Copyright is owned by the Author of the thesis. Permission is given for a copy to be downloaded by an individual for the purpose of research and private study only. The thesis may not be reproduced elsewhere without the permission of the Author. Corticosterone responses to stressors and the regulation of the hypothalamic-pituitary-adrenal axis in Japanese quail A thesis presented in partial fulfilment of the requirements for the degree of Doctor of Philosophy in Physiology at Massey University, Palmerston North New Zealand Wei-Hang Chua 2003 III Abstract Corticosterone is the major adrenal glucocorticoid in birds. It is secreted in response to stressors, with plasma concentrations typically rising to a peak within 10 to 15 minutes and then declining over 30 to 60 minutes once the stressor is removed. The increase in corticosterone is thought to help the animal to adjust to the stressor, although corticosterone may also inhibit the reproductive axis. The corticosterone response to a stressor varies between individuals, although the level of the hypothalamic-pituitary-adrenal axis at which this variation arises in birds has not been identified. The goals of this research were to determine in Japanese quail (Coturnix coturnix japonica): (1) the effects of corticosterone and of various stressors on sex steroid secretion; (2) corticosterone responses to 5, 10 or 15 min exposure to manual restraint and 15 min exposure to different stimuli; (3) quantifying individual variation in the corticosterone response; and (4) whether individual variation in the corticosterone response to stimuli used in goal 2 is regulated by differences in pituitary or adrenal sensitivity to corticotropin releasing factor (CRF) and adrenocorticotrophic hormone (ACTH) respectively. A prolonged elevation in plasma corticosterone concentrations can inhibit reproduction. The effects of a short elevation in corticosterone are less understood, so the relationship between corticosterone and sex steroid secretion in quail was characterised by administering 1 .2 mg corticosterone injections. Corticosterone treatment s timulated an increase in plasma corticosterone, while testosterone declined in both corticosterone-treated and control birds. Plasma LH was unaffected by challenge with corticosterone. Collectively, these data suggest that repeated handling associated with frequent blood sampling inhibited testosterone secretion directly at the testis. v Corticosterone responses to a variety of novel stimuli were characterised. Manual restraint for 10 or 15 min caused a significant response whereas restraint for 5 min or less did not lead to an increase in corticosterone 15 minutes from the start of the stressor. Corticosterone responses to 15 min of mechanical restraint varied between birds whereas 15 min of manual restraint elicited a maximal plasma corticosterone response in all birds. Manual and mechanical restraint caused declines in plasma testosterone of a similar magnitude. There was more variation within than between birds in their corticosterone response to mechanical restraint. Despite this, the general pattern of the corticosterone response was repeatable for individuals. Quail with low or high plasma corticosterone responses to 15 minutes of mechanical restraint were injected intrajugularly with a dose of ACTH that stimulated a sub-maximal corticosterone response. The plasma corticosterone response to ACTH did not differ between birds with low or high corticosterone responses to mechanical restraint, indicating that variation in corticosterone responses to restraint did not arise at the level of the adrenal gland. A preliminary study showed that injections of ovine CRF stimulated corticosterone secretion. However, CRF did not consistently stimulate an increase in plasma corticosterone in these birds and hence it was not possible to determine if pituitary responsiveness to CRF differed between birds with low or high corticosterone responses to restraint. These results indicate that variation in the corticosterone response between individual Japanese quail arises above the level of the adrenal gland in the HPA-axis, and may occur at the pituitary gland or due to differences in activation of neural pathways in the brain. �·I�-' Massey University � Institute of Veterinary, Animal and Biomedical Sciences Private Bag 11 222, Palmerston North, New Zealand T: 64 6 350 4525 F: 646350 5714 Institute of Veterinary, Animal and Biomedical Sciences 30th October 2004 CERTIFICA TE OF REGULA TORY COMPLIANCE This is to certify that the research carried out in the Doctoral Thesis entitled: "Corticosterone responses to stressors and the regulation of the hypothalamic-pituitary-adrenal axis in Japanese quail" in the Institute of Veterinary, Animal and Biomedical Sciences at Massey University, New Zealand: (a) is the original work of the candidate, except as indicated by appropriate attribution in the text and/or in the acknowledgements; (b) that the text, excluding appendices/annexes, does not exceed 100,000 words; (c) all the ethical requirements applicable to this study have been complied with as required by Massey University, other organizations and/or committees which had a particular association with this study, and relevant legislation. Candidate's Name: We i-Hang Chua Signature: � Date: /It/IO /2004 Supervisor's Name: John F. Cockrem Signature: AW Date: IX Acknowledgements I would like to thank my supervisor, Dr. John Cockrem, for his support, guidance, encouragement, and the opportunities to travel to Japan, India, Scotland and Antarctica during my studies. Thanks also go to Prof. Tim Parkinson for his constructive criticism and insight during my research. This research was supported by funding from the Institute of Veterinary, Animal and Biomedical Sciences (IVABS) Postgraduate Research Fund. Personal financial support was provided by the Massey University Scholarship Committee. A special thanks to Prof. Hugh Blair and Allain Scott of the IVABS Postgraduate Research Office. Many thanks go to past and present members of the Conservation Endocrinology Research Group; Janis Bridges, Jane Candy, Janie Girling, Guy Hessell, and Raewyn Wheeler for their help and ideas over the years. Thanks also go to numerous students and staff in the Institute of Veterinary, Animal and Biomedical Sciences for their support. I am grateful to Professor Susumu Ishii (Department of Education, Waseda University, Tokyo, Japan) for his kindness and hospitality during my visit to his laboratory in Tokyo. I would also like to thank Dr. Motoshi Kikuchi (Waseda University) who assayed blood samples for gonadotrophins and thyroid hormones, and Dr Richard Talbot (Roslin Institute, Midlothian, United Kingdom) who assayed plasma prolactin. I would like to acknowledge the advice, comments, and hospitality of Dr. Tim Boswell, Dr. Ian Dunn, and Prof. Peter Sharp during my stay at the Roslin Institute. Thanks go in particular to Tim for helping to get the quail c-fos mRNA assay setup when all looked lost. I am indebted to my fellow students, Ellen Bennett, Emma Hawke, Sharon Henare, David and Sarah Simcock, and Mhairi Sutherland . Thank you for all the help with my experimental work, but also for the years of encouragement through the thick and thin, the great laughs we've all had, and especially the friendships. Good luck to all you guys! Finally, I thank my parents and my brother, whom I will always be indebted to for their support and encouragement. Last but not least, appreciative pats and hugs go to my dogs Marley and Yuki for the stress relief and reminding me who gave them their mischievous streaks. Xl Table of contents Abstract ............................................................................................................................... iii Acknowledgements ...................................... .................................................................... ix Table of contents ............................................................................................................... xi List of figures ........................................... ....................................................................... xvii List of tables ..................................................................................................................... xxi Chapter 1: General introduction 1 1.1 Photoperiodic control of reproduction in birds .................................................... 2 1.1.1 Photoperiodic induced gonadal growth and regression ..................... ........... 2 1.1.2 The hypothalamic-pituitary-gonadal axis . . . . . . .... . . . . . . ... .... . . . . . . . . . . . . . . . .... ... . .... . . . . . . . 3 1.1.2.1 Gonadotrophin-releasing hormone .................. ............... ................................ 3 1.1.2.2 Gonadotrophins ........................................ . .. ... .............................................. .. 4 1.1.2.3 Gonadal steroids ............................................................................................. 4 1.1.2.4 Prolactin ..... ................................................................. ........................... ........ 5 1.1.2.5 Regulation of gonadotrophin secretion .............................. ............................. 5 1.1.3 Photorefractoriness .... ..... ......... ..................................... .................. ..................... 7 1.1.3.1 Absolute photorefractoriness ............................................................ .............. 8 1.1.3.2 Relative photorefractoriness ... . ......... .............................................................. 8 1.1.3.3 Thyroid hormones .......................................................................................... 9 1.1.4 Gonadal growth and regression under artificial light cycles in Japanese quail .. ............................................ ................................. .......... . . ....... ..... 9 1.1.4.1 Using changes in daylength to induce gonadal regression in sexually mature Japanese quail .............................................. . .. ................................. 12 1.1.4.2 The use of changes in day length to prevent gonadal growth in sexually immature Japanese quail .......... .................................................................... 14 1.1.4.3 The effect of environmental temperature on gonadal regression ......... ......... 14 1.2 Stressors, stress and the stress response .............................................................. 15 1.2.1 Stress . . . . . . .. .. . .... . . ..... . . . ...... . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ... . . . . . . . . . .. . . . . . .. . . . . . . . . . . .... . . . . . . . . . . ... . . . . . . . 16 1.2.1.1 5 tressors ......................... .............................................................................. 16 1.2.2 The physiological response to stressors ......................... .......... ........... . ...... ...... 16 1.2.3 The hypothalamo-pituitary-adrenal axis . . . . . .. . . .. . . . . . . . . . . . . . . ... . ... . . . . ....... . . . . . . . . . . . . . 18 1.2.3.1 Hypothalamo-hypophysiotrophic factors ............................. ......................... 18 1.2.3.1.1 Corticotrophin-releasing factor ...................... ................... ..................... 18 XlI 1.2.3.1.2 Other hypothalamo-hypophysiotrophic factors ...................................... 19 1.2.3.2 ACTH .................................................................................................... . ..... 20 1.2.3.3 Corticosterone ........................................... .... . . .. ................. .......................... 21 1.2.3.3.1 Circadian rhythm of corticosterone secretion ......................................... 22 1.2.3.3.2 Effects of increased corticosterone secretion ................... . ....... .............. . . 22 1.2.3.3.3 Control of glucocorticoid secretion ......................................................... 23 1 .2.4 Measurement of the stress response . .. .......... . . . . . ... ........ . .. ................. . .. ....... . .. .. 24 1 .2.5 Effect of stressors on reproduction ......... . .. . ...... . . . ................ . . ... ..... ......... ....... .. 25 1.2.5.1 Different types of stressors .......................... ....................... ....... . .................. 27 1.2.5.2 How do stressors affect reproduction? ......................................................... 28 1.2.5.2.1 Glucocorticoids ................. . ...... . ................. .. .... . . ...... . ... ........ .................. 29 1 .2.6 Individual variation in the corticosterone response ... ................... . .. . ............ 32 1.2.6.1 Where is individual variation of the adrenocortical response mediated? ..... 33 1.2.6.1.1 Adrenal gland ........................................................................................ 33 1.2.6.1.2 Anterior pituitary gland ... . . . ............ ....................... .......... ............ ......... 35 1.2.6.1.3 Central nervous system ......................................................................... 36 1 .3 Outline of thesis ............ . . ......................... . ......................... ...................................... 37 Chapter 2: Effect of a single corticosterone injection on reproductive hormones in Japanese quail 41 2.1 Introduction .............................................................................................................. 41 2.2 Methods and materials . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 43 2.2.1 Animals and housing ...... . ........ . . . ..... .... . ...... . ...... . . . .. .. . . . . . ..... ........ ... . ....... . ........... 43 2.2.1.1 Source and type of birds .............................................................................. .43 2.2.1.2 Housing conditions ...................................................................................... 43 2.2.1.3 Food and water ............................................................................................ 43 2.2.2 Experimental design ............... ......... . ............... . . .. . .......... . . . . . . ....... .... . ...... . ..... . .. . . 44 2.2.2.1 Experiment 1 - Acute response to a single subcutaneous corticosterone injection in 8 week old male Japanese quail. . . .............................................. .44 2.2.2.2 Experiment 2 - Acute response to a single subcutaneous corticosterone injection in 8 week old female Japanese quail . . ...... . . .............. . ......... . .. . . . . . .. .. .44 2.2.3 Data collection . . ............. . . . .. . . .... ... .... . .... ....... . . . . . . . . . . . . . ....... .. . . . ........ . . ..... ...... ..... . ... 45 2.2.3.1 Corticosterone administration ..... ........ . . .............. . . ..... . ... .. ....... .. ....... ... . ...... . 45 2.2.3.2 Tissue and blood samples ............................................................................. 45 2.2.4 Hormone assays .. ..... ... . . . . ... . ........ . .. . ... . . . .. ...... .. . . . . . . . . . . . ..... ......... . . . . . . . .. . . . . .... .. . ... ... 46 2.2.4.1 Corticosterone .............................................................................................. 46 2.2.4.2 Testosterone ........... . . . .. . . . . . ..... .... . . .. . .... ....... . ....... ......... . . . ........ .. . .................. . . 47 Xlll 2.2.4.3 Oestradiol ........................... .................................................... . . . ................... 49 2.2.4.4 Luteinising hormone (LH) ............ ................. .............................................. 50 2.2.5 Calculation of the area under the corticosterone curve . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 51 2.2.6 Statistical analyses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 51 2.3 Results ........................................................................................................................ 52 2.3.1 Experiment 1 - Acute response to a single subcutaneous corticosterone injection in 8 week old male Japanese quail ... . . . ........... ........ 52 2.3.1.1 Gross morphology ....................................... . ... ............................................. 52 2.3.1.2 Plasma corticosterone ................... .................................................... ............ 52 2.3.1.3 Plasma testosterone ...................................................................................... 61 2.3.1.4 Plasma LH . . ............................................................ ..................................... 63 2.3.2 Experiment 2 - Acute response to a single subcutaneous corticosterone injection in 8 week old female Japanese quail .. . ... . ...... ......... 65 2.3.2.1 Gross morphology .................................... .................................................... 65 2.3.2.2 Plasma corticosterone ....... ....................... . ...................................... ...... . ....... 66 2.3.2.3 Plasma oestradiol . ....................................................................... ................. 