counted for recovery. The remainder was s p l i t into .5 ml and 1 ml aliquots, which were evaporated to dryness and assayed for testosterone. Dividing the sample into .5 ml and 1 ml aliquots increases the likelihood of obtaining at least one value for testosterone that f a l l s within the usable portion of the standard curve. However, both values were recorded, along with two other determina-tions performed on corresponding aliquots from an extract of twice the volume of the same serum. Thus, four assays were performed on two d i f f e r -ent volumes of each serum sample. Testosterone-binding solution (TBS) 3 800 ul of the T H standard solution was pipetted into a 125 ml flask and evaporated to dryness. Fifty five ml TBS (enough for 100 assays) was prepared by adding .5 ml TBG plasma and 54,5 ml d i s t i l l e d water. 203 This dilution (1:110) was satisfactory for the assay of 0.2 to 2.0 ng testosterone. The TBS solution remained stable for several weeks at 4°C. Assay Procedure The 12 x 75 mm assay tubes containing .5 ml and 1 ml aliquots of the serum extracts and amounts of the standard testosterone solution (.01 ng/yl) corresponding to 0, 0.4, 0.8, 1.2, 1.6, 2.0, 3.0 and 4.0 ng testosterone were evaporated to dryness. One half ml (25,000 CPM or 58,000 DPM) of the TBS was added to each tube and mixed gently on a vor-tex mixer for 5 sec. The tubes were incubated at room temperature for 1 hour. The protein-bound-testosterone was precipitated by adding 0.5 ml saturated ammonium sulfate solution. Each tube was mixed b r i e f l y on a vortex mixer, and allowed to stand for 5 minutes before centrifuging at 3000 RPM for 10 min. A 0.5 ml aliquot of the supernatant was pipetted into a counting vi a l and 10 ml s c i n t i l l a t i o n f l u i d was added. The count-ing vials were shaken vigorously and allowed to settle before counting. Each sample was counted for 20 min in a Nuclear Chicago Isocap 300 liquid s c i n t i l l a t i o n counter. Calculations The standard curve was obtained by plotting the CPM in 0.5 ml of the supernatant against the amount of testosterone added to each assay tube. The assay tubes containing 0.5 ml and 1 ml aliquots of the serum extracts must be corrected for the small amount of tracer testosterone added for the purpose of determining recovery. Thus, the percent unbound S = 100 X -j; + i/2R ^ o r t n e m* fraction* a n d S 100 X _ D for the 1.0 ml fraction, where S represents the CPM 204 in 0.5 ml of the supernatant; T represents the total possible counts (CPM in 0.5 ml pipetted from an assay tube containing 0.5 ml TBS and 0.5 ml ammonium sulfate solution); and R lepresents the CPM in the 0.5 ml aliquot counted for recovery. The percent unbound was then expressed in CPM (% unbound x T = CPM) and the amount of testosterone contained in the assay tube was estimated from the standard curve. This value was corrected for recovery and the blank value subtracted before expressing . i t in ng per ml of serum. Results The standard curve (Figure 84) was obtained by plotting the CPM in 0.5 ml of the supernatant against the amount of testosterone added to each assay tube. Recovery The mean recovery of testosterone in 50 serum samples of 2 ml or less was 65%. The mean recovery of testosterone in 50 serum samples of 2 to 4 ml was 50%. Because of wide variations i n recovery when different volumes of serum are extracted i t was necessary to determine the recovery for each sample. Accuracy Accuracy was measured by analysis of 4 ml aliquots of d i s t i l l e d water to which 0, 0.5, 0.0, 1.5, 2.0 and 4.0 ng testosterone had been added, ("steroid free" deer serum was not available). Figure 85 shows the testosterone obtained in the analysis of 5 samples (n = 4) plotted as a function of the concentration expected. The mean blank value (0.113 +_ S.D.; n = 14) is also plotted in Figure 85. The analysis of variance gives y = .113 + .949 x, where y is the concentration measured and x is the concentration expected. 205 FIGURE 84. Standard curve: CPB assay for testosterone. The radioactivity (CPM)in the supernatant (unbound) is plotted as a function of the unlabeled testosterone content of each tube. 20% unbound = 2,800 CPM 80% unbound = 11,200 CPM TBG dilution =1:110 206 207 FIGURE 85. Accuracy o f CPB t e s t o s t e r o n e Determination 208 E X P E C T E D (ng) 209 Precision Precision was examined by measuring the testosterone content of deer serum samples in the range 2.66 to 7.69 ng/ml, processed in duplicate on different days (Table III). Thus, the mean coefficient of variation (6.53%) represents intra-assay and interassay precision combined. Specificity Specificity was determined by comparing the results of CPB testos-terone assays performed on human plasma and deer serum with the results * from another laboratory u t i l i z i n g the double-isotope-derivative tech-nique, which uses different TLC systems and gas chromatography with an electron capture detector. The results of the testosterone determinations performed on five human and two duplicate deer samples with these d i f f e r -ent techniques were similar (Table IV). Sensitivity The minimum detectable concentration of testosterone was calculated by determining the point at which the standard error of the blank value intersects the slope of the line in Figure 85. This value represents a displacement of .142 ng. When the blank value is subtracted from this figure a value of 0.029 ng is obtained. The latter value corresponds closely to the minimum concentration detectable (0.03 ng) on the standard curve. • The results obtained with this procedure are similar to, and in some respects are an improvement over those achieved with earlier applica-* Dr. F. Leung, CP. Lab,, Vancouver General Hospital TABLE III. 210 Precision of CPB testosterone determinations. (Duplicate analyses of the same serum samples run on separate days). Sample Mean Standard Standard Coefficient of Size Deviation error variation (%) 4 2.66 .13 .07 4.98 4 2.86 .22 .11 7.78 4 3.18 .36 .18 11.35 4 3.21 .21 .11 6.54 4 4.44 .19 .09 4.24 4 4.70 .20 .10 4.21 4 4.79 .22 .11 4.66 4 5.09 .36 .18 7.03 4 5.39 .37 .18 6.84 4 5.47 .39 .19 7.07 4 5.48 .31 .15 5.71 4 5.69 .25 .13 4.45 4 7.03 .83 .42 11.83 4 7.28 .33 .16 4.46 4 7.69 .52 .26 6.74 Mean coefficient of variation: 6.53% TABLE IV. Specificity: Comparison of the CPB method to the double isotope derivative technique N Sample # DID Method* CPB Method (ng/ml) (ng/ml) Y-16 4.5 4.92 Male Deer 5.0 5.46 5.40 ,. 5.82 X = 4.75 x = 5.40 Y-23 4.0 3.56 Male Deer 2.3 3.38 Sept. 1970 3.06 2.74 x = 3.15 x = 3.19 To 7.7 8.48 Human Male 8.32 x = 8.40 Du 9.0 10.94 Human Male 10.80 x = 10.87 Fr 8.2 7.56 Human Male 7.52 x = 7.54 Qu 4.76 Human Male 5.5 4.55 x = 4.66 Br .34 Human Male .30 .17 x = .26 CP. Lab, Vancouver General Hospital 212 tions of this technique. During my study, this method was used rou-tinely for determining testosterone concentrations in the range 0.1 to 30 ng/ml, on over 600 blood samples. 213 OX - AB - PAS - Haem - OG stain for deer pituitaries (after Herlant, 1960) 1. Dewax slides in xylene and hydrate to d i s t i l l e d water. 2, Oxidize in equal parts of 0.6% aqueous KMnO^ , and 0.6% aqueous H 2S0 4. 1-1/2 minutes 3. Rinse in d i s t i l l e d water. 4. Bleach sections in 3% aqueous sodium metabisulphite. 30 seconds 5. Wash in tap water. 5 minutes 6. Stain in 1% Alcian blue in 1% acetic acid at pH 3.0. 30 minutes 7. Wash i n tap water. 5 minutes 8. Oxidize in 10% aqueous periodic acid. 15 minutes 9. Rinse thoroughly in d i s t i l l e d water. 10. Treat with Schiffs reagent. 20 minutes 11. Rinse in d i s t i l l e d water. 12. Stain in Mayer's Haemalum. 3 minutes 13. Blue and colorize in hot running tap water until sections become magenta. 5-10 minutes 14. Rinse i n d i s t i l l e d water. 15. Stain in 3% Orange G in 1% acetic acid. 3 minutes 16. Transfer direct to 1% acetic acid. 5 minutes 17. Differentiate Orange G b r i e f l y i n d i s t i l l e d water. 18. Dehydrate, clear and mount. 214 OX - AT - PAS - Haem - OG stain for deer pituitaries (after Paget and Eccleston, 1960). 1. Dewax slides in xylene and hydrate to d i s t i l l e d water. 2. Oxidize i n equal parts of 0.6% aqueous KMnO^  and 0.6% aqueous H^ SO^ . 3 minutes. 3. Rinse in d i s t i l l e d water. 4. Bleach in 3% aqueous sodium metabisulphite. 30 seconds. 5. Wash in running tap water. 5 minutes. 6. Rinse in 70% alcohol. 7. Stain in Aldehyde Thionin. 2 hours. 8. Wash in running tap water. 5 minutes. 9. Rinse i n d i s t i l l e d water. 10. Oxidize in 0.5% periodic acid. 10 minutes. 11. Rinse thoroughly in d i s t i l l e d water. 12. Treat with Schiffs reagent. 30 minutes. 13. Rinse in d i s t i l l e d water. 14. Stain in Mayer's Haemalum. 3 minutes. 15. Blue and colorize in hot running tap water until sections become magenta. 5-10 minutes. 16. Rinse in d i s t i l l e d water. 17. Stain in 3% Orange G in 1% acetic acid. 3 minutes. 18. Transfer direct to 0.5% acetic acid. 5 minutes. 19. Differentiate Orange G b r i e f l y in d i s t i l l e d water. 20. Dehydrate, clear and mount. 215 Reagents: Schiffs reagent - prepared by "cold Schiff" procedure according to R.D. L i l l i e ' s "Histopathologic Technic and Practical Histochemistry" 3rd Ed., 1965, p. 270. Aldehyde Thionin - Thionin (Fisher Scientific) 0.5 gm. 70% ethyl alcohol 91.5 ml. paraldehyde 7.5 ml. concentrated HCl 1.0 ml Ripen stain in a tightly stoppered bottle for 1 to 2 weeks. VII APPENDIX B Tabulated Deer Data 217 TABLE V-A. Seasonal changes in body weight, serum testosterone and the antler growth cycle of captive black-tai led deer under various experimental treatments. Deer No. U26. Born: Jur.o, 1963 Date Body wt. (kg) Testoster-one ^ (ng/ml) Antlers Experimental treatment 28/1/72 antlers dropped outdoor control 25/2/72 pedicles healed 30/4/72 new growth 25/5/72 73.6 .27 2.5 cm 29/6/72 78.2 .37 14 cm. 28/7/72 77.3 .25 33 cm 25/8/72 79.5 1.98 39 cm 6/9/72 velvet shedding 29/9/72 87.3 4.67 polished 26/10/72 86.8 2.35 polished 1/12/72 82.3 8.33 polished 28/12/72 79.1 .96 polished 18/1/73 antlers dropped 31/1/73 73.6 .38 pedicles healing 2/3/73 71.8 .34 pedicles healed 5/4/73 70.5 .29 pedicles healed 2/5/73 71.4 .33 new growth; 1 cm 31/5/73 72.7 .29 3 cm 28/6/73 72.7 .24 12 cm 30/7/73 74.1 .47 30 cm 25/8/73 35 cm 10/9/73 velvet shedding 30/10/73 polished 23/1/74 * antlers dropped A l l testosterone values are expressed as the mean of duplicate determinations. For standard error and coefficient of variat ion, see Appendix A. J 218 TABLE VI-A. Seasonal changes in body weight, serum testosterone and the antler growth cycle of captive black-tailed deer under various experimental treatments. Deer No. Wl. Bom: June., 1965 Date Body wt, (kg) Testosterone (ng/ml) Antlers Experimental Treatment 20/3/69 25/7/69 3.00 polished r te polished 11/8/69 71.4 .90 18/8/69 antlers dropped 21/8/69 68.2 .50 pedicles healing 3/9/69 67.3 .27 pedicles healed 10/9/69 new growth 17/9/69 65.9 .35 1 cm 2/10/69 65.9 .29 2 cm 16/10/69 65.9 .28 8 cm 6/11/69 65.9 .42 17 cm 2/12/69 67.7 .45 30 cm 27/12/69 68.6 .56 40 cm 15/1/70 70.4 .81 50 cm 10/2/70 72.7 1.58 50 cm 27/2/70 velvet shedding 7/3/70 75.9 .98 polished 25/3/70 79.5 2.00 polished 15/4/70 80.0 4.04 polished 7/5/70 80.0 2.92 polished 27/5/70 80.0 2.99 polished 17/6/70 78.6 2.16 polished 17/6/70 78.6 2.16 polished 8/7/70 76.8 2.02 polished 29/7/70 74\5 1.08 polished velvet shed- 24L; (continuous light from 1/4/66; constant cont'd. 219 Table VI-A (cont'd) Date Body wt. Testosterone Antlers Experimental (kg) (ng/ml) Treatment 19/8/70 9/9/70 20/9/70 2/10/70 22/10/70 11/11/70 8/12/70 31/12/70 21/1/71 12/2/71 4/3/71 25/3/71 14/4/71 6/5/71 28/5/71 17/6/71 10/7/71 24/7/71 30/7/71 15/8/71 28/8/71 25/9/71 19/10/71 25/10/71 11/11/71 21/12/71 21/1/72 71.8 70.9 69.1 67.7 66.4 66.4 66.8 68.2 70.0 71.8 72.7 73.6 74.5 73.6 72.7 71.8 63.6 65.0 70.9 70.9 69.5 67.7 .50 .24 .37 .23 .20 .48 1.21 1.84 .89 2.22 2.00 5.46 1.84 2.40 .39 .20 .29 .26 1.40 1.51 3.50 3.35 polished antlers dropped new growth 1.5 cm 3 cm 12 cm 22 cm 45 cm 48 cm velvet shedding; 48 cm polished polished polished polished polished polished polished antlers dropped pedicles healing new growth 2 cm 6 cm 7 cm velvet shedding polished polished polished 24L; (continuous light from 1/4/66; constant temperature at 25°C) natural photoperiod cont'd 220 Table VI-A (cont'd) Date Body wt. Testosterone Antlers Experimental (kg) (ng/ml) Treatment 24/2/72 10/3/72 22/3/72 4/4/72 28/4/72 26/5/72 1/6/72 28/6/72 27/7/72 1/8/72 11/8/72 23/8/72 24/8/72 8/9/72 11/9/72 27/9/72 1/10/72 27/10/72 24/11/72 27/12/72 26/1/73 6/3/73 23/3/73 67.7 65.9 62.7 60.9 60.9 60.5 61.8 65.0 62.7 61.4 60.5 60.9 60.9 59.5 58.6 1.32 .55 .30 .37 .62 2.20 ,34 ,75 9.01 2.07 3.00 .98 .84 polished antlers dropped pedicles healed new growth 3 cm 3.5 cm 3.5 cm 4 cm 5 cm 5 cm velvet shedding polished polished antlers dropped scabs pedicles healing polishing pedicles pedicles bare polished polished polished polished bony caps shed methallibure; 1.25 mg/kg/day LTH: 3X933 IU/wk terminated LTH HCG: 3X933 IU/wk terminated HCG off methallibure 29/3/73 .40 new growth . cont' d 221 Table VI-A (cont'd) Date Body wt. Testosterone Ckg) (ng/ml) Antlers Experimental Treatment 9/4/73 1/5/73 30/5/73 1/6/73 26/6/73 27/6/73 4/7/73 30/7/73 14/8/73 1/9/73 11/9/73 30/9/73 21/10/73 22/2/74 56.4 56.4 56.4 56.8 .21 ,21 ,16 .34 5 cm 6 cm velvet dying; 6 cm 6 cm 8 cm 8 cm velvet dead; 8 cm 8 cm 8 cm renewed growth; 9 cm 12 cm 20 cm velvet shedding; 20 cm dropped antlers methallibure; 1.25 mg/kg/day LH (NIH-B8); 3 x 8 U/wk terminated LH off methallibure 222 TABLE VII-A. Seasonal changes i n body weight, serum testosterone and the antler growth cycle of captive black-tailed deer un-der various experimental treatments. Deer No. W3. Born: June, 1965 Date Body wt. (Kg) Testosterone (ng/ml) Antlers 10/4/69 in velvet; 10 cm 10/5/69 velvet shedding 25/7/69 83.6 5.30 . polished; 12 cm 21/8/69 82.3 4.21 polished 3/9/69 84.1 2.09 polished 17/9/69 82.7 2.13 polished 1/10/69 81.4 2.88 polished 16/10/69 80.9 4.30 polished 7/11/69 77.7 4.56 polished 4/12/69 76.4 1.44 polished 23/12/69 74.5 .51 polished 15/1/70 73.6 .39 polished 4/2/70 antlers dropped 13/2/70 72.3 .79 pedicles healing 6/3/70 70.9 .86 new growth 25/3/70 69.1 .25 3 cm 15/4/70 67.7 .51 8 cm 7/5/70- 71.8 .36 15 cm 27/5/70 75.0 1.55 20 cm 4/6/70 76.8 velvet shedding 17/6/70 78.2 5.08 polished; 20 cm Experimental Treatment 24L; (continuous light from 1/4/66; constant tempera-ture at 25°C) ....cont•d 223 Table VII-A (cont'd) Date Body wt. Ckg) Testosterone (ng/ml) Antlers 8/7/70 79.1 2.62 polished 29/7/70 75.0 .28 20 cm 3/8/70 antlers dropped 19/8/70 77.3 .92 scabs 9/9/70 78.2 3.83 scabs 26/9/70 polishing pedicles 2/10/70 77.3 8.88 pedicles bare 11/11/70 78.2 polished 7/12/70 80.9 3.69 polished 31/12/70 79.5 .45 polished 21/1/71 77.3 .33 bony caps shed 28/1/71 pedicles healing 5/2/71 new growth 13/2/71 75.5 .88 2 cm 4/3/71 74.1 .82 4 cm 27/3/71 75.0 1.00 7 cm 15/4/71 76.4 .65 15 cm 4/5/71 velvet shedding; 15 cm 6/5/71 79.1 1.15 polished 28/5/71 80.1 6.95 polished 17/6/71 81.4 6.32 polished 8/7/71 - 80.1 4.42 polished 10/7/71 polished i 29/7/71 80.1 .68 polished 17/8/71 antlers dropped Experimental Treatment terminated 24L natural photoperiod ,..,cont'd 224 Table VII-A cont'd Date Body wt. (kg) Testosterone (ng/ml) A n t l e r s Experimental Treatment 27/8/71 15/9/71 24/9/71 16/10/71 21/10/71 20/11/71 23/12/71 27/1/72 17/2/72 25/2/72 10/3/72 23/3/72 28/4/72 30/5/72 27/6/72 26/7/72 23/8/72 25/9/72 27/9/72 27/10/72 24/11/72 27/12/72 5/3/73 79.1 83.6 85.0 85.0 83.6 80.0 78.6 77.3 77.3 75.0 72.7 70.9 71.8 70.5 65.9 70.9 77.3 76.8 75.9 74.5 70.0 .14 1.10 7.09 9.44 2.95 .85 .45 .66 .21 .23 .16 .12 .30 1.07 .28 .40 .35 p e d i c l e s h e a l i n g new growth 1.3 cm v e l v e t shedding p o l i s h e d p o l i s h e d p o l i s h e d p o l i s h e d a n t l e r s dropped p e d i c l e s healed new growth .7 cm 2 cm 5 cm 10 cm 13 cm 15 cm v e l v e t shedding p o l i s h e d p o l i s h e d p o l i