72 2.4 Discussion .................................................. . ... . .......................................................... 74 2.4.1 Male quail . .... ........................ ............................................................................... 74 2.4.1.1 Corticosterone ............... ............................................................................. .. 74 2.4.1.2 Testosterone ................................... ..................................................... . .. ...... 75 2.4.1.3 LH .......................................................................................... ...................... 77 2.4.2 Female quail ...... ................................ . ................................ . .. .......... ... ........ ......... 78 2.4.2.1 Corticosterone .............................................................. .......... ............ .......... 78 2.4.2.2 Oestradiol ................................................ ..................................................... 78 2.4.3 Summary ... ........................... ....................... .. . ..... ......... ....................... ....... .... ..... 79 Chap-ter 3: Adrenocortical and testosterone responses of Japanese quail to a range of novel stimuli 81 3.1 Introduction ......................... ....... ........................................................................ . . . . .. 81 3.2 Methods and materials .............. ................................................... . ...... .......... ..... ..... 82 3.2.1 Animals and housing . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 82 3.2.1.1 Source and type of birds ............................................................................... 82 3.2.1.2 Housing conditions ......................................................................... ... . . . . . . . ... 83 3.2.1.3 Food and water .................................................... ......................................... 83 3.2.2 Experimental design ....................... . ... ................................ ............ . ................ .. 83 XlV 3.2.2.1 Experiment 1 - Quantification of the effect of manual restraint on plasma testosterone and corticosterone in the Japanese quail ....................... 83 3.2.2.2 Experiment 2 - Adrenocortical and testosterone responses to manual restraint, mechanical restraint and social stimuli in Japanese quail ............ 84 3.2.2.2.1 The effect of different stressors on the adrenocortical response .... . ......... 84 3.2.2.2.2 The effects of manual and mechanical restraint on testosterone and corticosterone ......................................................................................... 84 3.2.3 Data collection ........................ . . . . ........ ... . . . . ...... .... ......... ............ . ... ...................... 85 3.2.3.1 Administration of stressors .......................................................................... 85 3.2.3.1.1 Manual restraint ................................................................................... 85 3.2.3.1.2 Mechanical restraint ....... . . ..................................................................... 86 3.2.3.1.3 Exposure to an unfamiliar quail ............................................................ 86 3.2.3.2 Tissue and blood samples ....... ................................... . .................................. 86 3.2.4 Hormone assays .. . . . ....... . . .... . . . . .. ..................... . .............. ............ ............ . . . . . .... . . . . 87 3.2.4.1 Corticosterone ....................... ....................................................................... 87 3.2.4.2 Testosterone ................................................................................................. 87 3.2.5 Calculation of the area under the corticosterone curve .... . . ................ .... . ..... 87 3.2.6 Statistical analyses ..... . .. . . ............ ......... . . . . . ....... . . . .. . ............... . . . . ..... . .. . .. . .............. 87 3.3 Results ....................................................................................................................... 88 3.3.1 Experiment 1 - Quantification of the effect of manual restraint on plasma testosterone and corticosterone in the Japanese quail ..... ....... ..... . . . 88 3.3.1.1 Corticosterone .................. . .... . . ............................................................. ........ 88 3.3.1.2 Testosterone ............................................................................................... . . 89 3.3.2 Experiment 2 - Corticosterone and testosterone responses to manual restraint, mechanical restraint and social stimuli in Japanese quaiL . . . . . . .... 95 3.3.2.1 The effect of different stimuli on the corticosterone response .. ... . . ................ 95 3.3.2.2 The effect of mechanical and manual restraint on corticosterone and testosterone concentrations ........................................................................ 102 3.3.2.2.1 Plasma corticosterone ...... . . .................................................................. 102 3.3.2.2.2 Plasma testosterone ............................................................................. 102 3.4 Discussion .............................................................................. ................................. 108 3.4.1 Quantification of the effect of manual restraint on plasma testosterone and corticosterone . . . . .................... ... .... . . . . . . . . ........ . . . . . . . . ............ . . 109 3.4.1.1 Effect of manual restraint time on plasma corticosterone .......................... 109 3.4.1.2 Effect of duration of manual restraint on plasma testosterone ................... 111 3.4.2 Adrenocortical and testosterone responses to different stressors ............. 1 12 3.4.2.1 Adrenocortical responses to manual restraint, mechanical restraint and social stressors ..................................................................................... 112 3.4.3 Adrenocortical and testosterone response to manual restraint and mechanical restraint . ................ ..... . . .................................................... . . . . . . . . . .... 1 13 3.4.3.1 Plasma corticosterone ........... ......... . ............................................................ 113 xv 3.4.3.2 Plasma testosterone ................... .... ...... ........... . ......................... .. . ........ ....... 114 3.4.3 Summary .. ...... ... . . ..... . .... .... ... ........ . . . ... ...... . . ...... ...... . . ........ . . . ..... .. . . ..................... 115 Chapter 4: Variation within and between birds in corticosterone responses of Japanese quail 117 4.1 Introduction ............................................................................................................ 117 4.2 Methods and materials .......................................................................................... 118 4.2.1 Animals and housing .... .... ................. . ... . . ... . . ... . ....... . ....... . . ... ......... . . . ... . . ... . .... . . 118 4.2.1.1 Source and type of birds .............................. ......... . ............................. ........ 118 4.2.1.2 Housing conditions ..... ................................... . ......................... ............... ... 119 4.2.1.3 Food and water ........... ................ . .. ... .... ........ ................... . .... ................ ...... 119 4.2.2 Experimental design ........... ............ . . . . ... . .... . ............ . ...... ... . . . . .... . ... . . .... . ..... ..... . 1 19 4.2.3 Data collection .... .............. ........ ... . ... . . . ..... . . ............... .............. ........... ..... ... . . ..... . 1 19 4.2.4 Hormone assays ............ . . ... ... . . ... . ......... . .................. ....... .......................... ......... 120 4.2.5 Calculation of integrated corticosterone responses .................. . . ........ ......... 120 4.2.6 Statistical analyses . .......... ... ...... ....... ........... . . ... . .... ............. ........... . . . ... ...... . ... . . . . 121 4.3 Results ...................................................................................................................... 121 4.3.1 Corticosterone responses . .... . ....... ........... . ..... ..... ..... ......... ....................... ......... 121 4.3.2 Quantification of variation in corticosterone responses .............. .......... . . . .. 123 4.4 Discussion ............................................................................................................... 136 4.4.1 Variation in corticosterone responses .. .... . ... ..... . ..... .... ..................... . .... ...... ... 1 36 4.4.2 Quantification of variation in corticosterone responses .... .... . . ..... . .... . ........ 138 4.4.3 Summary . .... . . ... . . . . . . ...... . ... .................. .... . .... . ... . . ... ...... .... . . . . . ......... ....... . .... ......... 139 Chapter 5: Adrenal and pituitary responsiveness in quail with differing corticosterone responses to mechanical restraint 141 5.1 Introduction ............................................................................................................ 141 5.2 Methods and materials .......................................................................................... 143 5.2.1 Animals and housing .......... . ... . . . . .......... . ..... . ... . . . . . . . . . . ........ . ..... .............. ..... . . . . . . 143 5.2.1.1 Source and type of birds ................. ....... ....... . .. .............. . .. ... ..... .............. . . . . 143 5.2.1.2 Housing conditions ...... ................ .... .... ......... . . .............. . . .. ..... ......... . ...... .... 143 5.2.1.3 Food and water ....... .. .... ....................... . . ........ .................. . . . ...... ........ . .. ....... 143 5.2.2 Experimental design .................. ......... .. . .... . ....... . . . . ....... . ... . .... ... ............ . ..... .... . 144 5.2.2.1 Identification of birds with low or high corticosterone responses to mechanical restraint in individual quail . . ................................. . ................ 144 5.2.2.2 Relationship between the adrenocortical response and adrenal sensitivity to ACTH ....................... . . ... .......... . . ... ....... . ............. ......... .... . . . ... 144 XVl 5.2.2.2.1 Relationship between plasma corticosterone and dose of ACTH . . . . . . . . .. 144 5.2.2.2.2 Relationship between a sub-maximal dose of ACTH and plasma corticosterone concentrations in low and high response quail . . . . . . . . . . . . . 145 5.2.2.3 Relationship between the adrenocortical response and pituitary sensitivity to CRF . . . . . ..... . ... . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .. .. . . . . . . .. . . . . . . . . .... . 145 5.2.2.3.1 Relationship between plasma corticosterone and dose of CRF . . . . . . . . . . . . . 145 5.2.2.3.2 Relationship between a sub-maximal dose of CRF and plasma corticosterone concentrations in low and high adrenocortical response quail ...................................................................................... 146 5.2.3 Data collection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 146 5.2.3.1 Corticosterone administration . ... ....... . . . . . . . . . . . ... . .. . . . . . . . . . . . . ... . ... . .. . . .. . ... . . . . . . . . . 146 5.2.3.2 Tissue and blood samples ........................................................................... 146 5.2.3.3 Corticosterone assay . . .... . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . ... . . . . . .. . .. . . . . . . . . . . . . . . . . 147 5.2.4 Calculation of the area under the corticosterone curve . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . 148 5.2.5 Statistical analyses . . . . . . . . . . . ... ..... .... . ....... .. . . . . . . . . . . . . . . . . . . . . . . . . ... . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . 148 5.3 Results ..................................................................................................................... 149 5.3.1 Identifying low and high corticosterone responses to mechanical restraint. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .. . . .. . ... . . ... . . . . . ... . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . 149 5.3.2 Relationship between the adrenocortical response and adrenal sensitivity to ACTH . . . . . . ... ...... . . ... . . ... . . . . . . . . .... . . . . . . . . . . . . . . . . . . . . . .... . . . . . . . .. . . . . . . . . . . . . . . . . . . . 1 50 5.3.2.1 Relationship between plasma corticosterone and dose of ACTH . ...... . ..... .. . 150 5.3.2.2 Relationship between a sub-maximal dose of ACTH and plasma corticosterone concentrations in low and high response quail . .. . .... . . . . . . . . . . . . 159 5.3.3 Relationship between the adrenocortical response and pituitary sensitivity to CRF . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .. . . 1 65 5.3.3.1 Relationship between plasma corticosterone and dose of CRF . . . . . . . . . . . . . . . . . . . 165 5.3.3.2 Relationship between a sub-maximal dose of CRF and plasma corticosterone concentrations in low and high response quail .................... 171 5.4 Discussion ............................................................................................................... 176 5.4.1 ACTH dose response . . ..... . . . .. . . . . . . .. . . . . . . . . . . . . . ..... . . . . . . . . . . . . . . . . . . ... . . . . . . .. . .. . . . .. . . . . . .. . . ... 176 5.4.2 Adrenal sensitivity to ACTH in birds selected for low or high responses . ... . . ... .. ... . ... . . . . . . . . . ..... . . . . ... .. ... . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . ... . . . . . . . . .. . . . . . . . . . ..... .. 178 5.4.3 CRF dose response . . . . . . . . . . . ... . . . . . . . . . . . . . .. . .. . . . . . . . . . . . . . . . . . . . .. . . ... . . . . . . . . . . .... . . . . . . . . . . . . . . . . . . . 1 79 5 .4.4 Pituitary sensitivity to CRF in birds selected for low and high responses . . . . . . . . . . ... . . . . . ..... . . . . . . . . .. . . .. . .... . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . ... . . . . . . . . . . . . . . . . . . .. . .... 180 5.4.5 Summary .. .. . . ... . . . . . . . . . . . . . . . . . . ... . . . . .... . . . .. ... . . ......... . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . .. . . ... . . . . . . . . . . . . 181 Chapter 6: General discussion 183 Literature cited ............................................................................................................ 191 XVll List of figures Figure 1 .1 Eight week old male and female wild-type Japanese quail . . .... . . ... . . . . ... . ll Figure 1.2 Schematic representation of the hypothalamic-pituitary-adrenal and hypthalamic-pituitary-gonadal axes in birds and potential feedback pathways controlling the gonadal axis .. . . . ... . . ... . . . . . . . . . ... . . . . . . . . . . . . 17 Figure 2.1 Body weight, cloacal gland area, and paired testis weights in untreated male quail and in quail treated with oil or 1 .2 mg corticosterone . . . . . . . . . . . ... . . . . . . .. . . . . . . . . . . . . . . . . .. . ......... . . . .. . ..... . ... . . . . . . . . .... . . . . . ..... . . . . . . . .. 54 Figure 2.2 Plasma corticosterone, testosterone and LH in untreated male quail and in quail given a single injection of oil or 1 .2 mg corticosterone .... ...... ... . .. . . . .. . . . . . . . . . . .. . ........ . . . . . . . ...... . . . . . . . ... . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 55 Figure 2.3 Plasma corticosterone, testosterone, and LH in untreated male quail and in quail treated with oil or 1 .2 mg corticosterone . . . . . . . . . . . . . . . . . . .56 Figure 2.4 Plasma testosterone in male quail with initial concentrations < 1 nglml or 2: 1 nglml. Quail were untreated or given oil or corticosterone injections .. . . . ... . ..... . . . .. .. . . . . . . . . . . ..... . . . . . . . . . . . . . . . . . . . . ... . . . . . . . . . . . . . ... . . . 57 Figure 2.5 Total integrated responses for corticosterone, testosterone and LH in untreated male quail and in quail treated with oil or 1 .2 mg corticos terone ... ..... . . . . . .. . . . . . ..... . . ... . . . . . . . . . . . .. ........ . . . . . . . . .. . . ....... ... . ........ . . . . . . . ... .. . . 58 Figure 2.6 Body weight and ovary weight in untreated female quail and in quail treated with 1 .2 mg corticosterone . . . ..... .... . . . . . . . . . . . . . . . . . . ...... . . . .. .......... 67 Figure 2.7 Plasma corticosterone and oestradiol responses in female quail treated with oil or 1 .2 mg corticosterone .... . . . ..... . . . . . . . . . . . . . . . . . ..... .. . . . . . . . . . . . . . . . 68 Figure 2.8 Plasma corticosterone, oestradiol in female quail trea ted with oil or 1 .2 mg corticosterone . . . . . . . . . . . .. . . .. ... . . .. . ......................... .. . . . . . ... . . . . . . ... . . .. ... .. 