s h e d a n t l e r s dropped p e d i c l e s healed m e t h a l l i b u r e ; 1.00 mg/kg/day m e t h a l l i b u r e ; 1.25 mg/kg/day 225 TABLE VIII-A. Seasonal changes in body weight, serum testosterone and the antler growth cycle of captive black-tailed deer un-der various experimental treatments. Deer No. Y9. Born: June, 1967 Date Body wt, (kg) Testosterone (ng/ml) Antlers Experimental Treatment 23/7/69 - 1.09 in velvet 11/8/69 50.9 1.75 in velvet 1/9/69 velvet shedding 3/9/69 56.4 2.53 polished 20/9/69 58.2 6.73 polished 2/10/69 59.1 7.17 polished 22/10/69 60.9 10.54 polished 13/11/69 59.5 6.84 polished 4/12/69 55.5 6.30 polished 27/12/69 54.1 4.11 polished 15/1/70 54.5 1.53 polished 11/2/70 53.6 4.03 polished 7/3/70 52.7 .21 polished 19/3/70 antlers dropped 27/3/70 52.7 .18 pedicles healing 17/4/70 53.6 .29 pedicles healed 30/4/70 new growth 8/5/70 54.5 .25 1 cm outdoor control .corit' d 226 Table VIII-A cont'd Date Body wt. Testosterone Antlers Experimental (kg) (ng/ml) Treatment 28/5/70 55.5 .49 3 cm 19/6/70 57.3 .41 10 cm 10/7/70 58.2 .46 27 cm 3/8/70 60.9 1.81 36 cm 20/8/70 62.3 1.98 46 cm 28/8/70 velvet shedding 11/9/70 66.8 3.31 polished indoor control; 3/10/70 70.9 10.90 polished ( a r t i f i c i a l light 23/10/70 70.7 16.22 polished natural photoperi 13/11/70 70.5 17.93 polished od; constant tem-8/12/70 66.4 12.49 polished perature at 25°C) 2/1/71 64.5 1.51 polished 21/1/71 62.7 6.89 polished 13/2/71 62.7 .40 polished 6/3/71 61.8 .22 polished 27/3/71 62.7 .17 polished 10/4/71 62.7 antlers dropped 16/4/71 62.7 .02 pedicles healing 7/5/71 62.7 .13 pedicles healed 15/5/71 new growth 29/5/71 63.2 .28 5 cm 1/6/71 6 cm methallibure;l.25 18/6/71 61.8 .20 10 cm mg/kg/day 10/7/71 61.8 .52 15 cm 30/7/71 61.8 .16 18 cm 28/8/71 62.3 .16 18 cm 25/9/71 65.0 .24 18 cm ....cont'd 227 Table VIII-A cont'd Date 19/10/71 2/11/71 11/11/71 22/12/71 10/1/72 27/1/72 11/2/72 Body wt. Testosterone (kg) (ng/ml) Antlers 63.2 59.1 56.4 53.6 .26 .17 .28 .02 18 cm; velvet dying velvet dead; 18 cm 18 cm antlers dropped scabs scabs Experimental Treatment control for Y16 off methallibure 228 TABLE IX-A. Seasonal changes in body weighty serum testosterone and the antler growth cycle of captive black-tailed deer under var-ious experimental treatments. Deer No. Y16. Born: June.. 1967 Date Body wt, (kg) Testosterone (ng/ml) Antlers Experimental Treatment 23/7/69 66.4 1.25 in velvet 8/8/69 67.3 1.37 in velvet 20/8/69 67.3 1.99 in velvet 25/8/69 velvet shedding 3/9/69 71.8 1.50 polished 18/9/69 73.6 3.29 polished 2/10/69 76.8 3.88 polished 17/10/69 76.8 3.39 polished 7/11/69 ' 75.9 5.33 polished 3/12/69 73.2 13.15 polished 27/12/69 69.1 9.71 polished 17/1/70 67.7 5.00 polished 19/2/70 67.3 1.28 polished 2/3/70 antlers dropped 6/3/70 65.0 .39 pedicles healing 25/3/70 65.0 .39 pedicles healed 15/4/70 64.5 .43 new growth 7/5/70 65.9 .48 3.5 cm 25/5/70 67.3 .57 10 cm 17/6/70 70.5 .33 20 cm 8/7/70 72.7 .30 38 cm 29/7/70 76.4 .43 45 cm 20/8/70 79.5 1.15 velvet dying; 45 cm 4/9/70 Velvet shedding indoor control; ( a r t i f i c i a l light-natural photoperi-od; constant tem-perature 25°C) ....cont' d 229 Table IX-A cont'd Date Body wt. (kg) Testosterone (ng/ml) A n t l e r s 9/9/70 84.1 7.29 p o l i s h e d 2/10/70 85.0 3.88 p o l i s h e d 22/10/70 84.1 2.80 p o l i s h e d 11/11/70 81.8 11.83 p o l i s h e d 7/12/70 78.2 8.59 p o l i s h e d 31/12/70 75.0 5.38 p o l i s h e d 21/1/71 72.7 2.39 p o l i s h e d 12/2/71 71.4 2.64 p o l i s h e d 4/3/71 70.5 .34 p o l i s h e d 17/3/71 a n t l e r s dropped 25/3/71 69.5 .18 p e d i c l e s h e a l i n g 14/4/71 69.1 .47 p e d i c l e s healed 20/4/71 new growth 6/5/71 71.8 .06 3 cm 28/5/71, 74.5 ,30 14 cm 1/6/71 15 cm 17/6/71 76.4 .18 30 cm 8/7/71 73.6 .04 38 cm 29/7/71 76.4 .27 38 cm 27/8/71 78.6 .27 38 cm 24/9/71 75.5 .18 38 cm 21/10/71 71.8 .28 v e l v e t dying; 38 cm 2/11/71 69.5 v e l v e t dead; 38 cm 10/11/71 68.2 v e l v e t shedding 20/11/71 65.0 20.98 p o l i s h e d 20/12/71 59.1 p o l i s h e d 22/12/71 56.8 16.66 p o l i s h e d Experimental Treatment methallibure;1.25 mg/kg/day PMS:3X667 IU/wk terminated PMS ....cont'd 230 Table IX-A cont'd Date Body wt. Testosterone ' Antlers Experimental (kg) (ng/ml) Treatment 23/12/71 polished HCG:3X667 IU/wk 27/1/72 54.5 .16 polished 4/2/72 antlers dropped 11/2/72 53.6 .22 scabs terminated HCG 14/2/72 scabs off methallibure 231 TABLE X-A. Seasonal changes i n body-weight, serum te s t o s t e r o n e and the a n t l e r growth c y c l e o f c a p t i v e b l a c k - t a i l e d deer under var-. ious experimental treatments. Deer No. Y23. Born: June, i?67 Date Body wt. Testosterone A n t l e r s (kg) (ng/ml) 21/8/69 55.0 2.96 i n v e l v e t 27/8/69 59.1 v e l v e t shedding 18/9/69 61.8 1.68 p o l i s h e d 1/10/69 65.9 2.51 p o l i s h e d 17/10/69 68.2 4.54 p o l i s h e d 7/11/69 68.2 17.37 p o l i s h e d 3/12/69 65.9 10.63 p o l i s h e d 27/12/69 68.2 7.18 p o l i s h e d 17/1/70 69.5 2.24 p o l i s h e d 10/2/70 70.0 1.69 p o l i s h e d 27/2/70 a n t l e r s dropped 7/3/70 68.2 .34 p e d i c l e s h e a l i n g 25/3/70 65.9 .17 p e d i c l e s healed 10/4/70 65.0 new growth 15/4/70 65.0 .32 2 cm 7/5/70 65.9 .26 8 cm 27/5/70 65.9 .34 17 cm 17/6/70 68.2 .44 30 cm 8/7/70 69.5 • 47 45 cm 29/7/70 71.8 .45 57 cm 20/8/70 79.1 1.93 v e l v e t dying;57 cm 30/8/70 v e l v e t shedding 9/9/70 84.5 2.78 p o l i s h e d 2/10/70 87.7 2.00 p o l i s h e d Experimental Treatment indoor c o n t r o l ; ( a r t i f i c i a l l i g h t - n a t u r a l photoperiod; con-s t a n t temperature 25°C) , cont'd 232 Table X-A cont'd Date Body wt. Testosterone A n t l e r s Experimental (kg) (ng/ml) Treatment 22/10/70 85.0 16.65 p o l i s h e d 2/11/70 80.5 20.04 p o l i s h e d 7/12/70 78.2 14.13 p o l i s h e d 31/12/70 77.3 5.04 p o l i s h e d 21/1/71 76.4 1.80 p o l i s h e d 12/2/71 75.5 1.98 p o l i s h e d 5/3/71 74.1 .41 p o l i s h e d 10/3/71 a n t l e r s dropped 25/3/71 69.5 .42 p e d i c l e s h e a l i n g 14/4/71 67.3 .45 new grovth 6/5/71 68.2 .40 6 cm 29/5/71 70.0 .33 17 cm 17/6/71 73.2 .30 31 cm 8/7/71 73.6 .86 45 cm 29/7/71 75.0 .55 62 cm 25/8/71 v e l v e t shedding; 64 cm 27/8/71 76.8 2,33 p o l i s h e d 233 TABLE XI-A. Seasonal changes i n body weight, serum t e s t o s t e r o n e and the a n t l e r growth c y c l e o f c a p t i v e b l a c k - t a i l e d deer under var-ious experimental treatments. Deer No. Z l . Born: June, 1968 Date Body wt. Ckg) Testosterone (ng/ml) A n t l e r s Experimental Treatment 1/8/69 i n v e l v e t ; 15 cm 13/8/69 51.4 v e l v e t shedding 22/8/69 5.84 p o l i s h e d 6/9/69 53.2 6.21 p o l i s h e d 20/9/69 57.7 7.54 p o l i s h e d 2/10/69 59.1 1.95 p o l i s h e d 19/10/69 63.2 6.78 p o l i s h e d 13/11/69 64.5 5.36 p o l i s h e d 6/12/69 63.6 3.47 p o l i s h e d 27/12/69 60.0 2.89 p o l i s h e d 15/1/70 55.9 .79 p o l i s h e d 31/1/70 52.3 a n t l e r s dropped 11/2/70 49.5 .86 scabs 7/3/70 51.8 .33 p e d i c l e s h e a l i n g 27/3/70 53.2 .36 p e d i c l e s healed 17/4/70 55.0 .30 new growth 8/5/70 58.2 .33 2 cm 27/5/70 59.5 .35 6 cm 19/6/70 62.7 .20 14 cm 10/7/70 65.5 .23 33 cm 31/7/70 70.5 .47 40 cm 20/8/70 74.1 2.26 50 cm 1/9/70 v e l v e t shedding 11/9/70 78.6 4.39 p o l i s h e d outdoor c o n t r o l , cont'd Table XI-A cont'd 234 Date Body wt. kg. Testosterone (ng/ml) A n t l e r s Experimental Treatment 3/10/70 23/10/70 12/11/70 7/12/70 31/12/70 21/1/71 12/2/71 15/2/71 4/3/71 25/3/71 14/4/71 6/5/71 28/5/71 17/6/71 6/7/71 30/7/71 20/8/71 28/8/71 25/9/71 19/10/71 11/11/71 21/12/71 21/1/72 23/2/72 24/2/72 24/3/72 21/4/72 25/5/72 84.1 84.1 83.2 79.1 75.9 74.1 72.3 70.0 70.0 70.0 69.5 70.5 72.3 72.3 72.7 79.1 84.1 82.7 84.1 82.3 75.5 70.9 70.0 73.2 77.7 3.43 10.63 9.32 3.15 .74 .51 .31 .47 .14 .17 .21 .40 .27 .45 1.30 5.17 12.61 13.14 6.08 5.34 2.25 .23 .73 .41 .12 p o l i s h e d p o l i s h e d p o l i s h e d p o l i s h e d p o l i s h e d p o l i s h e d p o l i s h e d a n t l e r s dropped scabs p e d i c l e s healed new growth 2 cm i 6 cm 12 cm 30 cm 50 cm v e l v e t shedding; 52 cm p o l i s h e d p o l i s h e d p o l i s h e d p o l i s h e d p o l i s h e d a n t l e r s dropped scabs p e d i c l e s healed new growth 12 cm ..cont'd 235 Table XI-A cont'd Date Body wt. Testosterone A n t l e r s Experimental (kg) (ng/ml) Treatment 27/6/72 80.5 1.04 38 cm 28/7/72 82.7 1.14 60 cm 25/8/72 88.2 6.07 v e l v e t shedding; 65 cm 29/9/72 96.4 25.74 p o l i s h e d 26/10/72 95.5 13.97 p o l i s h e d 30/11/72 90.5 7.34 p o l i s h e d 28/12/72 84.5 1.11 p o l i s h e d 29/1/73 81.1 5.74 p o l i s h e d 2/3/73 81.1 .36 p o l i s h e d 6/3/73 a n t l e r s dropped 5/4/73 81.1 .51 p e d i c l e s healed 15/4/73 new growth 2/5/73 82.3 .54 5 cm 31/5/73 84.5 .51 20 cm 28/6/73 89.1 2.11 43 cm 30/7/73 1.94 62 cm TABLE XII-A. Seasonal changes in body weight, serum testosterone and the antler growth cycle of captive black-tailed deer under var-ious experimental treatments. Deer No. Z7. Born: June, 1968 Date Body wt. Testosterone Antlers Experimental (kg) (ng/ml) Treatment 3/10/70 77.7 14.60 polished 23/10/70 70.5 30.21 polished 3/11/70 65.0 4.78 polished 8/12/70 59.5 11.00 polished 2/1/71 58.6 2.71 polished 22/1/71 56.8 3.98 polished 13/2/71 55.5 3.03 polished 6/3/71 55.0 .58 polished 13/3/71 antlers dropped 27/3/71 59.5 .56 pedicles healing 16/4/71 60.9 .18 pedicles healed 30/4/71 new growth 7/5/71 65.0 .38 1.5 cm 29/5/71 65.9 .21 6 cm 18/6/71 68.2 .41 15 cm 10/7/71 71.8 .19 30 cm 30/7/71 73.2 .36 40 cm 28/8/71 81.4 6,87 velvet shedding; 42 cm 25/9/71 86.4 13.54 polished 19/10/71 80.9 14.04 polished 11/11/71 72.7 17.14 polished 21/12/71 66.8 14.38 polished 21/1/72 68.2 6.62 polished 24/2/72 65.0 9.04 polished outdoor control .cont'd 237 Table XII-A cont'd Date Body wt. Testosterone A n t l e r s Experimental (kg) (ng/ml) Treatment 24/3/72 68.6 1.15 p o l i s h e d 9/4/72 a n t l e r s dropped 21/4/72 70.5 .13 p e d i c l e s h e a l i n g 30/4/72 new growth 25/5/72 75.0 .18 4 cm 29/6/72 80.5 ,50 19 cm 28/7/72 83.6 .43 43 cm 25/8/72 82.7 4.27 v e l v e t dying; 43 cm 31/8/72 v e l v e t shedding 29/9/72 84.5 27.08 p o l i s h e d 26/10/72 77.3 18.44 p o l i s h e d 30/11/72 76.8 10.88 p o l i s h e d 28/12/72 76.4 10.82 p o l i s h e d 29/1/73 73.6 7.79 p o l i s h e d 2/3/73 69.1 4.70 p o l i s h e d 2/4/73 a n t l e r s dropped 5/4/73 77.3 .28 p e d i c l e s h e a l i n g 20/4/73 new growth 2/5/73 81.4 .22 1.5 cm 31/5/73 83.2 .30 6 cm 28/6/73 86.4 .95 20 cm 30/7/73 86.4 2.21 36 cm 30/8/73 v e l v e t shedding 30/9/73 p o l i s h e d 20/3/74 a n t l e r s dropped TABLE XIII-A. Seasonal changes in body weight, serum testosterone and the antler growth cycle of captive black-tailed deer under var-ious experimental treatments. Deer No. Bl. Born: June, 1970 Date Body wt. (kg) Testosterone (ng/ml) Antlers Experimental Treatment 29/10/71 7.74 polished; 16 cm outdoor control 6/12/71 8.90 polished 10/1/72 1.93 polished 11/2/72 .42 antlers dropped 10/3/72 28/4/72 55.5 50.9 .42 .40 new growth 3 cm methallibure; 1.25 mg/kg/day 31/5/72 50.0 .18 4 cm 1/6/72 4 cm LTH:3X933 I.U./wk 27/6/72 51.8 .20 4 cm 26/7/72 55.0 3.01 5 cm terminated LTH 1/8/72 5 cm HCG:3X933 I.U./wk 11/8/72 velvet shedding; 5 cm 23/8/72 60.0 ,70 polished terminated HCG 28/9/72 59.1 .09 polished 3/10/72 antlers dropped 4/10/72 56.8 scabs off methallibure 26/10/72 54.1 1.95 polishing pedicles 1/12/72 57.3 5.22 pedicles polished 28/12/72 53.6 .94 polished 26/1/73 shed bony caps 31/1/73 50.9 .32 pedicles healing . . enni-1 d 239 Table XIII-A cont'd Date 2/3/73 5/4/73 6/4/73 2/5/73 31/5/73 28/6/73 Body wt, (kg) 50.9 50.0 53.6 48.2 Testosterone (ng/ml) A n t l e r s ,34 ,21 .25 .57 p e d i c l e s healed new growth; 1 cm 1 cm 1 cm 1 cm 1 cm Experimental Treatment m e t h a l l i b u r e ; 1.25 mg/kg/day TABLE XIV-A. Seasonal changes i n body weight, serum t e s t o s t e r o n e and the a n t l e r growth c y c l e o f c a p t i v e b l a c k - t a i l e d deer under var-ious experimental treatments. Deer No. B2. Born: June, 1 9 7 0 Date Body wt. Testosterone A n t l e r s Experimental (kg) (ng/ml) Treatment 24/9/71 p o l i s h e d ; 18 cm outdoor c o n t r o l 28/10/71 6.04 p o l i s h e d 6/12/71 6.91 p o l i s h e d 10/1/72 1.96 p o l i s h e d 1/2/72 a n t l e r s dropped 11/2/72 52.3 .35 scabs 10/3/72 50.0 .16 p e d i c l e s healed m e t h a l l i b u r e ; 1.25 mg/kg/day 28/4/72 45.5 .15 .5 cm 31/5/72 43.2 .34 .5 cm m e t h a l l i b u r e ; 10/6/72 new growth; 1 cm 1.0 mg/kg/day 27/6/72 45.0 .06 5 cm 26/7/72 47.7 .11 10 cm 23/8/72 52.7 .28 16 cm 5/9/72 v e l v e t shedding; 16 cm 28/9/72 60.0 .42 p o l i s h e d 4/10/72 p o l i s h e d o f f m e t h a l l i b u r e 26/10/72 62.7 .77 p o l i s h e d 1/12/72 60.9 2.88 p o l i s h e d 28/12/72 60.9 4.23 p o l i s h e d 31/1/73 59.5 .13 p o l i s h e d 28/2/73 a n t l e r s dropped .cont'd Table XIV-A cont'd Date . Body wt. Testosterone (kg) (ng/ml) 2/3/73 55.5 .10 5/4/73 51.8 .14 6/4/73 2/5/73 52.7 .37 31/5/73 55.5 .18 28/6/73 58.2 .07 17/7/73 60.0 241 A n t l e r s Experimental Treatment scabs p e d i c l e s healed p e d i c l e s m e t h a l l i b u r e ; healed 1.25 mg/kg/day .5 cm 1.0 cm 1.5 cm 2 cm 242 TABLE XV-A. Seasonal changes in body weight, serum testosterone and the antler growth cycle of captive black-tailed deer under var-i ious experimental treatments. Deer No. B3. Born: June, 1070 Date Body wt. (kg) Testosterone (ng/ml) Antlers Experimental Treatment 22/3/72 4/4/72 12/4/72 28/4/72 29/5/72 1/6/72 23/6/72 27/6/72 27/7/72 3/8/72 4/8/72 23/8/72 24/8/72 19/9/72 27/9/72 13/10/72 27/10/72 24/11/72 27/12/72 26/1/73 56.4 3.90 60.0 62.3 57.3 60.9 60.9 60.0 56.8 53.6 49.5 45,9 .42 .24 15.28 ,23 12.49 .32 1.81 2.40 1.34 .53 polished spikes; 10 cm polished antlers dropped pedicles healed; 1 cm new growth; 2 cm 2 cm velvet shedding; 2 cm polished polished polished polished polished polished antlers dropped scabs pedicles healed pedicles healed pedicles healed pedicles healed pedicles healed outdoor control methallibure; 1.25 mg/kg/day PMS:3X933 IU/wk terminated PMS HCG:3X933 IU/wk terminated HCG off methallibure 243 TABLE XVI-A. Seasonal changes i n body weight, serum t e s t o s t e r o n e and the a n t l e r growth c y c l e o f c a p t i v e b l a c k - t a i l e d deer under var-ious experimental treatments. Deer No. C l . Born: June, 1?7'1. Date Body wt. (kg) Testosterone (ng/ml) A n t l e r s 5/7/71 7.7 .42 p e d i c l e s developing 29/7/71 11.4 .40 p e d i c l e s ; .5 cm 10/8/71 12.7 .5 cm 27/8/71 13.6 .34 1.0 cm 24/9/71 20.0 .40 1.5 cm 21/10/71 26.4 .35 2.0 cm 20/11/71 30.9 .29 2.5 cm 22/12/71 35.0 .24 3.0 cm 25/1/72 37.3 .24 3.5 cm 25/2/72 40.9 1.03 3.5 cm 24/3/72 40.0 .03 3.5 cm 21/4/72 43.6 .08 3.5 cm 25/5/72 44.1 .02 3.5 cm 1/6/72 new growth; 4 cm 29/6/72 46.8 .00 6 cm 28/7/72 48.2 .05 10 cm 25/8/72 52,3 .96 15 cm 29/9/72 59.1 1.71 15 cm 10/10/72 15 cm 11/10/72 v e l v e t shedding 26/10/72 60.9 1.19 p o l i s h e d 30/11/72 63.6 7.74 p o l i s h e d 28/12/72 58.2 .98 p o l i s h e d 29/1/73 59.1 .99 p o l i s h e d Experimental Treatment outdoor c o n t r o l m e t h a l l i b u r e ; 1.0 mg/kg/day o f f m e t h a l l i b u r e .cont'd 244 Table XVI-A cont'd Date Body wt. Ckg) Testosterone ng/ml) A n t l e r s Experimental Treatment 2/3/73 9/3/73 5/4/73 6/4/73 2/5/73 31/5/73 1/6/73 28/6/73 29/6/73 11/7/73 31/7/73 13/8/73 14/8/73 24/8/73 11/9/73 18/3/74 54.5 58.2 60.0 62.2 63.8 66.4 69.1 .18 .08 .16 .22 .11 .18 .40 . p o l i s h e d a n t l e r s dropped p e d i c l e s healed p e d i c l e s healed p e d i c l e s healed new growth: 1.5 cm 1, 2 2 2 2, 6 6 6 5 cm cm cm cm 5 cm cm cm cm v e l v e t shedding; 8 cm a n t l e r s dropped m e t h a l l i b u r e ; 1.25 mg/kg/day HCG:3X933 IU/wk terminated HCG DHEA*;3X33 mg/wk DHEA :3X66 mg/wk terminated DHEA o f f m e t h a l l i b u r e * dehydroepiandrosterone 245 TABLE XVTI-A. Seasonal changes i n body weight, serum t e s t o s t e r o n e and the a n t l e r growth c y c l e o f c a p t i v e b l a c k - t a i l e d deer under v a r -ious experimental