69 Figure 2.9 Total integrated responses for corticosterone and oestradiol in female quail treated with oil or 1 .2 mg corticosterone . . . . . . . .. .. ....... . . . . . . . . . . 70 Figure 3.1 �lasma corticosterone -and testosterone in male quail blood sampled at 0 min then manually restrained up to 5, 10 or 15 min after the initial blood sample, then blood sampled at 15, 30, 60, 120 or 240 min .... ... ..... . . . . .... . . . . . . . . .. . . .. ... . . .. .... . . . . . . . . ............... ... . . . ..... ..... . . . . . . . ... . 90 Figure 3.2 Plasma corticosterone in male quail that were blood sampled, or in addition were manually or mechanically restrained for 15 min, or exposed to an unfamiliar quail that was visible but separated by a wire mesh or in direct physical contact . . .. . ... .. . . . . . . .. . . . . .... . . . . . .... .... . . . . . 97 Figure 3.3 Plasma corticosterone and change in plasma corticosterone relative to 0 min in male quail that were blood sampled, or in addition were manually or mechanically restrained for 15 min, or XVlll exposed to an unfamiliar quail that was visible but separated by a wire mesh or in direct physical contact ............................................ ........ 98 Figure 3.4 Total and corrected integrated corticosterone responses in male quail that were blood sampled, or in addition were mechanically restrained or manually restrained for 15 min, or exposed to an unfamiliar quail that was visible but separated by a wire mesh or in direct physical contact ........................ ................................................ .... 99 Figure 3.5 Plasma corticosterone and testosterone male quail blood sampled at 0 min then manually or mechanically restrained for 15 min, then blood sampled at 15, 30, 60, 120 or 240 min ................................... 104 Figure 4.1 Plasma corticosterone in individual male quail handled for 15 min on three separate occasions at 10 day intervals ............................. 125 Figure 4.2 Mean plasma corticosterone concentrations in individual male quail handled for 15 rnin on three separate occasions at 10 day in tervals ..................................................................................... . . . ...... . . . ...... 126 Figure 4.3 Mean plasma corticosterone concentrations in individual male quail that were returned to their cages between samples or handled for 15 min on three separate occasions at 10 day intervals ... 127 Figure 4.4 Total and corrected corticosterone responses of male quail that were returned to their cages between samples; or handled for 15 min on three separate occasions at 10 day intervals ...................... .. 128 Figure 4.5 Mean corticosterone concentrations at 0, 15, 30, 60, and 120 min in individual male quail handled for 15 min on three separate occasions at 10 day intervals ................................................... .................. 129 Figure 4.6 Mean total and corrected integrated corticosterone responses in individual male quail handled for 15 min on three separate occasions at 10 day intervals ..................................................................... 130 Figure 5.1 Plasma corticosterone in individual male quail immobilised by mechanical restraint for 15 min for all birds, birds with a low response to restraint, and birds with a high response to handling .... 152 Figure 5.2 Mean plasma corticosterone concentrations and total and corrected integrated corticosterone responses in male quail immobilised by mechanical restraint for 15 min ..................... ............... 153 Figure 5.3 Plasma corticosterone concentrations in individual male quail that received a single injection of saline or 0.05, 0.10, 0.20, or 0.50 IU ACTH/bird immediately after the 0 min blood sample . . . . . . . . . . . . . . . . . . . 154 Figure 5.4 Mean plasma corticosterone concentrations in male quail that received a single injection of ACTH at a dose of 0.05, 0.10, 0.20 or 0.50 IU/bird; and mean plasma corticosterone concentrations 15 min after an injection of saline or ACTH .......................................... 155 Figure 5.5 Total and corrected corticosterone responses in quail treated with saline or ACTH at doses of 0.05, 0.10, 0.20 or 0.50 IU/bird . ................... 156 Figure 5.6 Plasma corticosterone in individual male quail with low or high responses to restraint. Birds were injected with saline or 0.10 ID XIX ACTI-Ilbird . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ... . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 161 Figure 5.7 Mean plasma corticosterone concentrations; and the total and corrected integrated corticosterone responses in male quail selected for low or high responses to restraint that were treated with saline or 0.10 ID ACTH/bird . . ... . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ... . . . .. . . .... . 162 Figure 5.8 Plasma corticosterone in individual male quail selected for low or high responses to restraint. Birds were injected with saline or 6 J..Lg CRF . . . . . . . . . . .... . . ... . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .... . . . . . . . . . . . . . . . . . . ... . ... . . . . . . . . . . . . . . . . . . 167 Figure 5.9 Mean plasma corticosterone concentrations; and the total and corrected integrated corticosterone responses in male quail treated with saline or 2, 4, 6, or 8 J..Lg CRF/bird .. . . . . . ... . . . . . . . . . . . . . . . . . ... . . . . . . . . . 168 Figure 5.10 Plasma corticosterone in individual male quail selected for low or high responses to restraint. Birds were injected with saline or 6 J..Lg of CRF . ... . . . . . . . . . . . . . . . . .... . ..... . . ... . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ..... . . . . . . . . . . . . . . . . . . . . . . . . . . . . 172 Figure 5.11 Mean plasma corticosterone concentrations; and the total and corrected integrated corticosterone responses in male quail selected for low or high responses to restraint that were treated with saline or 6 J..Lg CRF/bird . .... . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ... . . . . . . . . . . . . . . . . ..... 173 Figure 6 Schematic representation of the stress response in Japanese quail summarising the findings of this disserta tion . . . . ... . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 190 List of tables Table 2.1 Two-way repeated measures ANOV A for corticosterone concentrations in untreated male quail and in quail treated with XXI oil or 1 .2 mg corticosterone . ...... .................................................................. 59 Table 2.2 Two-way repeated measures ANOVA for testosterone concentrations in untreated male quail and in quail treated with oil or 1 .2 mg corticosterone . ........................................... ...................... .... ... 62 Table 2.3 Two-way repeated measures ANOV A for LH concentrations in untreated male quail and in quail treated with oil or 1 .2 mg corticosterone ................................................................................................ 64 Table 2.4 Two-way repeated measures ANOVA for corticosterone concentrations in quail treated with oil or 1 .2 mg corticosterone . ........ 71 Table 2.5 Two-way repeated measures ANOVA for oestradiol concentrations in quail treated with oil or 1 .2 mg corticosterone . . . . . . . . . 73 Table 3.1 Two-way repeated measures ANOV A of corticosterone concentrations in male quail that were manually restrained for 2, 5, 10 or 15 minutes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 91 Table 3.2 Two-way repeated measures ANOV A of testosterone concentrations in male quail that were manually restrained for 2, 5, 10 or 15 minutes .......... .............................................. . ........................... 93 Table 3.3 Two-way repeated measures ANOVA of corticosterone concentrations in male quail exposed to different stimuli . ........... ....... 100 Table 3.4 Two-way repeated measures ANOVA of corticosterone concentrations in male quail manually or mechanically restrained for 15 min ................ .. .............. .. ... ........................... . ...... . . . ..... .. . 105 Table 3.5 Two-way repeated measures ANOVA of testosterone concentrations in male quail manually or mechanically restrained for 15 min .............................. .................................................... 107 Table 3.6 One-way ANOV A corticosterone and testosterone concentrations in quail sampled on Day 0 versus 10 . . . . . . . . . . . . . . . . . . . . . . . . . . . . 108 Table 4.1 Statistical analysis for variation in the corticosterone response in birds that were returned to their cages between samples (untreated) on three separate occasions at 10 day intervals .............. ... 131 Table 4.2 Statistical analysis for variation in the corticosterone response in birds that were handled for 15 min on three separate occasions at 10 day intervals ............................................... .. .................... .... .. ............ 132 XXll Table 4.3 One-way ANOVA analyses for comparison of untreated and handled corticosterone concentrations for each bleed at each time . ............................................................................................................. 133 Table 4.4 One-way ANOVA analyses for corticosterone concentrations for each time and integrated corticosterone responses in birds that were handled for 15 min on three separate occasions at 10 day intervals . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 133 Table 4.5 Smallest and largest mean values of corticosterone concentrations and integrated corticosterone responses for individual quail. ....... ............................................................................... .. . 134 Table 4.6 Coefficients of variation of mean corticosterone concentrations and integrated corticosterone responses within and between quail. ............................................................................................................ 134 Table 4.7 Coefficients of variation of mean corticosterone concentrations and integrated corticosterone responses within and between quail. .... ........................................................................................................ 135 Table 5.1 Two-way repeated measures ANOVA for plasma corticosterone concentrations in male quail treated with ACTH .................................. 157 Table 5.2 One-way ANOVA for the integrated corticosterone responses in quail treated with ACTH . ..................................................................... 159 Table 5.3 Two-way repeated measures ANOV A for corticosterone concentrations in low and high response quail treated with 0.10 IU ACTli/bird . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1 63 Table 5.4 One-way ANOVA for the integrated corticosterone responses in low and high response quail treated with 0.10 IU ACTli/bird ....... 164 Table 5.5 Two-way repeated measures ANOVA for corticosterone concentrations in male quail treated with CRF . .................................... 169 Table 5.6 One-way ANOV A for the integrated corticosterone responses in quail treated with CRF .......................................................................... 171 Table 5.7 Two-way repeated measures ANOVA for corticosterone concentrations in low and high response quail treated with 6 J-Lg CRF/bird . ..................................................................................................... 174 Table 5.8 One-way ANOVA for the integrated corticosterone responses in low and high response quail treated with 6 J-Lg CRF . ....................... 176 Chapter One Chapter 1 General introduction 1 Birds respond to stressors with an increase in corticosterone secretion. The precise role of corticosterone in the stress response in not well understood in birds. It is thought to inhibit reproduction although this effect can vary between individuals. The effect of corticosterone on sex steroid secretion was investigated in Japanese quail. Handling was found to have a profound effect on testosterone and the effect of handling and restraint on corticosterone and testosterone were investigated. Differences in the corticosterone response between individuals to the same stressor were quantified and the reasons for how they arise were investigated. This is the first physiological study to use the New Zealand strain of Japanese quail and it was necessary to conduct pilot studies to identify the photoperiodic conditions for maintaining quail in the laboratory. These preliminary experiments are not included in this thesis, but the review of photoperiodic literature is included to provide relevant background information. The rest of the literature review provides an overview of the role of corticosterone in avian reproduction and variation of the corticosterone response. 2 Cha pter One 1 . 1 Photoperiodic control of reproduction i n b irds 1 . 1 . 1 Photoperiodic i nduced gonadal g rowth and regression I t is well established that most birds are seasonal breeders, with reproduction timed to coincide with periods of ideal environmental temperature and food availability (Follett, 1984). Typically, exposure to a daylength of 12 hours or greater (long days; LD) stimulate the growth of the gonads, with onset of reproductive activity also influenced by other "ultimate" factors such as availability of food, competition, nesting conditions, predation pressure, and climate (Follett, 1984) . Breeding is timed to occur when environmental conditions are such that the chances of successfully raising young are greatest. Near the end of the breeding season, most photoperiodic species undergo spontaneous gonadal regression while daylengths are the same or longer than those which stimulated vernal gonadal growth. This phenomenon is known as photorefractoriness (Deviche and Small, 2001). It occurs at a time of long daylengths, but prevents further breeding when any attempt to reproduce would be disadvantageous to the adult and potential young (Murton and Westwood, 1977) . The photorefractive mechanism causes the rapid and spontaneous regression of the gonads. Photorefractoriness appears to be "programmed" into the hypothalamic-pituitary-gonadal axis (HPG-axis) during the first few days of photostimulation as reported in the American tree sparrow (Spizella arborea) (Re inert and Wilson, 1996a). All seasonally breeding birds possess a unique critical daylength threshold, which once exceeded triggers the gonadal growth. Photic cues affect the hypo thalamic­ pituitary-gonadal axis via photoreceptive structures and centres within the avian brain involved with photoperiodic time measurement such as the mediobasal hypo thalamus, the suprachiasmatic nucleus and pineal gland (Yasua et al., 2003). Extra-ocular and extra-pineal receptors located close to or in the lateral septum and tuberal hypothalamic regions have been identified as the primary encephalic photoreceptors in the avian brain (reviewed by Deviche and Small, 2001) . Wilson (1989) and Wilson (1990a) described the function of extra ocular centres outside of the ocular photoreceptors in American tree sparrows (Spizella arborea). However, the photoreceptive mechanism may be mediated differently in different Chapter One 3 species. Siopes and Wilson (1980) demonstrated that the photorefractive mechanism was mediated through retinal photoreceptors in quail, as blinded birds did not respond to a reduction in daylength. Photostimulation of the neural pathways associated with the photoreceptors is thought to directly influence the maintenance and numbers of gonadotrophin-releasing hormone (GnRH) neurons and the pulsatile secretion of GnRH from hypothalamic centres, which ultimately affect the secretion of gonadotropins from the anterior pituitary gland. Exposure to long days has been shown to stimulate an increase in GnRH secretion in Japanese quail (Follett and Maung, 1978) and gonadotrophin secretion increases in quail after just one day of exposure to long days (Follett et al., 1977). 1 . 1 .2 The hypothalamic-pitu itary-gonadal axis Exogenous cues such daylength, temperature and food availability are integrated in the brain and affect the hypothalamic-pituitary-gonadal axis through changes in GnRH secretion from the hypothalamus. 1 . 1 .2.1 Gonadotrophin-releasing hormone The secretion of luteinising hormone (LH) is under the control of GnRH. The effect of GnRH on follicle-stimulating hormone (FSH) secretion in birds is as yet unclear as previous in vivo GnRH treatment studies have proved inconclusive in chickens and Japanese quail (Hattori et al., 1986). Gonadotropin-releasing hormone has been isolated from the chicken hypothalamus in two distinct forms, cGnRH-I and cGnRH-II (King and Millar, 1982; Miyamoto et al., 1984). GnRH-II is the more potent stimulator of LH secretion in vitro (Millar et al., 1 986), although GnRH-I is likely to be the more important in vivo as it is secreted directly from the median eminence. GnRH-I (from this point GnRH will be taken to mean GnRH-I) is secreted from the GnRH neurons which terminate in the median eminence. A fine network of blood vessels, also known as the hypothalamo-hypothyseal portal circulation, carries GnRH and other neuropeptides and neurotransmitters from the hypothalamus to the pituitary gland. GnRH stimulates gonadotrophin secretion from the anterior 4 Chapter One pituitary gland or pars distalis. Photo stimulation results in an increase in GnRH synthesis and secretion in the median eminence, as seen in Japanese quail in which GnRH immunoreactivity is higher in the median eminence in birds on long days than in birds held on short days (Foster et al., 1988). 