treatments. Deer No. C2. Born: June, ^971 Date Body wt, (kg) Testosterone (ng/ml) A n t l e r s Experimental Treatment 8/7/71 5.9 p e d i c l e s .5 cm 19/7/71 6.8 .5 cm 29/7/71 , 8.2 .39 .5 cm 10/8/71 10.0 1.0 cm 28/8/71 11.8 .60 1.5 cm 24/9/71 16.4 .55 2.0 cm 21/10/71 21.8 .24 2.5 cm 20/11/71 23.6 1.09 2.5 cm 10/12/71 v e l v e t shedding; 2.5 cm 23/12/71 28.2 .98 p e d i c l e s p o l i s h e d 27/1/72 29.5 .41 p o l i s h e d 25/2/72 30.9 .35 p o l i s h e d 10/3/72 29.1 buttons dropped 24/3/72 26.8 .09 scabs 21/4/72 30.5 .51 p e d i c l e s healed 5/5/72 new growth; 1 cm 25/5/72 35.0 .21 3 cm 29/6/72 38.6 .21 11 cm 28/7/72 42.7 1.53 23 cm 18/8/72 v e l v e t shedding; 23 cm 25/8/72 45.9 2.56 p o l i s h e d 29/9/72 55.5 1.06 p o l i s h e d 26/10/72 57.7 5.86 p o l i s h e d 1/12/72 50.9 8.96 p o l i s h e d .. . .cont'd 246 Table XVII -A cont'd Date Body wt. (kg) Testosterone ng/ml A n t l e r s Experimental Treatment 28/12/72 50.9 6.90 p o l i s h e d 29/1/73 50.9 6.05 p o l i s h e d 2/3/73 50.0 .24 p o l i s h e d 16/3/73 a n t l e r s dropped 5/4/73 47.7 .40 p e d i c l e s healed 6/4/73 p e d i c l e s healed m e t h a l l i b u r e ; 1.25 mg/kg/day 2/5/73 44.1 .34 .5 cm 31/5/73 46.8 .29 2.0 cm 1/6/73 2 cm HCG:3X933 IU/wk 28/6/73 50.0 .10 3 cm 29/6/73 3 cm terminated HCG 11/7/73 4 cm ASD:*3X17 mg/wk 31/7/73 51.8 .19 9 cm 1/8/73 9 cm ASD: 3X33 mg/wk 13/8/73 9 cm terminated ASD 14/8/73 55.0 .48 9 cm 23/8/73 9 cm o f f m e t h a l l i b u r e 31/8/73 v e l v e t shedding; 9 cm 11/9/73 p o l i s h e d 10/3/74 a n t l e r s dropped * androstenedione 247 TABLE XVIII-A. Seasonal changes in body weight, serum testosterone and the antler growth cycle of captive black-tailed deer under var-ious experimental treatments. Deer No. C3. Born:July 4,1971 Date Body wt. (kg) Testosterone (ng/ml) Antlers Experimental Treatment 4/7/71 6/7/71 29/7/71 10/8/71 27/8/71 2.3 5.0 7.3 9.1 24/9/71 14.1 1/10/71 21/10/71 21.8 20/11/71 27.7 22/12/71 31.8 25/1/72 30.0 25/2/72 31.8 23/3/72 31.8 24/4/72 34.5 26/5/72 36.4 1/6/72 28/6/72 40.0 27/7/72 42.7 23/8/72 24/8/72 50.0 6/9/72 16/9/72 28/9/72 53.2 ,17 .21 11 ,13 ,14 ,27 .16 ,13 .16 .17 .22 .17 .20 .23 .74 none none none pedicles developing pedicles developing pedicles developing pedicles .5 cm born in captivity methallibure; 1.0 mg/kg/day methallibure; 1.25 mg/kg/day .5 cm 1.0 cm 1.0 cm 1 cm 1 cm 1 cm 1 cm 1 cm i 1 cm new growth; 2.5 cm 4 cm 5 cm 5 cm velvet dying velvet shedding; 5 cm HCG:3X933 IU/wk terminated HCG 3.19 polished . cont'd 248 Table XVIII-A cont'd Date Body wt. (kg) Testosterone (ng/ml) A n t l e r s Experimental Treatment 27/10/72 30/11/72 27/12/72 18/1/73 55.5 58.2 55.5 26/1/73 51.8 3/3/73 6/3/73 43.6 29/3/73 42.7 6/4/73 9/4/73 1/5/73 30/5/73 27/6/73 44.5 47.7 52.3 51.8 11/7/73 30/7/73 31/7/73 1/8/73 13/8/73 14/8/73 54.5 23/8/73 7/10/73 1.24 1.60 1.51 .92 .46 .28 .14 .19 .12 ,19 1.76 p o l i s h e d p o l i s h e d p o l i s h e d a n t l e r s dropped scabs p e d i c l e s h e a l i n g p e d i c l e s h e a l i n g p e d i c l e s healed new growth; 10 cm 1.0 cm 1.5 cm 2.5 cm 9 cm 9 cm 9 cm 10 cm 10 cm 10 cm v e l v e t shedding p o l i s h e d a n t l e r s dropped o f f m e t h a l l i b u r e m e t h a l l i b u r e ; 1.25 mg/kg/day m e t h a l l i b u r e ; 1.50 mg/kg/day 19-nortestoster-one;3X17 mg/wk terminated nor-tes t o s t e r o n e t e s t o s t e r o n e ; 3X33 mg/wk terminated t e s t o s t e r o n e o f f m e t h a l l i b u r e 249 TABLE XIX-A. Seasonal changes i n body weight, serum t e s t o s t e r o n e and the a n t l e r growth c y c l e o f c a p t i v e b l a c k - t a i l e d deer under var-ious experimental treatments. Deer No. C5. Born: June, 1971 Date Body wt. (kg) Testosterone (ng/ml) A n t l e r s Experimental Treatment 25/12/71 28/12/71 25.5 28/1/72 24/2/72 10/3/72 24/3/72 27/3/72 4/4/72 28/4/72 26/5/72 1/6/72 28/6/72 27/7/72 23/8/72 24/8/72 27/9/72 27/10/72 30/11/72 19/12/72 26.4 28.6 30.5 32.3 34.5 38.6 40.9 45.9 47.7 49.1 48.6 1.79 .62 .47 .40 ,17 .17 ,20 .23 .15 .12 .64 .34 .33 v e l v e t shedding; 2.5 cm p e d i c l e s p o l i s h e d p o l i s h e d p o l i s h e d 1. button shed 1. p e d i c l e h e a l i n g r. button shed r. p e d i c l e h e a l i n g new growth; 2 cm 5 cm 6 cm 15 cm 22 cm 25 cm 25 cm v e l v e t dying v e l v e t dead 25 cm 25 cm outdoor c o n t r o l m e t h a l l i b u r e ; 1.25 mg/kg/day HCG:3X933 IU/wk terminated HCG 250 TABLE XXA. Seasonal changes in body weight, serum testosterone and the antler growth cycle of captive black-tailed deer under var-ious experimental treatmentr. Deer No. C6. Bom: June, 1971. Date Body wt. Testosterone Antlers Experimental (kg) (ng/ml) Treatment 28/12/71 29.5 .38 pedicles; 2.5 cm outdoor control 28/1/72 31.8 .83 2.5 cm 24/2/72 35.0 .46 2.5 cm 22/3/72 36.4 .21 2.5 cm 4/4/72 28/4/72 37.3 41.4 .04 2,5 cm 2.5 cm methallibure; 1.25 mg/kg/day 26/5/72 43.6 ,02 2.5 cm 1/6/72 2.5 cm PMS: 3X933 IU/wk 27/6/72 47.3 .16 2.5 cm 26/7/72 50.0 .14 2.5 cm 3/8/72 ,2.5 cm terminated PMS 4/8/72 2.5 cm HCG: 3X933 IU/wk 23/8/72 52.7 2.95 velvet dying terminated HCG 27/9/72 56.8 1.79 velvet dead 27/10/72 59.1 .36 2.5 cm 24/11/72 60.5 .19 velvet worn off 5/12/72 bony caps shed 27/12/72 60.5 .55 pedicles healing 26/1/73 58.2 .16 pedicles healed 6/3/73 60.0 .22 pedicles healed ....cont'd 251 Table XXA (cont'd) Date 29/3/73 1/5/73 30/5/73 1/7/73 26/6/73 27/6/73 11/7/73 31/7/73 1/8/73 13/8/73 14/8/73 23/8/73 24/8/73 30/9/73 Body wt. (kg) 56.8 54.5 53.6 56.4 58.2 59.5 61.4 Testosterone (ng/ml) .28 ,12 .24 .46 .39 1.03 A n t l e r s Experimental Treatment p e d i c l e s healed p e d i c l e s healed;. 5 cm .5 cm .5 cm 1.0 cm 1.0 cm 1.0 cm v e l v e t dying v e l v e t dead 1.0 cm 1.0 cm v e l v e t shedding v e l v e t shedding p o l i s h e d FSH:(NIH-P1) 3X6 IU/wk terminated FSH te s t o s t e r o n e ; 3X8 mg/wk te s t o s t e r o n e ; 3X16 mg/wk terminated t e s t o s t e r o n e o f f m e t h a l l i b u r e 252 TABLE V-B. Seasonal changes in testis volume, sperm production, and semen quality of captive black-tailed deer under various experimental conditions (see Table V-A). Deer No. U26. Date Testis Vol. (cm3) Semen Vol. (ml) Sperm Con-centration (106/ml) Motility* % live Remarks 25/5/72 29/6/72 28/7/72 25/8/72 29/9/72 26/10/72 1/12/72 28/12/72 31/1/73 2/3/73 5/4/73 2/5/73 31/5/73 28/6/73 30/7/73 12 14 14 18 20 25 25 12 10 10 8 8 7 8 8 0.1 0.2 1.6 0.6 0.9 1.5 1.6 2.2 1.1 1.5 2.1 1.4 0.1 1.5 0.4 n i l n i l n i l n i l n i l 287 157 21 n i l 86 n i l nil n i l 13 n i l excellent excellent poor n i l 90 94 94 63 pink semen yellow, vis-cous semen pink semen pink semen; 90% abnormal n i l * motility scale: excellent, v. good, good, f a i r , poor, n i l . 253 TABLE VI-B. Seasonal changes i n t e s t i s volume, sperm p r o d u c t i o n , and semen q u a l i t y o f c a p t i v e b l a c k - t a i l e d deer under v a r i o u s experimental c o n d i t i o n s (see Table VI-A). Deer No. W 1. Date T e s t i s V o l . (cm^) Semen V o l . (ml) Sperm Con-c e n t r a t i o n ( 1 0 6 m l ) M o t i l i t y % l i v e Remarks 3/11/71 11/11/71 21/12/71 28/1/72 24/2/72 22/3/72 28/4/72 30/5/72 28/6/72 27/7/72 24/8/72 27/9/72 27/10/72 24/11/72 27/12/72 26/1/73 6/3/73 29/3/73 1/5/73 30/5/73 27/6/73 30/7/73 15 20 20 15 15 12 10 8 .7 10 10 12 15 15 15 15 14 9 7 7 6 5 1.3 .7 1.5 1.4 1.7 1.9 .4 n i l .1 .4 .3 .4 .5 .5 .3 .9 .5 .2 .3 n i l n i l .6 18 28 51 69 52 12 0.4 n i l n i l n i l n i l 1.05 n i l 0.01 n i l 63 230 n i l good v. good f a i r good f a i r poor n i l poor n i l good poor 60 83 30 67 64 50 20 84 45 60% no t a i l couldn't e j a c u l a t e 100% no t a i l y ellow, v i s -cous semen 40% no t a i l couldn't e j a c u l a t e n i l 254 TABLE VTI-B. Seasonal changes in testis volume, sperm production and semen quality of captive black-tailed deer under various experimental conditions (see table VII-A). Deer No. W3 Date Testis Vol. (cm3) Semen Vol. (ml) Sperm Con-centration (106ml) Motility live Remarks 29/11/71 23/12/71 27/1/72 25/2/72 23/3/72 28/4/72 30/5/72 26/6/72 