1 . 1 .2.2 Gonadotrophins The avian gonadotrophins LH and FSH are each composed of two glycoprotein subunits, a common a-subunit and a ,B-subunit specific to each hormone, with both units necessary for biological activity (Burke and Papkoff, 1 979) . Gonadotropin secretion is pulsatile and the frequency of pulses differs during hours of light and darkness. LH and FSH have specific roles within males and females in maintaining and controlling testicular and ovarian function. LH induces ovulation of the largest ovarian follicle as well as stimulating differing modes of steroidogenesis in the largest ovarian follicles and small follicles. In contrast, FSH maintains and stimulates steroidogenesis in the smaller yolky follicles and the production of progesterone, androstenedione, and oestradiol from thecal cells of small follicles (reviewed in Scanes, 2000). LH stimulates Leydig cells to differentiate and produce testosterone in males, whilst FSH stimulates Sertoli cell differentiation and spermatogenesis (reviewed in Scanes, 2000). 1 . 1 .2.3 Gonadal steroids Gonadal steroids are synthesised and secreted in response to an increase in gonadotrophin secretion. In the female, a variety of gonadal steroids are secreted from the follicles and stromal tissue. Progesterone, oestradiol-17,B and oestrone secretion are at their highest 4-6 hours prior to ovulation at the time of the preovulatory surge in LH (reviewed by Johnson, 2000). The gonadal steroids play essential roles in maintaining the passage of the ovulated follicle through the oviduct and in shell formation. In males, gonadotrophins stimulate the secretion of testosterone and androstenedione by Leydig cells, and the steroids influence spermatogenesis and the maintenance of testicular ducts and secondary sexual characteristics (reviewed by Kirby and Froman, 2000). Chapter One 5 In addition to physiological effects, elevated gonadal steroids are associated with specific breeding behaviour. Oestrogens are involved in stimulating courtship and copulatory behaviour in females, with elevated progesterone and prolactin during the la tter stages of foIlicular development inducing nest-building and incubation behaviour. Testosterone in males mediates copulatory behaviour indirectly through aromatization to oestrogens in the preoptic nuclei of the hypothalamus. Other courting behaviours such as strutting and crowing are stimulated through reduction of testosterone to Sa- and 5t3-dihydrotestosterone (reviewed by Phillips et al., 1985). 1 . 1 .2.4 Prolactin Prolactin is secreted from the pituitary gland under the influence of vasoactive intestinal polypeptide (VIP) from the hypothalamus (reviewed by Sharp et al., 1998). Prolactin concentrations increase during photostimulation and prolactin is associated with brooding and incubation behaviour. Maximal prolactin concentrations during breeding are associated with and contribute to the suppression of LH secretion in incubating poultry and are also associated with the development of photorefractoriness (reviewed by Sharp et al., 1998). The site of action of prolactin in the HPG-axis is unknown, although evidence shows that it is likely to act directly on GnRH and gonadotrophin secretion at the hypothalamus and anterior pituitary respectively. In turkeys, administration of prolactin results in a decline in hypothalamic GnRH and plasma LH (El Halawani et al., 1991; Rozenboim et al., 1993). However, the onset of photorefractoriness is likely to be independent of increasing prolactin concentrations, with prolactin being more involved with gonadal regression than with the initiation and maintenance of photorefractoriness Ouss, 1993). Alexander and Wolfson (1970) showed that twice daily injections of prolactin for 25 days did not affect ovarian or testicular weights but interrupted egg lay in quail, although Camper and Burke (1977) showed that ovarian and oviductal regression occurred following ovine prolactin treatment. 1 . 1 .2.5 Regu lation of gonadotroph in secretion Gonadotrophin secretion is influenced by sex steroid feedback on GnRH secretion (reviewed by Scanes, 2000) . The effect is predominantly inhibitory with LH 6 Chapter One secretion being inhibited by oestrogens and androgens in male Japanese quail (Davies and Follett, 1980). While elevated oestradiol inhibits LH secretion in females, progesterone can have either a positive or negative feedback influence on LH secretion depending on the stage of the ovulatory cycle. The preovulatory LH surge in hens is stimulated by progesterone (Etches, 1990), although the stimulatory action of progesterone on GnRH and LH secretion has been suggested to only occur while oestrogen concentrations are elevated (Phillips et al., 1985) . Inhibitory gonadal steroid feedback on the hypothalamus could influence gonadal state during the photorefractory period. It has been suggested that hypothalamic sensitivity to gonadal steroid inhibitory feedback increases in response to prolonged exposure to long days (Nicholls et al., 1988) . However, removal of the main sources of gonadal steroids by castration does not abolish the onset of photorefractoriness, with photorefractoriness manifesting itself in castrated birds at the same time as intact birds (Nicholls et al., 1988) . Wilson (1985) demonstrated in passerines that hypothalamic sensitivity to inhibitory gonadal steroid feedback decreased rather than increased after exposure to long daylength. A reduced hypothalamic sensitivity to negative feedback from steroids allows gonadal steroid secretion to increase resulting in the development of reproductive behaviour and development of accessory glands required during breeding (Nicholls et al., 1988). Hypothalamic sensitivity to gonadal steroids is restored upon exposure to short days. As a result, a low gonadal steroid concentration inhibits any increase in gonadotropin secretion from the anterior pituitary until daylength increases (Nicholls et al., 1988) . Exposure of a photosensitive bird to long days results in a decrease in hypo thalamic sensitivity to negative feedback inhibition from gonadal steroids, allowing an increase in GnRH and gonadotropin secretion, gonadal development and increased steroid secretion (Nicholls et al., 1988) . Hypothalamic sensitivity to gonadal steroids varies between species (Cockrem, 1995). Foster et al. (1987) and Goldsmith et al. (1989) reported a proliferation of GnRH perikarya in the median eminence of the hypothalamus in starlings once they were transferred from the photorefractory state to short days (8L:16D). Chapter One 7 However, the increase in GnRH perikarya and GnRH immunoreactive regions of the hypothalamus which occur as the bird regains photosensitivity are not reflected by a corresponding increase in gonadotropin secretion when kept under short days (Wilson, 1990b). This indicates that photostimulation and the resulting increase in gonadotrophin secretion will not occur in the photosensitive bird without an increase in daylength. LH secretion did not change in photosensitive American tree sparrows in response to increasing daylength while photoperiod remained below the photostimulation threshold (Wilson, 1990c), and LH remained suppressed even after photosensitivity has been fully restored. LH only increased when daylength increased beyond the critical daylength threshold (Wilson, 1 990b) . However, in the non-passerine species that have been studied, photorefractoriness does not appear to be associated with a change in the sensitivity of the hypothalamus to negative feedback inhibition from gonadal steroids (Nicholls et al., 1988) . 1 . 1 .3 Photorefractoriness During the breeding season whilst daylength may be still increasing, gonadal regression can occur as birds lose their responsiveness to long days (Cockrem, 1995). The loss of photosensitivity generally occurs whilst photoperiod is still above the critical daylength threshold for photo stimulation (Murton and Westwood, 1977). Photorefractiveness is a "reversible state of unresponsiveness to daylengths of gonadostimulatory duration" (Wilson, 1989), where the same daylength that stimulates the reproductive system (photo stimulation) also induces the shutdown of reproduction (photorefractoriness) . The photorefractory process encompasses a complex myriad of neuroendocrine mechanisms entrained to a variety of environmental cues or proximate factors but occurs to ensure that breeding terminates before environmental factors have the potential to impede reproductive success. Photorefractoriness is initiated at or above the level of the hypothalamus (Cockrem, 1995) . Associated with photorefractoriness is a decrease in GnRH and gonadotrophin secretion. It is unlikely that the pituitary gland or gonads become unresponsive to GnRH or gonadotrophins as gonadal development can be readily 8 Chapter One induced by administration of synthetic GnRH and gonadotropin respectively (reviewed by Phillips et al., 1985). Elevated gonadal steroid inhibitory feedback after photostimulation by long days is unlikely to be the primary cause of photorefractoriness. Wilson (1986) reported that the decrease in photosensitivity associated with photorefractoriness is independent of elevated plasma testosterone concentrations in American tree sparrows (Spizella arborea). However, it is important to note that the strength of the photorefractory response (degree of unresponsiveness to long days) varies between species (Cockrem, 1995) and species can be absolutely or relatively photorefractory. 1 . 1 .3 . 1 Absolute photorefractoriness Birds that undergo photorefractory gonadal regression and become completely unresponsive to daylengths that would otherwise be photostimulatory are known as absolutely photorefractory. In the starling (Sturnus vulgaris), a classic photorefractory model species, there is a reduction in areas of GnRH immunoreactive staining in the hypothalamic GnRH perikarya and median eminence as gonadal regression is nearly complete associated with the onset of photorefractoriness (Deviche and Small, 2001). Species such as the starling, house finch (Carpodacus mexicanus), and American tree sparrow (Spizella arborea) remain photorefractory as long as they are exposed to long days, and photosensitivity is only restored upon exposure to short days (reviewed by Deviche and Small, 2001) . 1 . 1 .3.2 Relative photorefractoriness Species that are relative photorefractory retain gonadal photosensitivity to extremely long daylengths and do not undergo gonadal regression until there is a decrease in daylength. The Japanese quail is one such species that is not completely unresponsive to increasing daylength during the photorefractory process. Vernal gonadal development is stimulated in photosensitive quail at a photoperiod of 1 1 .5 hours in quail at 52°N, whereas photorefractoriness is subsequently stimulated when the daylength falls below 14.5 hours (reviewed by Cockrem, 1 995). However, rather than being completely unresponsive to long days during the photorefractory period, large gonads can be maintained indefinitely by chronic exposure of Chapter One 9 photorefractory quail to unnaturally long day lengths, i.e. 20 hours (reviewed by Cockrem, 1 995). 1 . 1 .3.3 Thyroid hormones Thyroid hormones are imperative for the regulation and control of whole body metabolism of birds. The effects of environmental factors such as diet, temperature and daylength on the release of thyroid hormones may explain how environmental cues influence reproductive development (reviewed by Phillips et al., 1985). In many species, photorefractoriness is associated with the post-nuptial moult, and the cessation of mating behaviours and egg laying (reviewed by Phillips et al., 1985). Thyroid hormones have been implicated as the primary humoral factor initiating and maintaining the photorefractory state (Wilson and Reinert, 1993), as well as having a primary role in the moult process. Thyroidectomy abolishes the main source of thyroid hormones, and blocks the onset of photorefractoriness (Wilson and Reinert, 1993). The secretion of T4 and its metabolites increase after photostimulation, an effect which is required for gonadal growth (Reinert and Wilson, 1996b). However, thyroid secretion increases rapidly before the onset of photorefractoriness to concentrations that inhibit gonadotropin secretion at the same time as an increase in prolactin which also inhibits gonadotropin synthesis (reviewed by Sharp et al., 1998; Sharp and Blache, 2003). Thyroid hormones are known to stimulate the growth of new feathers, which eject the old feathers. In starlings, thyroidectomy a few days after photostimulation abolishes the onset of photorefractoriness and there is no gonadal regression (Dawson et al., 1986). 1 . 1 .4 Gonadal g rowth and regression under a rt ificial l ight cycles in Japanese quai l Japanese quail have been used extensively to study photoperiodic responses. Along with the starling and the domestic chicken, the photoperiodic response of the Japanese quail is one of the most extensively studied of any bird species (reviewed by Robinson and Follett, 1982; Wingfield, 1983; Follett and Nicholls, 1984; and Follett and Pearce-Kelly, 1990). The relative ease with which the timing of reproduction can be manipulated has made the Japanese quail an ideal species for photoperiodic studies. Japanese quail have also been selected for specific 10 Chapter One photoperiodic responses due to the relative ease of housing quail and short time to sexual maturity. Male and female Japanese quail (Figure 1 . 1 ) raised on long days from hatch are sexually mature by 5 and 6 weeks respectively (Siopes and Wilson, 1974). At sexual maturity, males and females of the wild-type randombred variety are sexually dimorphic by plumage. The increase in gonad size and androgen secretion results in the expression of a range of behaviours and secondary sexual characters such as crowing and development of a large cloacal gland (Oishi and Konishi, 1983) . The cloacal gland produces a thick foam used during copulation (Adkins-Regan, 1 999). Females are on average heavier than males and begin laying eggs between 6-8 weeks of age at a regular daily interval. Wilson and Huang (1962) showed that females laid 75% of their eggs in the late afternoon between 1500 - 1800 when raised on a 12L:12D photoperiod (lights on : 0600 - 1800) . Robinson and Follett (1982) reported that quail became photorefractory and underwent gonadal regression at 15 weeks of age when raised on natural daylengths of 13.5 - 14.5 hours of daylight. However, being relative photorefractory, gonadal regression can be prevented by maintaining birds on daylight hours greater than 15 hours (Robinson and Follett, 1982). Conditions for maintaining Japanese quail with large gonads indefinitely are well established (Robinson and Follett, 1982), but maintaining birds with small gonads is more problematic due to differences in responsiveness to short days between strains of quail (Oishi and Konishi, 1983). Previous reports show that there are significant differences in the photoperiodic response between different populations or "strains" of Japanese quail. Some studies report a high degree of success inducing gonadal regression after a decrease in daylength (Robinson and Follett, 1982; Follett and Nicholls, 1984), while others have taken a different approach by preventing gonadal growth by raising chicks on short days (Follett and Farner, 1966; Wakabayashi et al., 1992) . The response of the different strains of quail to short days varies greatly and in some instances, additional stimuli such as Chapter One Figure 1.1 Eight week old male (a) and female (b) wild-type Japanese quail. 1 1 12 Chapter One environmental temperature have been used in addition to a decrease in daylength to induce gonadal regression (Wada et al., 1990). The Japanese quail used in previous studies vary in origin from commercial "randombred" stock produced for egg or meat production, to strains genetically selected for particular photoperiodic responses. Japanese quail in New Zealand have been bred for both egg and meat production and nothing has been published on their photoperiodic response. Their photoperiodic response therefore had to be characterised in the present study as a prerequisite to studies of the effects of corticosterone on the control of the reproductive system. 1 . 1 .4 .1 Using changes in daylength to induce gonadal regression in sexually mature Japanese quail Transferring quail with large gonads from long to short days can decrease reproductive function (Wilson et al., 1965; Arrington et al., 1969; Stein and Bacon, 1976; Robinson and Follett, 1982; Wada, 1993). Wilson et al. (1965) showed that egg production ceased 3-5 days after a reduction in daylength from 14L:I0D to 8L:16D in 26-34 week old quail. Arrington et al. (1969) also showed that egg production had completely ceased 13 days after 12 week old birds were transferred from 14L:I0D to a shorter daylength of 4L:20D. A similar pattern was repeated by Stein and Bacon (1976) where transfer from 14L:I0D to 6L:18D caused a cessation in lay after 6 or 15 days in 89 or 140 day old females, respectively. Regression of the gonads occurs as a result of a decrease in gonadotrophin secretion, as LH injections maintain lay in quail transferred from long to short days (Wilson et al., 1965) . There is also involvement of thyroid hormones as pharmacological doses of T4 result in increases in FSH, LH and prolactin in quail transferred to short days (Follett et al., 1988) . Thyroid hormone concentrations also play a role in maintaining small gonads and are inherently higher in quail held on short days versus those on long days (Almeida, 1982) . Japanese quail with small gonads held on short days are extremely sensitive to subtle changes in daylength, and the utmost care must be taken in preventing an Chapter One 1 3 increase i n daylight hours or inadvertent exposure to any light outside the normal light regime. Hatanaka and Wada (1988) showed that 15 minutes exposure to light 12.5 - 14.5 hours after dawn during a 8L:16D photoperiod was enough to elevate LH. Plasma LH concentrations have been reported to increase within one day of exposure to increasing in daylength (Follett et al., 1977; Wada, 1979). There is a significant decrease in size of the androgen-dependent cloacal gland in some males when transferred from long to short days (Sachs, 1967; Siopes and Wilson, 1974; Siopes and Wilson, 1980; Follett and Pearce-Kelly, 1990) . Cloacal gland area had been shown to be highly correlated with the endocrine function of the testis (Siopes and Wilson, 1975). Follett and Pearce-Kelly (1990) showed that the cloacal gland started to reduce in size 1-2 weeks after a transfer from long to short days. In some studies (Siopes and Wilson, 1974; Siopes and Wilson, 1 980), testicular regression was inferred based on a reduction in cloacal gland area. Delville et al. (1 985) showed that plasma testosterone concentrations dropped by up to 50% within a week of being transferred to short days and were lowest after 28-35 days of short days. However, without any increase in daylength, testosterone concentrations and cloacal gland area spontaneously increased after 40 days of short days in this strain of quail. A decrease in daylength has also been shown to suppress LH secretion in quail (Gibson et al., 1975; Urbanski and Follett, 1982a; Urbanski and Follett, 1982b; Robinson and Follett, 1982; Follett and Nicholls, 1984). Decreasing daylength can cause complete gonadal regression in some strains of quail, but these have been quail selected for gonadal regression in response to a reduction in daylength (Robinson and Follett, 1 982; Follett and Nicholls, 1984). The response is much more varied in randombred quail sourced commercially, and in many of these strains some birds do not respond to decreases in daylength (Oishi and Konishi, 1983; Wada et al., 1990) . Oishi and Konishi (1983) showed that there was a correlation between the photoperiodic response and feather plumage in the Japanese commercial strain of quail, which underlines the genetic basis of this variation. 14 Chapter One 1 . 1 .4.2 The use of changes i n daylength to prevent gonadal g rowth i n sexual ly immature Japanese quai l Another approach used to maintain birds with small inactive gonads i s to raise immature birds on non-photostimulatory photo periods and prevent gonadal maturation. Testicular growth is inhibited by raising male quail on less than 12L :12D of light (Tanaka et al., 1965; Chaturvedi et al., 1992) . Follett and Farner (1966) showed that testicular growth was delayed for 170 days in males raised on 6L:18D, whereas males raised on long days reach sexual maturity by 35 days. Raising male quail on 8L: 16D delayed the onset of testicular growth to 70 days of age (Konishi et al., 1965; Siopes and Wilson, 1974), and this increased to 102 days where the photoperiod was 6L: 18D (Siopes and Wilson, 1974) . Testicular growth is delayed due to low gonadotrophin concen trations, as quail raised on 12L:12D respond to injections of ovine LH (McFarland et al., 1964). Ovarian growth and the onset of lay is also delayed by raising female quail on less than 12L:12D of light (Tanaka et al., 1965; Siopes and Wilson, 1980; Delville et al., 1985; Brain et al., 1988), while the onset of lay is normal in birds raised on 14 hours or more of light (Wilson and Huang, 1 962) . Raising quail on short days delays gonadal growth with the duration of the delay varying with daylength and the strain of quail. Despite the delay, spontaneous gonadal growth inevitably occurs in some if not all birds under non­ photostimulatory photoperiods. 1 . 1 .4.3 The effect of environmental temperature on gonadal regression Responses to short days in different strains of quail range from complete gonadal regression (Wilson et al., 1965; Arrington et al., 1969; Stein and Bacon, 1 976; Follett et al., 1977; Robinson and Follett, 1982) to varying degrees of gonadal regression (Oishi and Konishi, 1983), or no regression (Wada et al., 1992). A range of environmental factors influence seasonal breeding in birds, and one of primary importance is temperature. As mentioned previously, testicular growth is delayed until 70 days of age in quail raised on 8L:16D (Konishi et al., 1965) . Chapter One 15 However, Kato and Konishi (1968) showed that by lowering the environmental temperature from 25°C to 7-12°C, testicular regression occurred within a week in quail that had undergone testicular growth on 8L:16D, while no regression occurred in birds exposed to the same decrease in temperature but held on a continuous light regime. It has subsequently been shown that low temperatures along with short days are required to induce gonadal regression or prevent gonadal growth in the quail strain used in Japan (Tsuyoshi and Wada, 1992; Wada, 1993). Oishi and Konishi (1978) showed that both T4 and T3 concentrations are elevated in birds on short days, and that plasma T3 concentrations increased once short day quail were moved from 23 to 9°C, while LH decreased rapidly leading to regression of the gonads (Wada, 1993). Thyroid hormones have some influence in short day and cold temperature induced gonadal regression, although the mechanism by which this occurs is unknown (Wingfield et al., 1997a). Clearly genetically selecting Japanese quail that undergo gonadal regression when exposed to short days results in a quail strain that is highly responsive to a decrease in daylength as used by Robinson and Follett (1982) . Quail of a commercial origin such as those in New Zealand and Japan differ markedly in that the gonadal response to changes in daylength varies between individuals since no selection for photoperiodic response has occurred. Therefore the photoperiodic response of the New Zealand quail must be characterised before further studies on the effects of corticosterone on the control of the reproductive system. 1 .2 Stressors, stress and the stress response Individuals are exposed to challenges from the internal and external environment (stressors) on a daily basis. These challenges, sometimes called stress, can range from subtle environmental changes to an event that may threaten the survival of the individual. How the individual perceives the threat governs the magnitude of the stress response which includes behavioural and physiological responses that allow the animal to maintain or restore homeostasis. The individual's life history and experiences also contribute to how a threat is perceived and responded to; 16 Chapter One therefore a neural component is just as much a part of the response as the physiological. An inability of an individual to respond to threats to homeostasis results in compromises to a multitude of body functions (e.g. muscular-skeletal, reproduction), leading to debilitation through injury and disease and may result in death. 1 .2 . 1 Stress Stress is a term that is often used to broadly describe the causes and effects of challenges that an animal may encounter. There is no correct definition of stress and the term can often be misleading. Silverin (1998) defines stress as "an animal is stressed when it is exposed to adverse conditions which produce physiological responses in the individual that adapt the animal to the stressor and return it to a homeostatic condition". In this study, "stress" refers to the state when an animal experiences a stress response. 1 .2 .1 . 1 Stressors Stimuli that challenge homeostasis are commonly called "stressors" (Tilbrook et al., 2000) and in this study, the term will define any stimuli which activate the hypothalamic-pituitary-adrenal (HPA) axis. Stressors in birds can originate from a variety of sources such as the environment (e.g. extremes in temperature (Beuving and Yonder, 1978), transport or housing (Carlisle et al., 1998), physiological (injury or disease), psychological (fear or anxiety due to con specifics or predators), or nutritional (starvation or food deprivation, Tanabe et al., 1981)). 1 .2 .2 The phys iological response to stressors Exposure to a stressor activates the HPA-axis. Stimuli from the internal and external environment are integrated into neural information within the central­ nervous system (CNS), where they may be conSCiously and/or subconsciously perceived as stressful or benign (Harvey et al., 1984) . The way in w hich the stimulus is perceived will determine if the sympathoadrenal and HPA-axis are activated. In mammals as well as birds, stimulation of the HPA-axis (see Figure 1 .2) is characterised by activation of corticotrophin-releasing factor (CRF) neurons in the Chapter One 17 Brain e HPA axis HPG axis Stimu l i Autonomic ne�ous�--------------------------------� system PVN AVT CRF MT Corticotroph ACTH Adrenal Hypothalamus ME •.. . . +/ . . . . . . . . . . . . . . . . . . . . . . .. 8 e GnRH LH FSH e \ Gonad Anterior pitu itary Q gland '-.:::;J · · · ·e · · · · · · · · · · · · · · · · Glucocorticoids Catecholamines Sex steroids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Figure 1.2 Schematic representation of the hypothalamic-pituitary-adrenal (HPA) and hypothalamic-pituitary-gonadal (HPG) axes in birds, along with potential feedback (-ve and +/- indicate negative and positive/or negative feedback respectively) pathways which may regulate the gonadal axis. On the HPA-axis, the hypothalamo-hypophysial factors corticotrophin-releasing factor (CRF), arginine vasotocin (A VT) and mesotocin (MT) are released from neurons in the paraventricular nucleus (PVN) into the portal circulation stimulating adrenocorticotrophin hormone (ACTH) secretion from the adrenal gland. Glucocorticoids are secreted from the adrenal gland in response to ACTH. On the HPG-axis, gonadotrophin-releasing hormone (GnRH) is secreted from neurons terminating in the median eminence (ME) of the hypothalamus, and is released into the portal circulation stimulating secretion of the gonadotrophins luteinizing hormone (LH) and follicle-stimulating hormone (FSH) from gonadotrophs in the anterior pituitary gland. The gonadotrophins stimulate steroidogenesis in the gonads resulting in an increase in sex steroid secretion. 18 Chapter One paraventricular nucleus (PVN) and secretion of hypothalamo-hypophysiotrophic factors into the hypophysial portal system. CRF stimulates the production of adrenocorticotrophic hormone (ACTH), ,B-endorphin and a-melanocyte-stimulating hormone from corticotrophs in the pituitary gland, which are released during the stress response (Tilbrook et al., 2000). ACTH stimulates the synthesis of glucocorticoids from the adrenal gland and their secretion into the systemic circulation. An increase in plasma corticosterone concentration occurs during the avian stress response and directs physiological and behaviour modifications that allow the bird to maintain or restore homeostasis. In this study, a stress response has not occurred if there is no increase in plasma corticosterone concentrations. The sympathoadrenal system is also activated when an individual is exposed to a stressor and is composed of the sympathetic nervous system and adrenal chromaffin cells. Activation of the sympathoadrenal system coupled with the preganglionic innervation within parts of the adrenal gland results in an increase in catecholamine secretion into the systemic circulation, principally adrenaline (Tilbrook et al., 2000) .Catecholamine secretion from avian adrenal chromaffin cells has been shown to be stimulated by ACTH (reviewed by Carsia and Harvey, 2000) and in many cases the glucocorticoids and catecholamines act synergistically at target tissues. The effects of the catecholamines are not included in this review which will concentrate on the HPA-axis. 1 .2.3 The hypotha lamo-pituitary-adrenal axis 1 .2.3.1 Hypothalamo-hypophysiotrophic factors 1 .2 .3. 1 . 1 Corticotroph in-releasing factor The hypophysiotrophic role of CRF in mammals has been well established (reviewed by Rivier and Plotsky, 1986) . CRF is a 41-amino acid polypeptide synthesised in the PVN and released into the median eminence (ME) and hypophysial portal system resulting in activation of the pituitary corticotrophs (Lovejoy and Balment, 1 999) . Expression of genes encoding CRF are also influenced by changes in serotonin, acetylcholine, cytokines and interleukins 1 and 6 (Chadwick et al., 1993) . In mammals, CRF is acknowledged as the predominant Chapter One 19 hypophysiotrophic factor that stimulates ACTH secretion from the anterior pituitary during the stress response (Rivier and Plotsky, 1986) . The actions of CRF during the stress response are partly mediated through the two identified CRF receptor (CRF-R) subtypes; CRF-R type 1 is the main subtype and is present in the pituitary and brain, while CRF-R type 2 is present in the periphery and some areas of the brain (reviewed by Aguilera, 1998). The responsiveness of the HPA-axis during the stress response is controlled in part by changes in expression of CRF in the PVN as well as regulation of pituitary CRF receptors by CRF and glucocorticoid concentrations, and the possible role of a CRF binding protein (Behan et al., 1995; Aguilera, 1998) . Despite the well established function of CRF in mammals, an avian CRF is yet to be characterised (Speiss et al., 1998; Carsia and Harvey, 2000). However, there is measurable ovine CRF-immunoreactivity in the median eminence of the hypothalamus in Japanese quail (Mikami and Yamada, 1984) and chickens 06zsa et al., 1984), while the avian pituitary corticotrophs are responsive to mammalian CRF. Ovine CRF treatment of avian pituitary cells in vitro increases ACTH secretion (Castro et al., 1986; Carsia et al., 1986), while Japanese quail respond to in vivo treatment with human CRF (Launay et al., 1993). In addition, one of the two mammalian CRF receptor sub types, CRF-R type I, has been cloned in domestic fowl (Yu et al., 1996). 1 .2 .3 . 1 .2 Other hypotha lamo-hypophysiotroph ic factors In mammals, arginine vasopressin (A VP) is another hypothalamo­ hypophysiotrophic factor that controls ACTH concentrations (Konakchieva et al., 1997; Ma et al., 1997; Chrousos, 1 998) . AVP has a limited effect in birds (Gilles et al., 1982; Westerhof et al., 1992). Arginine vasotocin (AVT) is the natural hypothalamo­ hypophysiotrophic factor in birds (Westerhof et al. , 1992), while mesotocin (MT) is also known to control ACTH secretion. Nerve fibres in the median eminence of Japanese quail show immunoreactive staining to both A VT and MT (Mikami and Yamada, 1984). In vitro studies using duck and chicken pituitary cell cultures have shown A VT and MT to be ACTH secretagogues (Castro et al., 1986), although 20 Chapter One treatment with AVT or MT have produced inconsistent results. I t is possible that AVT and CRF act synergistically, with AVT potentiating the actions of CRF, a s reported by Westerhof e t al. (1992) and Romero and Wingfield (1998), where treatment with both factors stimulated a larger corticosterone response. The secretion of catecholamines immediately after exposure to a stressor and before activation of the HPA-axis is also thought to potentiate the responsiveness of pituitary to CRF, and thus enhance ACTH secretion (Harvey e t al., 1984) . In addition to changes in the synthesis and secretion of these hypothalamo­ hypophysiotrophic factors, the actions of CRF, AVT and MT in the stress response are also mediated by changes in responsiveness of the pituitary corticotrophs. The plasticity of pituitary responsiveness to CRF, AVT and MT has been extensively reported in wild, free living species (Romero and Wingfield, 1998). Seasonal differences in the stress response, as measured by the corticosterone response, are associated with changes in pituitary and adrenal responsiveness to hypothalamo­ hypophysiotrophic factors and ACTH respectively (Romero et al., 1997; Romero et al., 1998a; Romero et al., 1998b; Romero and Wingfield, 1998) . These results illustrate how the stress response is controlled at several different levels of the HPA­ axis. As well acting on the HPA-axis, AVT is also known to act on the HPG-axis. Male sexual behaviour is activated predominantly through the aromatization of testosterone to oestradiol. In Japanese quail, aromatase cell bodies and AVT fibres in the medial preoptic nucleus have been shown to be closely associated (Panzica and Viglietti-Panzica, 1999) . A VT has been shown to have an inhibitory effect on reproduction in male Japanese quail, intracerebroventricular or systemic treatment with AVT has a profound inhibitory effect sexual behaviour in male Japanese quail (Panzica and Viglietti-Panzica, 1999; Panzica et al., 2002) . 