26/7/72 23/8/72 27/10/72 24/11/72 27/12/72 26/1/73 45 35 27 18 15 10 10 10 10 8 27/9/72 15 15 20 13 10 1.0 .8 1.5 1.6 1.1 .9 .1 n i l .05 .7 .3 .5 1.0 .2 .4 325 183 880 365 66 133 46 n i l .85 137 101 338 677 v. good excellent v. good good poor n i l poor n i l f a i r good v. good f a i r 91 88 excellent 90 80 37 7 0 5 0 67 85 93 77 pink semen 95% abnormal, 75% no t a i l 95% no t a i l couldn't ejaculate 95% abnormal, 65% no t a i l 100% abnor-mal, 70% no t a i l 30% no t a i l 255 TABLE VTII-B. Seasonal changes in testis volume, sperm production, and semen quality of captive black-tailed deer under variou; experimental conditions (see Table VIII-A). Deer No. Y9 Date Testis Semen Sperm Con- Motility % Remarks Vol. Vol. centration live Can3) (ml) (106ml) 3/11/71 7 1.2 n i l 20/11/72 10 1.0 n i l 22/12/71 15 0.7 n i l 10/1/72 10 0.7 n i l 27/1/72 8 1.0 n i l 11/2/72 8 0.2 n i l 256 TABLE X-B. Seasonal changes in testis volume, sperm production, and semen quality of captive black-tailed deer under various experimental conditions (see Table X-A). Deer No. Y16 Date Testis Vol. (cm3) Semen Vol. (ml) Sperm Con-centration (106ml) Motility % Remarks live 2/11/72 6 0.3 n i l 20/11/71 12 1.2 n i l 22/12/71 15 0.7 .1 poor 10/1/72 18 1.1 2.1 poor 2 70% no t a i l 27/1/72 12 0.8 n i l 11/2/72 10 0.2 n i l 257 TABLE XI-B. Seasonal changes i n t e s t i s volume, sperm p r o d u c t i o n and semen q u a l i t y o f c a p t i v e b l a c k - t a i l e d deer under various experiraen-t a i c o n d i t i o n s (see Table XI-A). Deer No. Z l Date T e s t i s V o l . (cm 3) Semen V o l . (ml) Sperm Con-c e n t r a t i o n (10 6ml) M o t i l i t y % l i v e Remarks 11/11/71 45 1.8 671 v. good 58 21/12/71 30 1.3 31 f a i r 25 21/1/72 25 1.3 164 v. good 42 24/2/72 15 2.3 724 e x c e l l e n t 85 24/3/72 12 1.2 50 f a i r 27 21/4/72 12 1.2 106 good 50 25/5/72 13 2.8 26 f a i r 65 29/6/72 14 1.3 0.6 poor 70 28/7/72 20 1.0 41 poor 64 pink semen 25/8/72 30 1.1 2.8 poor 75 29/9/72 50 1.7 441 e x c e l l e n t 92 26/10/72 45 3.8 224 e x c e l l e n t 93 30/11/72 30 2.0 386 e x c e l l e n t 93 28/12/72 20 2.8 314 v. good 95 29/1/73 15 1.5 433 e x c e l l e n t 97 2/3/73 13 2.0 86 v. good 77 pink semen 5/4/73 12 1.4 271 v. good 95 2/5/73 8 1.8 116 f a i r 56 pink semen 31/5/73 8 0.4 161 good 80 ^ pink semen 28/6/73 10 1.2 87 f a i r 77 pink semen 30/7/73 10 0.5 103 f a i r 258 TABLE XII-B. Seasonal changes i n t e s t i s volume, sperm pr o d u c t i o n and semen q u a l i t y o f ca p t i v e b l a c k - t a i l e d deer under various experimental c o n d i t i o n s (see Table X I I - A ) . Deer No. Z7". Date T e s t i s V o l . (cm 3) Semen V o l . (ml) Sperm Con-c e n t r a t i o n (10 6ml) M o t i l i t y % l i v e Remarks 3/11/71 30 1.5 190 f a i r 44 11/11/71 30 1.4 582 good 64 21/12/71 30 1.3 148 good 41 21/1/72 20 1.4 107 v. good 50 24/2/72 20 1.5 28 good 40 24/3/72 18 2.2 158 good 80 21/4/72 18 1.4 662 good 40 25/5/72 14 1.0 95 poor 65 35% no t a i l 29/6/72 12 1.1 5.8 n i l 0 70% no t a i l 28/7/72 16 1.0 14 poor 40 25/8/72 30 0.5 0.05 n i l 0 29/9/72 40 1.0 524 e x c e l l e n t 90 26/10/72 35 0.5 243 v. good 92 30/11/72 20 2.0 184 v. good 95 28/12/72 15 1.7 58 f a i r 96 29/1/73 12 1.2 407 e x c e l l e n t 94 2/3/73 12 1.9 378 v. good 97 5/4/73 12 4.5 174 good 82 sperm clump 2/5/73 8 1.0 150 poor 55 i n g 31/5/73 8 3.5 5.3 poor 8 80% no t a i l 28/6/73 8 1.3 8 n i l 0 90% no t a i l 30/7/73 12 1.8 16 n i l 0 259 TABLE XIII-B. Seasonal changes in testis volume, sperm production and semen quality of captive black-tailed deer under various experimental conditions (see Table XIII-A). Deer No. Bl Date Testis Vol. (cm3) Semen Vol. (ml) Sperm Concen- Motility tration (10 6/ml) % Remarks live 1/11/71 20 1.0 422 v. good 67 6/12/71 25 .5 20 poor 94 partial ejaculate 10/1/72 17 .8 523 excellent 80 11/2/72 12 1.0 87 good 72 sperm clumping 10/3/72 12 .7 288 good 39 60% abnor-mal, 30% no t a i l 28/4/72 10 • 8 n i l 31/5/72 7 .2 n i l 27/6/72 7 .08 n i l 27/7/72 8 .8 n i l 23/8/72 7 .05 n i l 28/9/72 7 .5 n i l 26/10/72 8 n i l couldn't ejaculate 1/12/72 13 .4 n i l 28/12/72 13 .3 n i l 31/1/73 8 .4 691 poor 50 2/3/73 6 .3 50 poor 20 80% abnormal 5/4/73 5 .1 n i l 2/5/73 4 1.1 .25 n i l o 31/5/73 3.5 n i l couldn't ej aculate 260 TABLE XIV-B. Seasonal changes i n t e s t i s volume, sperm production and semen q u a l i t y of captive b l a c k - t a i l e d deer under various experimental conditions (see Table XIV-A). Deer No. B2 Date Te s t i s Semen Sperm Con- M o t i l i t y % Remarks Vol. Vol. centration l i v e (cm 3) (ml) (10 6/ml) 29/10/71 15 1.5 119 v. good 6/12/71 17 1.0 172 excellent 91 10/1/72 15 1.0 297 exc e l l e n t 91 11/2/72 10 .7 302 good 72 sperm clump-ing 10/3/72 10 1.4 54 poor 46 sperm clump-ing, 30% no t a i l 2 8 / 4 / 7 2 10 1.9 n i l 31/5/72 . 7 .05 n i l 27/6/72 7 .5 29 n i l 0 100% no t a i l 27/7/72 8 n i l couldn't ejaculate 23/8/72 8 .3 n i l 28/9/72 7 .3 n i l 26/10/72 12 .9 29 poor 40 70% abnormal 45% no t a i l 1/12/72 12 1.1 . 491 excellent 97 28/12/72 8 2.2 381 v. good 95 31/1/73 8 1.0 208 poor 95 2/3/73 6 .5 90 f a i r 67 60% abnormal 5/4/73 5 1.1 .45 n i l 0 100% abnor-mal 2/5/73 4 .4 .40 n i l 0 100% abnor-mal 315/73 3.5 n i l couldn't ejaculate 28/6/73 3 .8 n i l 261 TABLE XV-B. Seasonal changes in testis volume, sperm production and semen quality of captive black-tailed deer under various experimen-tal conditions (see Table XV-A). Deer No. B3 Date Testis V o l . (cm3) Semen Vol. (ml) Sperm Con-centration (106/ml) Motility % live 23/3/72 12 .5 821 excellent 70 28/4/72 10 1.8 109 poor 35 30/5/72 7 .2 4 n i l 0 27/6/72 9 .7 n i l 27/7/72 11 .4 n i l 24/8/72 8 .2 n i l 27/9/72 6 .3 n i l 27/10/72 8 • 2 29 f a i r 90 24/11/72 7 .5 36 f a i r 95 27/12/72 7 n i l 26/1/73 6 n i l Remarks couldn't ejaculate couldn't ejaculate 262 TABLE XVI-B. Seasonal changes in testis volume, sperm production and semen quality of captive black-tailed deer under various experimental conditions (see Tabl e XVI-A). Deer No. Cl Date Testis Semen Sperm Con- Motility % Remarks Vol. (cm3) Vol. centration live (ml) C106/ml) 4/11/71 1 .25 n i l 23/12/71 1.5 n i l 27/1/72 2 n i l 25/2/72 2 n i l 24/3/72 3 n i l 21/4/72 4 .4 n i l 25/5/7] 2 n i l 27/6/72 2 .5 n i l 28/7/72 4 1.2 n i l 25/8/72 3.5 .7 n i l 29/9/72 5 .6 n i l 26/10/72 10 .9 18 f a i r 30/11/72 17 1.8 113 poor 85 yellow, viscous - semen 28/12/72 13 1.0 458 v. good 92 29/1/73 8 .7 357 excel-lent 95 2/3/73 8 2.1 203 f a i r 87 viscous semen 5/4/73 6 .7 121 poor 40 sperm clumping 2/5/73 3.5 1 1.6 10 n i l 0 31/5/73 2.5 n i l couldn't ejaculate 28/6/73 4 n i l couldn't ejaculate 31/7/73 4 1.0 n i l 14/8/73 3 n i l couldn't ejaculate 263 TABLE XVII-B. Seasonal changes i n t e s t i s volume, sperm production and semen q u a l i t y o f c a p t i v e b l a c k - t a i l e d deer under various experimental c o n d i t i o n s (see Table XVII-A). Deer No. C7 Date T e s t i s Semen Sperm Con- M o t i l i t y % Remarks V o l . V o l . c e n t r a t i o n l i v e (cm 3) (ml) (10 6/ml) 4/11/71 3 .2 n i l 23/12/71 7 .8 n i l 27/1/72 8 .1 5.3 25/2/72 10 .5 22 24/3/72 5 n i l 21/4/72 3 .4 n i l 25/5/72 4 .9 n i l 29/6/72 7 .5 n i l 28/7/72 11 .05 n i l 25/8/72 14 .1 6 29/9/72 18 .7 339 26/10/72 22 .8 512 1/12/72 18 1.4 809 28/12/72 10 .8 679 29/1/73 8 1.0 303 2/3/73 8 .6 184 5/4/73 6 .5 51 2/5/73 4 .9 n i l 31/5/73 3.5 n i l 28/6/73 3.5 n i l 31/7/73 3.5 .05 n i l 14/8/73 3 n i l f a i r 50 good 67 couldn't e j a -c u l a t e poor 75 v. good 75 v. good 88 e x c e l l e n t 96 sperm clumping e x c e l l e n t 91 v. good 90 f a i r 91 poor 9 70% abnormal, 40% no t a i l ; pink semen couldn't e j a c u l a t e couldn't e j a c u l a t e 264 TABLE XVIII-B. Seasonal changes in testis volume, sperm production and semen quality of captive black-tailed deer under various experimental conditions (see Table XVIII-A). Deer No. C3 Date Testis Semen Sperm Con- Motility % Remarks Vol. Vol. centration , live (cm3) (ml) (106/ml) 28/4/72 2 n i l 25/5/72 2.5 n i l 28/6/72 