1 .2.3.2 ACTH In both mammals and birds, the synthesis and secretion of corticosteroids from adrenocortical cells is primarily under the control of adrenocorticotrophic hormone (ACTH) which is secreted from the anterior pituitary (reviewed by Mikami and Chapter One 21 Yamada, 1984; Mikami, 1986; Chrousos, 1998; Carsia and Harvey, 2000). As discussed previously, CRP and other hypothalamo-hypophysiotrophic factors, catecholamines and indoleamines stimulate ACTH secretion from the anterior pituitary (Harvey et al., 1984). ACTH is a 39-amino acid peptide synthesised from proopiomelanocortin, a large precursor protein complex (Hayashi et al., 1991) . ACTH is synthesised in the corticotrophic cells in the cephalic lobe of the anterior pituitary gland (Hayashi et al., 1991) . It is secreted under the control of hypothalamic-hypophysiotrophic factors (Carsia et al., 1986; Castro et al., 1986) and is also inhibited by glucocorticoid feedback (Carsia et al., 1986; Herold et al., 1992). ACTH is released into the systemic circulation and acts on adrenal cortical cells in the adrenal glands to stimulate the synthesis and secretion of corticosteroids and catecholamines. In birds, ACTH stimulates the secretion of the glucorticoid corticosterone (Freeman and Manning, 1 979; Davison et al., 1980; Davison et al., 1985; Beuving and Vonder, 1986), the mineralocorticoid aldosterone (Radke et al., 1984; Radke et al., 1985), and catecholamines (Zachariasen and Newcomer, 1 971). Adrenal corticosterone release in the Japanese quail is under the control of pituitary ACTH (Mikami and Yamada, 1984; Mikami, 1986) and Carsia et al. (1988) showed that corticosterone secretion increased in quail adrenocortical cells incubated with ACTH. Corticosterone concentrations have been shown to increase within 5 minutes of treatment with ACTH in a variety of birds (reviewed by Carsia and Harvey, 2000). Plasma ACTH concentrations have been shown to peak 5-1 0 minutes after initiation of a stress response in geese (Kovacs and Peczely, 1991) followed by a peak in corticosterone secretion several minutes later. 1 .2.3.3 Corticosterone Corticosterone is the primary avian glucocorticoid (Siegel, 1971; Harvey et al., 1984; Carsia and Harvey, 2000). ACTH stimulates an increase in adrenocortical cell secretion of corticosterone and this increase is indicative of activation of the HPA­ axis and a "non-specific s tress response" (Harvey et al., 1984) . Corticosterone is carried in the systemic circulation in free and bound forms, the latter being bound 22 Chapter One to corticosteroid binding globulins (CBG) (Wingfield et al., 1984) which reduce the bioactivity and clearance of corticosterone (Carsia and Harvey, 2000). 1 .2 .3 .3 . 1 Circadian rhythm of corticosterone secretion There is a distinct daily rhythm of corticosterone secretion as shown in Japanese quail (Boissin and Assenmacher, 1970; Kovacs and Peczely, 1983) and a variety of other birds (reviewed by Carsia and Harvey, 2000). Changes in basal corticosterone concentrations controlled by the HPA-axis appear to be mediated via the suprachiasmatic nucleus (King and Follett, 1997). Plasma corticosterone concentrations typically increase and peak during hours of darkness (Boissin and Assenmacher, 1970; Beuving and Vonder, 1977). Corticosterone concentrations remain low during hours of light, increase at the onset of darkness and remain elevated before decreasing at the onset of light (Boissin and Assenmacher, 1970). The timing of the peak varies and is influenced by photoperiod (Boissin and Assenmacher, 1970; Kovacs et al., 1983). In other Japanese quail studies, corticosterone has also been shown to peak in late morning and then decline (Kovacs and Peczely, 1983; Kovacs et al., 1983) . However, despite the change in concentrations during the day, overall basal corticosterone concentrations are low relative to the increase associated with a stress response. 1 .2 .3 .3 .2 Effects of increased corticosterone secretion on body and behaviour The increase in corticosterone secretion in response to a stressor is thought to help the bird make physiological and behavioural modifications that allow it to maintain homeostasis. During the stress response, the main actions of glucocorticoids are on intermediary metabolism, immune function and behaviour (reviewed by Siegel, 1995; Carsia and Harvey, 2000). The effects on intermediary metabolism vary between species, but the most immediate change is in glucose metabolism, associated with gluconeogenesis via fat and protein catabolism (reviewed by Harvey et al., 1984; Wingfield et al., 1998; Carsia and Harvey, 2000) . An increase in glucocorticoid secretion also alters immune function, although prolonged glucocorticoid concentrations can result in involution of the thymus, spleen and bursa of Fabricius (Carsia and Harvey, 2000). Chapter One 23 There is clear evidence that corticosterone influences behaviours in a mechanism that allows an individual to alter behaviour to maximise short term and long term survival. Elevated plasma corticosterone is known to have a profound effect on behaviour in birds, including decreases in aggression, parental and reproductive behaviours (Wingfield and Silverin, 1986) and increases in locomotor activity (Breuner et al., 1998) and foraging behaviour (Wingfield et al., 1998) . Breuner et al. (1998) avoided the disturbance caused by manually administrating corticosterone by infusing it directly into food, and clearly demonstrated an increase in locomotory activity in Gambel's White-Crowned Sparrows (Zonotrichia leucophnjs gambelii). Fearfulness also appears to be related to plasma corticosterone (reviewed by Jones, 1996; Carsia and Harvey, 2000). Corticosterone treatment of laying domestic hens increased tonic immobility, an indicator of fearfulness Gones et al., 1988). In Japanese quail, selection for low or high adrenocortical responses to mechanical restraint also caused a divergence in behaviour. Quail with a high corticosterone response to mechanical restraint showed greater fearfulness than that of random-bred quail Gones et al., 1992a; Jones et al., 1992b; Jones et al., 1994a). Prolonged exposure to a stressor can result in suppression of the reproductive axis in birds as well as other vertebrates (reviewed by Deviche, 1983; Harvey et al., 1984; Carsia and Harvey, 2000). Inhibition of the HPG-axis has been shown to occur at the hypothalamus, pituitary and at the gonads in mammals (Rivier and Rivest, 1991; Tilbrook et al., 2000; Tilbrook et al., 2002) and is thought to occur a t the same levels in birds (see Figure 1 .2) (reviewed by Carsia and Harvey, 2000). 1 .2 .3 .3 .3 Control of g lucocorticoid secretion Corticosterone secretion is regulated by feedback loops which are well established in mammals. Glucocorticoids exert a negative feedback affect on the hypothalamus and pituitary, through downregulation of brain corticosteroid receptors, and suppression of CRF, AVT and ACTH (Harvey et al., 1 984; Plotsky et al., 1989). Glucocorticoids have been shown to inhibit hypothalamic CRF mRNA expression (Ma et al., 1997) and release from the ME (Spinedi et al., 1 991) . Glucocorticoids also inhibit POMC transcription resulting in a decrease in ACTH secretion by the corticotrophs (Levin and Roberts, 1991), as well as reducing adrenal sensitivity to ACTH (Harvey et al., 1984). Negative feedback regulation limits glucocorticoid 24 Chapter One secretion, although the response of specific levels of the HPA-axis to feedback may vary between species and be specific to the type of stressor the bird has experienced (Chadwick et al., 1993) . In rats the pattern of HPA-axis activation differs according to the type of stressor. Desensitization of ACTH responses to repeated stimuli occur for stressors such as cold exposure or repeated immobilization, while the responses are preserved after repeated stimuli of other stressors (foot shock, insulin hypoglycaemia and intraperitoneal hypertonic saline injections) (Aguilera, 1998) . 1 .2.4 Measurement of the stress response The exposure of an individual to a stressor and activation of the HPA-axis can be assessed by measuring the hormones released in an adrenocortical response. Plasma concentrations of ACTH, catecholamines, and glucocorticoids all increase during the response , and measuring the change in concentrations is an ideal means of defining the stress response. Measuring plasma corticosterone is the widely accepted measure for defining the stress response given corticosterone is the predominant glucocorticoid secreted during activation of the HPA-axis (Carsia and Harvey, 2000). Plasma corticosterone concentrations increase rapidly (1-3 minutes) in response to a stressor in a variety of birds (Beuving and Yonder, 1 978; Harvey et al., 1980; Satterlee and Johnson, 1988) and the magnitude and duration of the response reflect the severity of the stressor (Harvey et al., 1984) . In Japanese quail, immobilisation causes corticosterone concentrations to increase within one minute, reaching a peak after four minutes and remained elevated during 20 minutes of immobilisation (Satterlee and Johnson, 1988) . Basal corticosterone concentrations vary between species and are low in domesticated species such as chickens (Freeman and Flack, 1 980), Japanese quail (Satterlee and Johnson, 1988) and ducks (Harvey et al., 1980), while significantly higher in some free-living species (Astheimer et al., 1994; Smith et al., 1994; Silverin, 1998). The corticosterone response is assessed experimentally by the application of a stressor to each bird. The method described by Wingfield ( 1994) termed the Chapter One 25 "capture stress protocol" is commonly used and involves the application of a standardised stressor (capture and handling). In wild birds, blood samples are collected before and then during the stressor, whereas in domesticated species housed in cages, samples can be collected after the stressor. The measurement of corticosterone concentrations allow the magnitude and duration of the response to be characterised. The collection of blood samples is in itself invasive and can stimulate a corticosterone response, therefore minimising the duration of the sampling procedure is important. Littin and Cockrem (2001) reported no increase in corticosterone concentrations after 15 minutes when blood was collected within 3 minutes in chickens, while there was no increase in Japanese quail after 15 minutes when samples were collected within 2 minutes (Boyd, 2000). However, Satterlee and Johnson (1988) showed plasma corticosterone increased 1 minute after being restrained and in further studies, blood was collected within 45 seconds to minimise measuring an adrenocortical response Gones et al., 2000). The magn itude of the corticosterone response has been shown to vary and can be influenced by factors such as the bird's age (Romero et al., 1998c), sex Gones et al., 1994b; Astheimer et al., 1995), or stage of the breeding season (Romero et al., 1997; Romero et al., 1998a; Romero et al., 1998b; Romero and Wingfield, 1998; Romero and Wingfield, 1999; Romero and Remage-Healey, 2000). These are factors to be mindful of when measuring the stress response in birds, although they can be minimised in laboratory species such as quail where birds are kept under controlled conditions. 1 .2 .5 Effect of stressors on reproduction Corticosteroids are an integral component of the avian reproductive system and have a range of effects both stimulatory and inhibitory (reviewed by Carsia and Harvey, 2000; Wingfield and Sapolsky, 2003). It is generally accepted that the prolonged exposure to a stressor suppresses reproduction in birds (reviewed by Deviche, 1983; Harvey et al., 1984) and this review will concentrate on the inhibitory effects of stress. However, the inhibitory relationship between stress and 26 Chapter One reproduction cannot be generalised for all individuals of a species, some individuals exhibit an inhibitory response while others do not (Moore and Jessop, 2003). From an ecological perspective, suppression of the reproductive system during which a bird is undergoing a period of stress is advantageous. Reproduction is an energetically expensive part of the life cycle, and reproduction during periods such as inclement weather and poor food availability when reproduction would be futile or compromise the survival of the individual is not ideal. Plasma corticosterone secretion can increase under such conditions, and exposure to continual high circulating concentrations of corticosterone for many days or even weeks contribute to suppression of the reproductive system in birds (reviewed by Deviche, 1983; Silverin, 1998; Wingfield et al., 1998; Carsia and Harvey, 2000). Treatment of male and female Japanese quail with corticosterone or ACTH decreased oviposition in females and suppressed LH (Martin et al., 1984) . The inhibitory effects of stress on reproductive function have been extensively studied in mammals and in particular rodents (reviewed by Rivier and Rivest, 1991; Tilbrook et al ., 2000; Tilbrook et al., 2002). In almost all cases, the main result of prolonged exposure to a stressor is a decrease in GnRH secretion, which in turn influences gonadotrophin secretion and the gonads. It is important to highlight that the effects on reproduction depend very much on the type and duration of stressor, and the effects differ between sexes. Studies in the mammalian literature show that the neural and endocrine pathways activated during the stress response vary depending on the type and duration of the stressor, and these differences alter how the reproductive system is affected (reviewed by Rivier and Rivest, 1991; Tilbrook et al., 2000; Tilbrook et al., 2002). In this review, the effect of stressors on reproduction will concentrate on the avian literature; however, reference will be made to mammalian research where appropriate. Chapter One 27 1 .2 .5 .1 Different types of stressors In mammals, a wide variety of physicat metabolic and psychological stressors have been shown to activate the HPA-axis, resulting in a decline in LH secretion (reviewed by Tilbrook et al., 2002). Similarly, birds show an adrenal response to a wide range of stressors (reviewed by Harvey et al., 1984) . While an inhibitory effect of corticosteroids has been shown in birds, there are relatively few studies which report the effects of different stressors on the reproductive system in species other than poultry (reviewed by Deviche, 1983). Repeated handling associated with frequent blood sampling has been shown to cause a decline in gonadotrophin and sex steroid secretion in poultry (Wilson and Sharp, 1975; Eskeland and Blom, 1979; Wilson et al., 1979; Johnson, 1981) . The results of Heiblum et al. (2000) conflict with these earlier studies, since short term mechanical restraint stimulated an increase in plasma testosterone concentrations at the same time as the corticosterone response in cockerels. The adrenocortical response to different stressors has also been extensively studied in the Japanese quail. Plasma corticosterone increases in response to a variety of stimuli such as capture and brief manual restraint by the experimenter, social challenges, cold, crating, and food and water deprivation Uones et al., 2000). The magnitude and duration of HPA-axis activation may vary between the types of stimuli indicating that different stress response pathways may be activated. Differences in adrenocortical responsiveness between low and high stress response lines of Japanese quail have been shown to result in different changes in cloacal gland physiology to mechanical restraint between the lines (Satterlee et al., 2002). Studies in rats (Hennessy and Levine, 1978; Hennessy et al., 1979) and starlings (Nephew et al., 2003) show that the size and duration of the corticosterone response reflect the strength of the stressor. Plasticity of the corticosterone response to varying intensities of stimulation is likely to be an adaptive mechanism to minimise the change in energy balance involved in the stress response. Activation of a response of a precise size adequate to deal with the stressor is advantageous, in contrast to a non-specific maximal response to any stressor irrespective of type or duration. The size of the elicited response will also reflect an animal's individual 28 Chapter One experiences and its own perception of whether the stimuli was a threat or benign. Modulation of the corticosterone response in free-living birds also extends to seasonal variation (Romero et al., 1997; Romero et al., 1 998a; Romero et al., 1998b; Romero and Wingfield, 1998) and the corticosterone response and reproduction are altered by unseasonal climatic conditions (Wingfield et al., 1999). Different stressors activate different parts of the brain, and the location of neuronal activity has been detected by localisation of immediate early gene expression in rats (reviewed by Akana and Dallman, 1997; Bonaz and Rivest, 1998; Van de Kar and Blair, 1999) . In rats, the paraventricular hypothalamus and central amygdala are two brain regions noted for activation of c-fos in response to exposure to stressful stimuli (Campbell et al., 2003; Zelena, 2003). Detection of the c-fos immediate early gene protein is used in the avian brain to detect neuronal activity following photoperiodic or sexual stimuli (Meddle and Follett, 1995; Meddle et al., 1999) and also distress in quail (Takeuchi et al., 1996) . Further work in this area will aid in the understanding of how different stressors activate a response in the bird and how this could explain the difference in the magnitude and duration of the stress response for a given stimuli. 1 .2.5.2 How do stressors affect reproduction? The stress-induced activation of the HPA-axis could influence the avian reproductive system at the hypothalamus, pituitary gland or gonads or any number of these levels as it does in mammals (see Figure 1 .2) . In mammals, the main effects of stressors are generally accepted to be on the secretion of GnRH from the hypothalamus and LH from the anterior pituitary gland (Brann and Mahesh, 1 991; Rivier and Rivest, 1991; Tilbrook et al., 2000; Tilbrook et al., 2002). The effects are mediated directly at the gonadotrophs that secrete the gonadotrophins and indirectly through interactive effects that influence gonadotroph function such as feedback from gonadal steroids or inhibin (reviewed by Rivier and Rivest, 1991; Tilbrook et al., 2000). In part, gonadotrophin suppression is mediated by glucocorticoid suppression of gonadotrophin release by acting at the level of the pituitary gland (Calogero et al., 1999) . Corticosterone Chapter One 29 treatment of quail pituitary gonadotrophs in vitro has been shown to inhibit LH secretion (Connolly and Callard, 1987), and continuous corticosterone infusion by osmotic pump in vivo causes a decline in plasma LH concentrations in hens (Etches et al., 1984; Williams et al., 1985; Petitte and Etches, 1989) . Despite the decline in LH, Etches et al. (1984) showed that the pituitary was just as sensitive to GnRH, indicating that corticosterone did not influence gonadotroph sensitivity to GnRH or the synthesis of LH. In contrast, subjecting Japanese quail to 3 days of food deprivation (nutritional stress) results in a decline in plasma LH caused by a decrease in gonadotrophin subunit mRNA expression (Kobayashi and Ishii, 2002). Wingfield and Silverin (1986) showed that subcutaneous corticosterone implants depressed testosterone but not LH in male song sparrows (Melospiza melodia). The differences in gonadotrophin response to corticosteroids in vivo and in vitro can be attributed to differences in species, sex and experimental protocol (Connolly and Callard, 1987). Differences in the way the HPG-axis is affected by the HPA-axis can also be attributed to the origin of the stressor that elicits the stress response. In the mammal, the origin and type of stressor has been clearly demonstrated to affect the neural pathways activated and how the HPA-axis is stimulated (reviewed by Day et al., 1999; Tilbrook et al., 2002). Further to this, it is likely that there are also differences in the way an individual responds to a stressor of psychological or psychogenic origin depending on the unique life experiences of that individual. Prolonged (chronic) exposure to noxious stimuli generally results in a decrease in gonadotrophin secretion and the decline in the reproductive system. However, the effects of acute or repeated acute exposure to stressful stimuli in mammals as well as birds are less understood as there are varying reports ranging from no effect, to stimulatory and inhibitory effects on reproduction (reviewed by Brann and Mahesh, 1991; Tilbrook et al., 2000). Differences in adrenocortical and reproductive axis responses to acute or chronic stressors are likely to be influenced by the type, intensity and duration of the stimuli, and the individual's perception of the stimuli. 1 .2 . 5 . 2 . 1 Glucocorticoids The structures of the HPG-axis in mammals and birds are fundamentally similar, and the avian and mammalian reproductive systems are likely to be influenced by 30 Chapter One stressors in similar ways. In mammals, chronically elevated glucocorticoids exert an inhibitory effect on reproductive function, and the effect is thought to be mediated in part by influencing gonadal sex steroid feedback and the modulation of peptides that regulate GnRH synthesis and secretion (Ahima and Harlan, 1992) . Central sites within the avian brain are thought to be the predominant sites of corticosteroid action in birds, and are generally accepted to influence gonadotrophin secretion directly and indirectly by influencing GnRH neurons (Deviche, 1983) . Glucocorticoid receptors (GR) have been localised in GnRH neurons in several phylogenetic groups (fish (Teitsma et al., 1999), mammals (Ahima and Harlan, 1992), and in the Japanese quail brain (Kovacs et al., 1989)) . In the rat brain, expression of GnRH mRNA and GnRH secretion are inhibited by corticosterone (Calogero et al., 1999; DeFranco et al., 1994) and the synthetic glucocorticoid dexamethasone (Attardi et al., 1997; Calogero et al., 1999; Schiml-Webb et al., 2001) . The mammalian GnRH neuron receives synaptic input from more than 18 known peptides and neurotransmitters, two of which are ACTH and CRF (reviewed by Tilbrook et al., 2002). CRF and GnRH neurons are found in some of the same regions of the Japanese quail brain and many perikarya terminate in the same regions (Mikami and Yamada, 1984; Mikami et al., 1988). However, no functional relationship between the two (CRF and GnRH neurons) has yet been demonstrated in birds. In the rat, direct synaptic connections between CRF and GnRH neurons have been found in the preoptic area, with CRF immunoreactive terminals impinging upon GnRH neurons (reviewed by Lovejoy and Balment, 1999) . The ability of glucocorticoids to directly and indirectly inhibit GnRH synthesis and secretion suggests that the hypo thalamus is one site of action where glucocorticoids may act to suppress the reproductive system. While the inhibitory effects of corticosterone on the avian reproductive system have been long recognised (Deviche, 1983), there has been little work to determine if corticosterone acts on GnRH in birds. Intra-hypothalamic corticosterone implants decreased LH and induced gonadal regression in photostimulated tree sparrows (Wilson and Follett, 1975), and daily corticosterone injections (3 mg/day) caused testicular regression and plasma LH to decline in ducklings (Deviche et al., 1979) . The continuous Chapter One 31 infusion of corticosterone via osmotic pumps resulted in ovarian regression and a decline in plasma LH and oestradiol concentrations (Etches et al., 1984; Williams et al., 1985; Petitte and Etches, 1 991) . The anterior pituitary remained responsive to GnRH, suggesting that the decline in gonadotrophin secretion was caused by glucocorticoids influencing GnRH secretion. However, quail pituitary gonadotrophs show a significant reduction in LH secretion when incubated with corticosterone (Connolly and Callard, 1987) indicating that corticosterone action is not limited to GnRH secretion. The association between elevated corticosterone and a decline in LH secretion is not always so clear, especiaUy where the stimulus is of short duration. In Japanese quail, a single injection of corticosterone (1 .2 mglbird) caused a significant increase in plasma corticosterone without affecting plasma LH (Boyd, 2000). Corticosterone delivered via subcutaneous implants in male tree sparrows did not affect LH or testosterone secretion (Astheimer et al., 2000). The direct actions of glucocorticoids on gonadal steroidogenesis are poorly known as the effects are often transitory and rapidly reversible, and are often masked by the overall suppression of GnRH and gonadotrophin secretion at higher centres. However, it is well accepted that short term exposure to stressors can have a direct effect on gonadal steroidogenesis, as shown in mammals. Restraint in rats caused an increase in plasma corticosterone, which was shown to inhibit testicular steroidogenesis resulting in a decrease testosterone (Bambino and Hsueh, 1981; Mann and Orr, 1990; Orr and Mann, 1992; Monder et al., 1 994). Elevated corticosterone alters testicular steroidogenesis in avian testicular cells in vitro (reviewed by Deviche, 1 983). Deviche et al. (1982) showed that corticosterone injections in Japanese quail reduced the production of androstenedione which led to a decrease in plasma testosterone concentration and also inhibited photoinduced cloacal gland growth. In light of the evidence in avian and mammalian in vitro studies, the acute stress response and transitory elevation in corticosterone are a likely cause of transitory reductions in sex steroid concentrations. A similar mechanism may also be acting in studies where a short 32 Chapter One period of handling (Wilson and Sharp, 1975; Wilson et al., 1979; Johnson, 1981) or an injection of corticosterone (Boyd, 2000) has resulted in a rapid decrease in testosterone but where LH has remained unchanged. 1 .2.6 Individual variation i n the corticosterone response The corticosterone response also varies within and between birds as shown in quail, domestic fowl and great tits (Satterlee and Johnson, 1988; Littin and Cockrem, 2001; Cockrem and Silverin, 2002a) . Individual differences in adrenocortical responsiveness have been used by Satterlee and Johnson (1988) to select birds for divergent adrenocortical responses over many generations. These quail are an ideal model for testing hypotheses on the relationships between behavioural, physiological and genetic traits and different aspects of stress and the adrenocortical response Gones et al., 1992a; Jones et al., 1992b; Jones et al., 1994a; Jones et al., 2000; Satterlee et al., 2002; Od eh et al., 2003a; Odeh et al., 2003b) . Variation between birds in their corticosterone responses have been attributed to differences in sex, age, reproductive status and body condition, as well as changes in environmental conditions such as rainfall, temperature and food availability (reviewed by Wingfield et al., 1992; Moore and Jessop, 2003; Wingfield and Sapolsky, 2003). The adrenocortical response to a standardised stressor has also been shown to vary during the season in some wild passerines, and is often referred to as "adrenocortical modulation" (Wingfield and Romero, 2001) . Modulation of the adrenocortical stress response appears to be regulated by mechanisms both upstream and downstream of the anterior pituitary (Wingfield and Romero, 2001) . There has been less attention to differences in corticosterone responsiveness to stressors within and between individual birds under constant conditions, where variation due to differences in body condition and the external environment can be minimised under laboratory conditions. Birds have been selected for divergent adrenocortical responses to stressors such as the cold (Brown and Nestor, 1973), social stress (Gross and Siegel, 1985), or immobilisation (Satterlee and Johnson, Chapter One 33 1988), indicating a genetic basis to this varia tion (as in other vertebrates, reviewed by Mormede et al., 2003). It has been assumed that a single measurement of a corticosterone response is sufficient to define the response for an individual bird. Studies in our laboratory on the domestic fowl and great tits (Littin and Cockrem, 2001; Cockrem and Silverin, 2002a) are the first to quantify within bird varia tion of the corticosterone response. The results showed that despite some variation, the adrenocortical response to 15 minutes of handling is generally repeatable within the same bird. The magnitude of the adrenocortical response to the same stressor varies between individuals in Japanese quail as shown by Satterlee and Johnson (1988). Despite the well established difference in adrenocortical responsiveness to immobilisation between individual quail, nothing is known of the causes of this variation. 1 .2.6.1 Where is individual variation of the adrenocortical response mediated? Varia tion between birds in their responses to emotional stressors could be due to differences in their perception of the stressor, or differences in the functioning of the HPA-axis. Differences in corticosterone responses are likely to be regulated at the CNS level (i.e. the individual's perception of the perturbation) and HPA-axis. 1 .2 .6 . 1 . 1 Adrenal gland Individual variation in the adrenocortical response could be due to differences in adrenal sensitivity to ACTH. Carsia et al. (1988) showed that maximal ACTH­ induced corticosterone secretion from isolated adrenocortical cells was greater in a strain selected for high corticosterone responses to immobilisation over random­ bred quail. Carsia et al. (1988) postulated that selection for a high corticosterone response altered the maximal adrenocortical response to ACTH without affecting cellular sensitivity to ACTH, as basal corticosterone secretion did not differ between high response and random-bred quail. McIlroy et al. (1999) showed that the ACTH­ induced corticosterone response of adrenocortical cells was enhanced by protein restriction in domestic fowl, and suggested that this was caused by either changes in adrenal sensitivity to ACTH, or changes in trans-membrane and intracellular 34 Chapter One pathways associated with ACTH binding. The results of these two studies show that the adrenocortical cells in birds can intrinsically differ between individuals in their sensitivity and responsiveness to ACTH. Studies by Satterlee and Johnson (1988) and Carsia et al. (1988) show that the adrenocortical response and adrenosteroidogenic properties of adrenocortical cells can differ significantly between quail exposed to a standardised stressor. Jones et al. (1992b) showed that selecting quail for divergent corticosterone responses to immobilisation also selected for other traits including different behaviour responses to stressors. Differences in intrinsic cellular properties of adrenocortical cells are not the sole cause of individual variation in the corticosterone response. Low or high corticosterone responses to immobilisation are clearly heritable, and differences are likely to be mediated at different levels of the HPA-axis as well as the CNS. In vitro treatment of adrenocortical cells from random-bred and high response quail showed differences between lines in maximal ACTH-induced corticosterone secretion but not basal secretion (Carsia et al., 1988). There does not appear to be an in vivo study where quail with differing corticosterone responses to mechanical restraint have been treated with a sub-maximal dose of ACTH. Such a study would elucidate whether differences in adrenal sensitivity to ACTH exist in vivo in quail with differing corticosterone responses. ACTH challenges have been used to identify changes in adrenal responsiveness in a range of passerines (Romero et al., 1998a; Romero et al., 1998b; Romero and Wingfield, 1998) . Measuring changes in plasma corticosterone concentrations following an ACTH challenge has been shown to be an accurate test for adrenal sensitivity in birds, since corticosterone is stimulated directly by ACTH. Plasma corticosterone increases rapidly « 30 minutes) in response to intra-muscular (i .m.), intra-venous (i.v.), and intra-peritoneal (i.p.) administration of mammalian ACTH in a variety of bird species (Beuving and Vonder, 1978; Harvey et al., 1980; Etches and Croze, 1983; Noirault et al., 1999; Wilson and Holberton, 2001; Faure et al., 2003). The use of the synthetic ACTH Synacthen ™ in chickens stimulated corticosterone secretion within 30 minutes (Beuving and Vonder, 1978; Harvey et al., 1980). Chapter One 35 1 . 2.6. 