3 n i l 27/7/72 3.5 .05 n i l 24/8/72 6 .1 n i l 28/9/72 9 2.0 n i l 27/10/72 12 .3 8 30/11/72 10 .4 89 27/12/72 8 .9 26 26/1/73 5 .2 2.5 6/3/73 3.5 .8 n i l 29/3/73 4.5 n i l 1/5/73 4 .6 n i l 30/5/73 4 n i l 27/6/73 2.5 2.0 n i l 31/7/73 3 .05 n i l 14/8/73 2.5 .1 n i l couldn't eja-culate couldn't eja-culate couldn't eja-culate poor poor 84 70% abnormal poor 30 yellow, viscous semen n i l 0 couldn't eja-culate couldn't eja-culate 265 TABLE XIX-B. Seasonal changes in testis volume, sperm production and semen quality of captive black-tailed deer under various experimental conditions (sse Table XIX-A). Deer No. C5 Date Testis Semen Sperm Con- Motility % Remarks Vol. Vol. centration live Ccm3) (ml) (106/ml) 28/12/71 5 .3 n i l 28/1/72 5 .9 n i l 24/2/72 5 .5 n i l 24/3/72 4 1.2 n i l 28/4/72 3.5 .3 n i l 25/5/72 3.5 .3 n i l 28/6/72 4 • 3 ni l 27/7/72 4 .4 n i l 24/8/72 2.5 .7 n i l 28/9/72 2.5 .4 n i l 27/10/72 2.5 .2 n i l 30/11/72 2.5 .2 n i l 266 TABLE XX-B. Seasonal changes i n testis volume, sperm production and semen quality of captive black-tailed deer under various experimen-t a l conditions (see Table XX-A). Deer No. C6 Date Testis Semen Sperm Con- Motility % Remarks Vol. Vol. centration live (cm3) (ml) (106/ml) 28/12/71 4 .3 n i l 28/1/72 4 .1 n i l 24/2/72 4 .5 n i l 22/3/72 4 .3 n i l 28/4/72 4 .3 n i l 25/5/72 4 .4 n i l 28/6/72 4 .3 n i l 27/7/72 3.5 .4 n i l 23/8/72 3.5 .2 n i l 27/9/72 3.5 .4 n i l 27/10/72 6 .3 n i l 24/11/72 10 .5 36 27/12/72 3.5 .4 n i l 26/1/73 3.5 .2 n i l 6/3/73 3.5 .35 n i l 29/3/73 3.5 .4 n i l 1/5/73 3 .5 n i l 30/5/73 2 .5 n i l 27/6/73 2 .1 n i l 31/7/73 2.5 .6 n i l 14/8/73 2.5 n i l poor 75 couldn't ejaculate 267 Key to the abbreviations used in a l l the tables in the C series SGOT serum glutamic-oxaloacetic transaminase LDH l a c t i c dehydrogenase Alk Phos alkaline phosphatase T B i l i total b i l i r u b i n Creat creatinine BUN blood urea nitrogen Choi cholesterol Inor Phos inorganic phosphorus Ca calcium Alb albumin TT^ total thyroxin (Murphy-Patee) Resin T^U resin triiodothyronine uptake (Clark) F Index "free" thyroxin index Table VC. Serum enzymes, metabolic wastes, cholesterol, phosphorus, calcium, proteins, and thyroxin of captive black-tailed deer under various experimental conditions (see table VA) • Deer number U26 Date SGOT IU/l LDH IU/l Alk Phos IU/l T B i l l mg io Creat mg i Uric' Acid mg $ BUN mg i Choi mg i Inor Phos mg % Ca mg % Alb g i Total TT^ Protein ug$ g $ Resin U $ F Tl| Index 25. 5.72 1^8 1+12 9U -3 2.0 0 30 73 8.U 9-0 .6 6.7 9-2 36.9 3-39 28. 7.72 112 UlO 218 • 3 1.9 0 31 80 7.9 8.2 • 5 6,7 10.7 35. h 3 79 29. 9-72 112 1+90 80 • 3 2.2 0 1+3 87 8.6 9.7 .7 7-2 8.6 36.2 3-11 1.12.72 96 37U 66 • 3 2.9 0 32 51 7.9 9-h .7 7.2 7.2 1+3-1 3.10 31. 1-73 12U 508 1+0 • 3 1.9 0 37 91 8.3 8.8 .6 7.1 9.U 35.8 3-37 5 U.73 86 388 38 • 3 1.9 0 32 92 8.1+ 8.8 .6 6.9 7.3 36.2 2.6U 31- 5-73 95 1+1+8 95 • 3 1.6 0 35 67 10.2 8.6 • 7 6.7 7.7 28.5 2.19 Table VIC. Serum enzymes, metabolic wastes, cholesterol, phosphorus, calcium, proteins and thyroxin in captive black-tailed deer under various experimental conditions (see table VIA). Deer number Wl Date SGOT LDH Alk T B i l l Creat Uric BUN Choi Inor Ca Alb Total TT1+ Resin F IU/l IU/l Phos mg io mg $ Acid mg f0 mg % Phos mg % g % Protein ug % T 3 Index IU/l mg i mgi g i U i 6. 2.73 53 3U9 59 -3 2.U 0 ko 79 7.0 9.6 .9 7.6 30. 5-73 89 372 77 .h 1.9 0.1 32 60 10.3 9-6 .9 6.9 7.5 35.8 2.69 Table ViIC. Serum enzymes, metabolic wastes, cholesterol, phosphorus, calcium, proteins and thyroxin in captive black-tailed deer under various experimental conditions (see table VIIIA) . Deer number Y9 Date SGOT IU/l LDH IU/l Alk Phos IU/l 7. 3.70 7h 23U 36 8. 5-70 100 327 51 10. 7.70 9h 259 139 11. 9.70 10U 337 55 13.11.70 93 268 37 22. 1.71 60 23U 20 27. 3-71 85 278 29 29- 5-71 ikl 366 61 30. 7.71 82 317 3h 25. 9-71 79 351 19 11.11.71 78 3kk 23 27. 1.72 79 311 16 T B i l i Creat Uric BUN mg % mg io Acid mg % mg % .3 l . l 0 26 .2 l - l 0 25 .2 l . l 0 23 .3 1.2 0 30 .k 2.8 0 30 .2 1.2 0 28 • 3 l.k 0 28 • 3 1.2 0 23 • 3 1.5 0 18 • 3 1.^  0 19 • 3 1.3 0 2k .k 1.7 0 18 Choi Inor Ca Alb mg% Phos mg % mg % g % 55 7.3 7.9 .6 52 l.k 7.7 .6 55 8.0 7.5 .6 66 8.6 8.2 .6 kg 7.9 9-3 .7 ki 6.0 5.8 .5 ki 8.6 l.k .6 52 8 .U 1.9 .6 57 8.1 7.6 .5 51 8.k 8.5 .6 60 8.5 8.2 .6 5k 9-0 8 .U .6 Total TT U Resin F Tl^ Protein ug i T3 Index g i 5.8 11.5 28.5 3.28 6.2 12.2 28.5 3.U8 5.9 10. k 29.2 3.0U 6.7 10.6 33.8 3.58 7.3 8.k ki.i 3.*+5 U.7 12.2 35.8 k.31 5.7 11.9 28.8 3M 5-9 9.8 29.6 2.90 5.6 7.5 30.8 2.31 6.7 7.2 32.7 2.35 6.8 6.1 32.7 1.99 6.6 9.1 33.5 3.05 Table VIIIC. Serum enzymes, metabolic wastes, cholesterol, phosphorus, calcium, proteins, and thyroxin i n . captive black-tailed deer under various experimental conditions (see table IXA).. Deer number Yl6 Date SGOT IU/l LDH IU/l Alk Phos IU/l T B i l l mg % Creat mg % Uric Acid mg i BUN mg % Choi mg i Inor Phos mg % Ca mg % Alb g i Total Protein g i TTli ug % Resin u i F TU Index 6. 3-70 57 336 27 .3 1.1+ 0 29 63 6.8 9-6 .7 6.7 10.2 31.9 3.25 7. 5-70 81 3^1 58 • 3 1.3 0 25 60 7-5 9-3 .6 6.2 10.8 32.3 3.»+9 8. 7.70 71 352 103 .5 1.3 0 27 65 8.1 8.9 .7 6.2 10.7 33.5 3.58 9- 9.70 80 1+1+8 1+1 .5 1.1+ 0 25 77 8.1+ 9.6 .8 7.1 12.5 32.7 1+.09 11.11.70 99 335 39 .6 2.9 0 22 71 7.5 9 A .8 7.1 10.2 i+o.8 U.16 21. 1.71 56 295 52 •6 2.2 0 29 79 6.2 9-5 • 9 6.5 11.1 38.1 1+.23 k. 3-71 63 319 39 .6 1.9 0 25 57 5.9 9-7 .8 6.3 11.5 31*.2 3-93 28. 5.71 101+ 1+1+1+ 121 •3 1.1+ 0 29 7U 7.5 9.2 .7 6.3 11.1 31.9 3-5lr 29- 7-71 63 331 39 .5 1.6 0 22 62 8.2 7.5 .7 5.9 9.9 30.1+ 3.01 2k. 9.71 66 3^8 25 • 5 1.6 0 18 63 8.3 8.1 .7 6.0 10.1+ 31.2 3.2U 20.11.71 262 630+ 1+1 1.5 2.7 0 10 81 8.1 8.6 .1+ 6.3 6.0 1+2.7 2.56 27. 1.72 123 630+ 23 .8 2.9 0 20 116 8.7 8.1 .7 5-8 8.6 38.5 3.31 Table IXC. Serum enzymes, metabolic wastes, cholesterol, phosphorus, calcium, proteins, and thyroxin in captive black-tailed deer under various experimental conditions (see table XIA). Deer number Z l Date SGOT IU/l LDH IU/l Alk Phos IU/l 2k. 3-72 21 U89 ko 25. 5.72 193 508 26 28. 7.72 13k k92 368 29. 9.72 130 519 kl 30.1.72 13 k66 kl 29. 1.73 162 1+85 3k 31. 5.73 159 602 25k T B i l i Creat Uric BUN mg % mg % Acid mg % mg $ .3 1.+ 0 3 .3 !-3 0 2k .2 1.3 0 36 • 3 2.9 0 kl .2 2.6 0 k2 • 3 2.0 0 kl .2 1.+ 0.1 39 Choi Inor Ca Alb mg io Phos mg % g % mg % 63 l.k 9-7 .6 81 6.6 9-3 .6 90 7.7 8.1+ .6 78 7.8 9.5 .7 67 8.0 9-7 .8 67 6.9 9-9 .6 94 8.7 9-6 1.0 .Total Protein g $ TT1+ ug io Resin u % F Tk Index 6.8 1.1+ 35.0 3.9 6.7 10.7 36.5 3.91 6.6 1.0 31+.6 3.81 7.1+ 9-7 37.7 3.6 7.0 10.5 U0.8 1+.28 6.9 9-3 35.8 3.3 6.9 9.8 3I+.6 3-39 Table XC. Serum enzyes, metabolic wastes, cholesterol, phosphorus, calcium, proteins, and thyroxin in captive b lack-ta i led deer under various experimental conditions (see table XIIA) . Deer number Z7 Date SGOT I U / l LDH I U / l Alk Phos I U / l T B i l i mg i Creat mg % Uric Acid mg % BUN mg i Choi mg i Inor Phos mg % Ca mg $ Alb g i Total Protein g i TTii ug i Resin u i F T U Index 2k. 3.72 98 317 95 .2 1.6 0 36 81 6.5 9.3 .6 7.2 17.8 3^.2 6.09 25. 5.72 193 392 125 .2 '1.3 0 27 85 7-3 8.8 .6 6.6 15.8 31+.6 5.^7 28.-7-72 103 392 lUo .2 1.3 0 3h 90 7.3 8.9 .6 6.8 11+.6 33.5 I+.89 29. 9-72 h9 230 50 .2 2.U 0 28 52 7.k 8.1 .6 7.3 10.0 1+0.0 1+.00 30.11.72 hi 253 85 .2 1.8 0 33 56 7.0 7.9 .5 l.h 8.6 1+1.5 3-57 29. 1-73 82 329 100 .2 1.7 0 38 61 7.2 7.8 .1+ 6.7 7.8 1+2.7 3-33 5- h.73 129 355 78 .2 1.8 0 l+l 89 7.6 9.8 .7 7.5 10.2 1+1.1 1+.19 31. 5.73 216 399 106 .2 l.U 0 33 81+ 8.8 7.7 .6 6.1+ 11.7 36.2 1+.21+ Table XIC. Serum enzymes, metabolic wastes, cholesterol, phosphorus, calcium, proteins, and thyroxin of captive black-tailed deer under various experimental conditions (see table XIVA). Deer number B2 Date SGOT LDH Alk T B i l l Creat Uric BUN Choi Inor Ca Alb Total TT. Resin F T^ IU/l IU/l Phos mg % mg % Acid mg % mg % Phos mg % g % Protein ug % To Index IU/l mg $ mg % g io XS i . 31. 5-73 107 U82 90 .k 1.6 .1 35 6 l 9-6 8.7 .7 7.0 10.2 36.9 3-76 Table XIIC. Serum enzymes, metabolic wastes, cholesterol, phosphorus, calcium, proteins, and thyroxin in captive black-tailed deer under various experimental conditions (see table XIXA). Deer number C5 Date SGOT IU/l LDH IU/l Alk Phos IU/l T ] mg 28. 1.72 172 556 52 .k 2h. 3-72 12k kkl 108 .2 26. 5.72 122 1+U3 71 .2 28. 6.72 153 kjk 99 .2 27. 7.72 lH6 1+31 73 .2 2k. 8.72 120 kl2 59 .2 28. 9.72 105 U8l 50 .2 30.11.72 136 k5h 25 .1+ Creat Uric BUN Choi Inor Ca mg $ Acid mg i mg i mg $ Phos mg % mg $ 1.2 0 36 62 9.8 8.2 1.1 0 2k 57 9-1 9.2 1.2 0 23 51 9 A 8.6 1.2 0 28 5U 9-7 8.5 1.3 0 26 56 9-6 8.3 1.3 0 29 *9 9-5 8.2 l.k 0 26 62 9-5 8.7 1.9 0 12 57 9.0 9.1 Alb g i Total Protein g 1o TT^ ug i Resin T3 .IT* F TLi Index .3 5-5 11.1 35.8 3-97 ,5 5.6 10.7 36.2 3.87 .6 5-7 8.3 33-5 2.78 .6 5.6 8.8 37-3 3.28 .6 5.7 8.6 35.8 3.08 .6 5-7 8.3 35.0 2.91 .7 6.2 7.9 35-0 2.77 .h 6.0 9.0 32.7 2.9*+ Table XTI'IC. Serum enzymes, metabolic wastes, cholesterol, phosphorus, calcium, proteins, and thyroxin of captive black-tailed deer under various experimental conditions (see table XXA) . Deer number C6 Date SGOT LDH Alk T B i l i Creat Uric BUN Choi Inor Ca Alb Total TT^ Resin F T^ IU/l IU/l Phos mg ii mg io Acid mg io mg % Phos mg % g % Protein ug % T~ Index IU/l mg i mg $ g % U% 30. 5.73 173 557 77 -3 1-9 .1 39 58 10.5 9-5 -9 7.7 5.h 28.1 1.52 277 Table V-D. Uri n e output, s p e c i f i c g r a v i t y , i n o r g a n i c i o n co n c e n t r a t i o n and o s m o l a r i t y of normal and m e t h a l l i b u r e - t r e a t e d c a o t i v e b l a c k - t a i l e d deer (see t a b l e IXA). Deer number Y l 6 Date Volume S p e c i f i c Na K C l Osmolarity ( l i t e r s ) g r a v i t y (mE/l) (mE/l) (mE/l) ( m i l l i -osmols/l) 18. 5-71 1.990 1.030 176.0 165 166 19- 5-71 1.77U 1.029 120.0 185 11+8 20. 5.71 1.7k2 1.033 120.5 175 li+2 21. 5.71 1.370 1.029 118.0 150 11+7 22-2k. 5.71 5-315 1.030 136.0 158 152 1309 25. 5-71 1.33*+ 1.025 106.5 131 132 979 26. 5.71 2.027 1.026 126.5 11+1+ ll+3 1331+ 27. 5-71 1.568 1.026 108.0 137 129 1012 28. 5-71 2.395 1.030 130.5 217 138 998 29- 5.71 1.651 1.031 ltr0.5 207 151 107I+ 30-31. 5.71 3.72k 1.032 117.5 199 li+2 1003 1. 6.71 1.1+00 1.030 ll+l.O 195 1U9 10U5 1. 6.71 J started methallibure treatment 2. 6.71 2.2k9 1.026 103.0 11+1+ 112 830 3- 6.71 k.371 1.015 5k.0 69 1+6 l+l+l 1+. 6.71 k.026 I.OO8 Ul.5 52 38 369 5. 6.71 3.525 1.020 78.0 93 81 601+ 6. 6.71 3.911 1.016 76.5 89 63 569 7. 6.71 3.^ 26 1.017 68.0 87 68 616 8. 6.71 2.867 1.020 81+.5 108 92 671+ 9- 6.71 U.519 1.012 61 5 76 5l+ 1+72 278 Table V-D continued Date Volume Specific Na K ( l i t e r s ) gravity (mE/l) (mE/l) Cl Osmolarity (mE/l) ( m i l l i -osmols/l) 10. 6.71 1+.191 1.012 1+9.0 65 51 1+1+5 11. 6.71 3.786 1.012 52.0 62 5h 1+92 12-13. 6.71 8.828 1.011 50.5 61 51 696 Ik. 6.71 3.990 1.013 U8.5 60 1+9 502 15. 6.71 h.3lk 1.010 kk.7 57 kk 616 16. 6.71 k.5hk 1.012 kk.3 60 kk k76 17-18. 6.71 9.152 1.008 M+.5 51 38 506 19-20. 6.71 9-355 1.011 1+0.3 51 1+0 580 21-23. 6.71 12.577 1.011 M+.5 55 1+0 1+80 2k-26. 6.71 12.380 1.012 50.3 61 577 27-29. 6.71 11.103 1.013 52.0 68 50 71+6 30. 6.71 3.029 1.021 53.0 56 1+2 1+21 1-2. 7.71 9.029 1.013 68.5 82 60 612 3-5. 7.71 15-800 1.011 1+1+.3 56 37 539 6-8. 7.71 13.708 1.011 U0.5 59 36 5^3 279 Table VI-D. Urine output, specific gravity, inorganic ion concentration and osmolarity of normal and methallibure-trated captive black-tailed deer (see table XA). Deer number Y23 (control) Date Volume Specific Na K Cl Osmolarity ( l i t e r s ) gravity (mE/l) (mE/l) (mE/l) ( m i l l i -osmols/l) 18. 5.71 1.1+16 1.035 173.0 175 191+ 19- 5.71 1.208 1.038 127.0 225 169 20. 5.71 1.336 1.035 11+8.0 205 196 21, 5.71 1.697 1.036 155.0 251 180 117U 22-2*+. 5.71 1+.576 1.035 157-5 209 18U 1328 25. 5.71 1.115 1.033 173.0 225 176 1168 26. 5.71 1.852 1.032 153.0 236 161+ 1187 27. 5.71 1.208 1.035 151.0 236 166 1275 28. 5.71 2.211 1.032 ll+l.O 211+ 151+ 1066 29. 5-71 1.551+ I.03I+ 1U0.0 215 138 1070 30-31. 5.71 3.335 1.030 139.5 160 ll+l 1060 1. 6.71' 1.206 1.035 151+.0 209 182 1210 2. 6.71 1.950 1.031 131.0 193 159 1321 3. 6.71 3.01+7 1.022 81+.0 131 102 821+ u . 6.71 2.161+ 1.023 89.5 127 108 8I+5 5. 6.71 2.719 1.031 121.5 159 138 1065 6. 6.71 2.082 1.028 109.0 ll+5 132 91+0 7. 6.71 2.313 1.032 133.0 166 ll+9 11+23 8. 6.71 2.221+ 1.028 103.0 11+8 120 970 9- 6.71 2.107 1.030 125.0 163 ll+2 1105 10. 6.71 2.186 1.026 lll+.O 130 132 982 280 Table VI-D continued Date Volume Specific Na (l i t e r s ) gravity (mE/l) K (mE/l) Cl (mE/l) Osmolarity ( m i l l i -osmols/l) 11. 6.71 1.712 1.031 114.0 140 155 1177 12-13. 6.71 4.725 1.027 106.5 120 138 1050 Ik. 6.71 2.084 1.030 120.0 139 138 1548 15- 6.71 2.337 1.024 93.0 118 111 980 16. 6.71 1.953 1.031 137.5 151 145 1298 17-18. 6.71 3.910 1.030 118.0 131 121 1260 19-20. 6.71 4.063 1.031 149.0 140 161 1280 21-23. 6.71 4.906 1.032 119.0 144 138 1570 24-26. 6.71 4.592 1.034 106.5 162 156 1342 27-29. 6.71 2.478 1.042 94.5 184 211 1710 30. 6.71 .968 i.oko 102.5 176 210 1660 1-2. 7-71. 3.228 1.037 157-5 164 211 1455 3-5. 7.71 4.320 1.035 153.5 154 168 1450 6-8. 7.71 4.906 1.029 131.0 140 144 1126 Key to the abbreviations used in table XXI. SGOT serum glutamic-oxaloacetic transaminase LDH lactic dehydrogenase Alk Phos alkaline phosphatase T B i l i total bilirubin Creat creatinine BUN blood urea nitrogen Choi cholesterol Inor Phos inorganic phosphorus Ca calcium Alb albumin TT. 4 total thyroxin (Murphy-Patee) Resin TjU resin triiodothyronine uptake (Clark) F T 4 Index "free" thyroxin index Table XXI. Serum enzymes, metabolic wastes, cholesterol, phosphorus, calcium, proteins and thyroxin of wild black-tailed deer on Vancouver Island. Animal SGOT LDH Alk T B i l l Creat Uric BUN No.Sex Age IU/l IU/l Phos IU/l mg % mg $ Acid mg $ mg Mar. 25-29 8 M 1.8 263+ 537 35 0 1.9 .6 9 10 M 1.8 263+ 20k 33 0 1.1+ .6 1+ 3^ F 1.8 93 303 18 0 l.k • 3 18 38 F* 2.8 67 300 31 0 1.6 •3 2 May 2U-25 26 M 2.0 129 506 81 0 .8 .8 19 27 M 2.0 263+ 632+ 85 0.1 1.2 0 27 28 M 3-0 263+ 625 117 0 .9 0 19 July 23 12 M 3-2 65 632+ 102 0 1.1 .5 12 13 M 2.2 263+ 632+ 161 0 •1.1 .6 11 Ik M 1.2 7h 632+ 66 0 .9 .8 11+ Choi mg % Inor Phos mg i Ca mg # Alb g i Total Protein g i ^h. ug % Resin T 3 U F T^ Indes 51 8.6 8.1 .3 1+.2 5.9 39-6 2.3h 19 8.6 6.8 .3 2.7 3.9 1+0.8 1.59 3h 8.1+ 7.3 .1+ 1+.1 7.7 37.7 2.90 28 8.3 7.5 .3 1+.6 l+.U 38.1 1.68 55 9.3 7.1* .5 h.<? 15.3 36.9 1.96 76 8.7 8.3 .6 7.6 7-3 1+0.1+ 2.95 1+8 7.5 6.2 .5 5.6 12.1 35.0 1+.2U 5h 9.0 10.7 .7 6.0 12.1+ 28.8 3.57 1+8 11.0 .6 5.h 9.9 32.7 3.21+ hi 9-8 10.3 .5 5.h 9.7 33-5 3.25 Table XXI continued Animal SGOT No.Sex Age IU/l Sept. 11-12 18 M 4.3 263+ 19 M 1.3 263+ 20 M 5-3 263+ 16 M .3 183 Nov. 16-30 2 M 1.5 85 4 M 5-5 263+ 5 M 3.5 183 6 M 6.5 263+ 1 M .5 214 7 M .5 75 12 F 3-5 240 15 F 4.5 263+ LDH Alk T B i l l Creat Uric BUT IU/l Phos mg % mg % Acid mg IU/l mg i 632+ 68 0 1.4 0 13 632+ 111 0 1.1 0 14 632+ 58 0 1.5 0 9 632+ 163 0 1.0 .2 15 257 27 0 1.7 0 6 632+ „ 32 0 2.4 0 14 472 25 0 2.4 .1 12 632+ 160 0.1 2.8 0 20 328 36 0 .8 0 7 347 78 0 1.3 0 13 588 40 0 1.2 0 8 632+ 23 0 .8 .1 9 Choi Inor mg $ Phos mg % 51 7.1 52 9.0 51 6.9 64 8.1 52 . 7.2 48 7.7 43 8.1 65 8.3 24 5-3 53 7.1 51 7.5 64 5.3 Ca Alb Total mg io g io Prote: g i 9.8 .7 7.2 8.6 .6 6.4 8.7 .6 6.7 8.9 • 5 4.2 9.6 .7 6.3 6.7 .6 5-9 9.3 .4 5.9 7.2 .7 7.0 5.0 .3 2.6 9-9 .7 5.8 8.8 .6 6.5 5.9 .4 h.3 TTit Resin F T. ug % Tg U Index 8.1 36.5 2.96 7.8 36.9 2.88 9-8 35.4 3.47 18.9 34.2 6.46 6.8 38.2 2.60 5.4 45.4 2.45 4.2 4l.l 1.73 3.0 43.5 1.31 8.2 37.3 3.06 8.2 31.9 2.62 7.6 31.2 2.37 7.7 29.2 2.25 Table XXI continued Animal SGOT LDH Alk T B i l i No.Sex Age IU/l IU/l Phos mg % IU/l Jan. 18-21 22 M 3-7 263+ 632+ 32 0.2 25 M 2.7 263+ 632+ 72 0.7 23 M .7 263+ 632+ 97 O.k * pregnant Creat mg lo Uric Acid rng i BUN mg io Choi mg io Inor Phos mg % Ca mg i0 1.7 0 2 71 6.1 9.1* l.U 1.3 0 91 10.5 7.9 1.3 U.2 0 73 8.9 7-5 Alb Total TT^ Resin P T^ g io Protein ug % To U Index si  3 .8 7.0 7.7 32.3 2.U9 .2 6.8 k.3 hO.O 1.72 .5 6.0 5.0 33.8 I.69 K > OO 

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