1 .2 Anterior pitu itary gland The anterior pituitary corticotrophs of domestic fowl are responsive to CRF peptides of mammalian origin (Carsia et al., 1986). In addition, AVT and MT have ACTH releasing properties in the isolated duck pituitary (Castro et al., 1986). However, in ter-specific differences in pituitary sensitivity to the three avian hypothalamo-hypophysiotrophic factors have been reported (Castro et al., 1 986; Westerhof et al., 1992). CRF is established as the primary ACTH secretagogue in mammals (Owens and Nemeroff, 1991). Individual variation in adrenal responsiveness could be mediated by differences in pituitary sensitivity to these releasing factors resulting in different rates of ACTH secretion between individuals. The administration of ACTH releasing factors in vivo has been shown to increase plasma corticosterone concentrations via action on ACTH secretion in pigeons (Calumba livia domestica) (Westerhof et al., 1992), Japanese quail (Launay et al., 1993), and a variety of free-living passerines (Romero et al., 1997; Romero et al., 1998a; Romero et al., 1998b; Romero et al., 1998d; Romero and Wingfield, 1 999). Assessing the effectiveness of ACTH-releasing factors by measuring corticosterone secretion is not ideal since they do not stimulate corticosterone directly (Romero et al., 1998b) . However, plasma corticosterone should reflect an ACTH response to CRF, AVT or MT. Pituitary responsiveness to mammalian CRF was reported by Westerhof et al. ( 1992) where the intravenous treatment of pigeons with ovine CRF (oCRF) with doses of 1 0 or 100 j.Lglkg BWT stimulated increases in plasma corticosterone at 15 and 60 minutes. Japanese quail also showed a small but significant increase in corticosterone after human CRF (15, 30, 45 j.Lglkg BWT) was administered sub­ cutaneously (Launay et al., 1993) . Clearly the Japanese quail pituitary is sensitive to mammalian CRF in vitro and in vivo (Carsia et al., 1986; Launay et al., 1993). The importance of a permissive action of the other two avian hypothalamo­ hypophysiotrophic factors A VT and MT cannot be d iscounted. Romero and Wingfield (1999) noted that that CRF and AVT administered together stimulated a more robust corticosterone response in Gambel's white-crowned sparrow than did 36 Chapter One CRF alone. The assessment of the corticosterone response to CRF, AVT or MT has been used successfully to measure differences in pituitary responsiveness in a variety of species. 1 .2 .6 . 1 .3 Central nervous system Variation in the corticosterone response between individual quail is also likely to be mediated at the CNS. As discussed previously, the adrenocortical response is only activated after the processing of the stimuli within the CNS. Stimuli are first detected by the bird visually, aurally, or through the somatosensory system; the information is then processed and integrated within the CNS and an HPA and sympathoadrenal response initiated where the stimuli pose a threat to homeostasis. Subtle changes in the internal or external environment or immediate threats such as the presence of a predator, stimulate different parts of the brain. In rats, the amygdala and paraventricular nucleus are well established as two brain regions involved in the processing of stressful or fearful stimuli (Campbell et al., 2003). Similarly, using C-fO$ immunoreactivity as a marker of immediate early gene expression, Takeuchi et al. (1996) showed that specific regions of the brain were activated in response to a social isolation stressor. Variation in the adrenocortical response between individuals is also influenced by the perception of the severity of the stressor by that individual based on its own previous experience and associative memory. These non-genetic differences in the brain associated with each individual's experiences will influence the responses of birds to stimuli. In mammals, exposure to acute stressful stimuli has been shown to alter an individual's subsequent response and enhance the later ability to acquire new memories about associations between stimuli (Beylin and Shors, 2003). In rats, the elevation in glucocorticoids following exposure to noxious stimuli enhance the recognition and association of the stressor as a source of stressful stimuli in future challenges (Beylin and Shors, 2003). The importance of a bird's perception of different stimuli and resulting activation or absence of a corticosterone response is illustrated by the study of Cockrem and Silverin (2002b). In their study, a marked corticosterone response was stimulated Chapter One 37 by exposing great tits to a moving stuffed owl (a recognisable predator), whereas birds shown a moving box of similar size and colouration did not have a corticosterone response. This clearly showed that an individual's perception of the stimuli within the CNS, whether innate or learned, is a primary regulatory component of the adrenocortical response. Few studies have addressed the variation in adrenocortical responsiveness between individuals by examining differences in neural activation within specific regions of the brain. Measuring C105 protein immunoreactivity to indicate expression of the immediate early gene has been the main method used to detect neuronal activation in the brain in mammals and birds (Meddle and Follett, 1995; Takeuchi et al., 1996; Meddle et al., 1999; Briski and Gillen, 2001). However, such measurements in birds with differing corticosterone responses to a standardised stressor have not been made. Underlying the psychophysiological level of HPA-axis control is a genetic component to the adrenocortical response which has been reported in a wide range of vertebrates (reviewed by Mormede et al., 2003). The Japanese quail is one such species which has been genetically selected over many generations for divergent adrenocortical responses to brief mechanical restraint (Satterlee and Johnson, 1988; Jones et al., 1992a; Jones et al., 1994a; Jones et al., 1994b) . The participation of genetic factors in variability of the stress response may occur at many levels of the HPA­ axis, although the prevalence of molecular biology techniques and improved availability of avian gene sequences are making research in this area more accessible (Mormede et al., 2003). 1 .3 Outl ine of thes is The overall aim of the research described in this disserta tion was to determine the effects of different stressors on corticosterone and sex steroid secretion in Japanese quail, and to investigate the neuroendocrine basis of variation between birds in their responses to stressors. The Japanese quail was used for this research as it is a convenient laboratory bird that had previously been studied in our laboratory. Boyd (2000) described 38 Chapter One corticosterone responses of quail to 15 minutes of handling and found marked variation between birds in their responses. She also found that daily corticosterone injections for 6 days reduced the size of the cloacal gland, an androgen-dependent structure, but that testosterone concentrations 24 hours after corticosterone injections were not reduced. This discrepancy was resolved in a further study in which corticosterone was shown to be elevated and testosterone reduced 3 and 6 but not 12 hours after a corticosterone injection. These findings led us to investigate the detailed time course of changes in corticosterone and testosterone after a corticosterone injection, the dynamics of testosterone during physiological corticosterone responses to stressors compared with pharmacological responses to corticosterone injections, and the sites within the hypothalamic-pituitary-adrenal axis at which variation in corticosterone responses arises. The current study addressed the following questions: 1 . What relationships are there between corticosterone, sex steroids and LH in the first 3 hours after a pharmacological elevation in plasma corticosterone? 2. What effect does the duration of a manual restraint stressor have on the magnitude and duration of the corticosterone response and testosterone secretion in male quail? 3. What effect do different stressful stimuli have on the magnitude and duration of the corticosterone response in male quail? 4. Are corticosterone responses to handling repeatable for individuals and what is the variation within and between individual quail? 5. Is variation in the corticosterone response between individual quail caused by differences in adrenal or pituitary sensitivity to ACTH or CRF respectively? Chapters 2 and 3 describe a series of experiments which were designed to address questions one, two and three. Chapter 4 addresses question four, while Chapter 5 Chapter One 39 describes a series of experiments designed to address question five. Thesis results and future directions for research are addressed in Chapter 6 (general discussion). Chapter Two Chapter 2 Effect of a s ingle corticosterone injection on reproductive hormones in Japanese quail 2 . 1 Introduction 41 Stress in its various forms is well documented as having the potential to have a detrimental effect on the reproductive system in a wide variety of vertebrates (reviewed by Rivier and Rivest, 1991) . Stressors which can be physical, psychological, nutritional, environmental, or social, trigger the release of adrenal steroids from the adrenal gland, which in turn stimulate a variety of physiological responses which allow the animal to respond or adapt to the stressor. Increased hypothalamic-pituitary-adrenal (HPA) axis activity can affect the function of the hypothalamic-pituitary-gonadal (HPG) axis in combination with other factors such as photoperiod and temperature, which were discussed in chapter 1. Exposure to a stressor results in an increase in plasma corticosterone concentrations in birds. Corticosterone is the major avian stress hormone released from the adrenal gland (Carsia and Harvey, 2000) . The roles of corticosterone are diverse and it plays an important function in the maintenance and termination of breeding during the reproductive cycle in birds (reviewed by Deviche, 1983). The inhibitory effects of corticosterone on the avian reproductive system have been long recognised (Deviche, 1983), and prolonged corticosterone treatment can suppress LH secretion and induce gonadal regression in a variety of species. Wilson and Follett (1975) showed that intra-hypothalamic implants of corticosterone caused gonadal regression and a decrease in LH secretion in 42 Chapter Two photo stimulated tree sparrows (Spizella arborea). Likewise, a prolonged but subtle increase in plasma corticosterone concentrations caused gonadal regression and the cessation of lay in hens (Etches et al., 1984; Williams et al., 1985; Petitte and Etches, 1991) . While the effects of chronically elevated plasma corticosterone concentrations on reproduction have been studied extensively, the effect of an acute increase in corticosterone on sex steroid concentrations has not been well characterised. Boyd (2000) showed that plasma testosterone concentrations in Japanese quail decreased 3 hours after a single corticosterone injection that had raised plasma corticosterone above normal physiological concentrations. The current study was designed to provide more detailed information on the relationship between plasma corticosterone and testosterone in male Japanese quail, and to characterise how oestradiol, the major sex steroid in females, is affected by a single corticosterone injection. The nature of the ovulatory cycle with surges in sex steroid concen tration at different stages of the cycle required that the timing of corticosterone injections in females was carefully planned. Van Tienhoven (1961) and Etches and Cunningham (1976) showed that exogenous corticosterone could induce ovulation in hens, while Wilson and Sharp (1976) and Etches and Croze (1983) showed that corticosterone could induce an increase in LH secretion. The aims of the experiments were : 1 . To determine the relationship between a dose of corticosterone and plasma concentrations of corticosterone, testosterone and luteinising hormone (LH) in male Japanese quail. 2. To determine the relationship between a dose of corticosterone and plasma concentrations of corticosterone and oestradiol in female Japanese quail given a single corticosterone injection. Chapter Two 2.2 Methods and materia ls 2.2 .1 Animals and housing 2.2.1 . 1 Source and type of bi rds 43 Seven week-old wild-type male and female Japanese quail (Coturnix coturnix japonica) were purchased from our supplier (Rangitikei Game Birds Ltd., Bulls). The birds had been raised under a long-day photoperiod (15L :9D) at air temperatures of 20 to 25°C in mixed sex groups. The 7 week-old birds were sexually mature at time of purchase with all females laying eggs, while all males had developed c10acal glands and cloacal foam was present. Sexually mature males could be selected by choosing males with enlarged cloacal glands (relative to males with no c10acal gland) as this has been shown to be a positive indicator of spermatogenesis, an increase in testicular weight and testosterone secretion (Ottinger and Brinkley, 1979) . 2 .2. 1 .2 Housing cond itions Each bird was identified with a numbered leg band and housed in an individual cage measuring 20 cm (W) x 35 cm (D) x 24 cm (H) in animal rooms in the Veterinary Science building. Quail were held on a long day photoperiod (16L:8D; lights on from 0900 - 0100 h). An extractor fan provided ventilation for each room, and only one sex was held in a room during each experiment. A temperature control unit enabled air temperature to be maintained at 20°e. Light in each room was provided by two 75W incandescent light bulbs controlled by a 24 hour;7 day time switch (HPM Excel Light Switch and Timer, Cat XL770T). The light intensity in the quail rooms was 130 lux in cages at the top, and 15 lux in cages near the floor of the room (Minolta Illuminance meter). 2.2. 1 .3 Food and water Quail were provided with fresh water and food (Harvey Farms quail layer mash) ab libitum. The food was the same as that used by Rangitikei Game Birds. 44 Chapter Two 2.2.2 Experimenta l design 2.2.2.1 Experiment 1 - Acute response to a single subcutaneous corticosterone injection in 8 week old male Japanese quai l Twenty one 7 week old male Japanese quail were held with ad libitum access to food and water under long days (16L :8D) at 20°C for one week. The birds were then assigned into an untreated control group and two treated groups (n =7 each) . These groups received either a single subcutaneous injection of a vehicle (sesame oil) or 1 .2 mg corticosterone per bird (6 mglkg) in sesame oil. A blood sample was taken at 0900 h (0 min, immediately before the injection) and then further samples were taken at 30 min, 1, 3, and 6 h after the injection. A terminal blood sample was obtained at 12 h when the birds were euthanased. Blood samples were collected from control birds at the same times. Body weight and the area of the cloacal gland were measured before the treatment and at 12 h. Plasma concentrations of corticosterone, testosterone and luteinising hormone (LH) were measured in all blood samples. 2.2.2.2 Experiment 2 - Acute response to a single subcutaneous corticosterone injection in 8 week old female Japanese quai l Twenty 7 week old female Japanese quail were obtained from our supplier and held with ad libitum access to food and water under long days (16L :8D) at 20°C for one week. The female experiment was conducted several months after the male study outlined above. The birds were then assigned into an oil- or corticosterone­ treated group (n= 10 each). These groups received either a single subcutaneous injection of a vehicle (sesame oil) or 1 .2 mg corticosterone per bird (6 mglkg) in sesame oil. Blood samples were collected at 0 min before the first injection, and then 30, 90 min, 3, 6, and 12 h after the injection. A terminal blood sample was obtained at 24 h when the birds were euthanased. The first blood sample was taken at 2000, in order to avoid the rise in corticosterone about the time of ovulation which occurs 15 - 30 min within oviposition (Woodard and Mather, 1964). The time of oviposition varied between 1400 - 1900 in these females. Body weight and the diameter of the cloacal opening were measured before the treatment and at 24 h. Plasma concentrations of corticosterone and oestradiol were measured in all blood samples. Chapter Two 45 The experiments were conducted under a protocol approved by the Massey University Animal Ethics Committee. 2.2.3 Data col lection 2.2.3.1 Corticosterone administration The corticosterone solution was prepared the day before the experiment and then kept on a magnetic stirrer (Chiltern Scientific) at 4°C. 250 mg of corticosterone (1 1�,21-dihydroxy-4-pregnene-3,20-dione, Sigrna Chemical Co., St Louis, USA) was dissolved in 1 ml of benzyl alcohol (Sigrna Chemical Co., St Louis, USA) then added to 9 ml of sesame oil (Sigma) producing a corticosterone 25 mg / ml stock solution. A solution was made from the stock producing a dose of 6 mg / kg of body weight in a 100 /-LI volume injection. 6 mg / kg was approximately 1 .2 mg per bird, a dose used previously (in our laboratory) to stimulate a strong pharmacological corticosterone response and decrease in testosterone in male Japanese quail (Boyd, 2000). Subcutaneous injections were made under the skin on the abdomen o f the quail using 1 ml insulin syringes and needles. 2.2.3.2 Tissue and b lood samples Blood samples were collected by venipuncture of the brachial vein in the wing with a 25 g needle, with 200 /-LI of blood collected into heparinised capillary tubes. All samples were collected within two minutes from the time the bird was removed from the cage. The final blood sample was collected when the birds were euthanased by stunning followed by decapitation. The blood was stored on ice until centrifugation at 2 000 g for 15 minutes at 4°C (Heraeus Christ 5000S refrigerated centrifuge) . The plasma was removed and stored at -20°C until assayed. The width and height of the c10acal protuberance was measured (±0.1 mm) in males and body weight was measured (±0.1 g) in all birds on the day before the experimental treatment. The reproductive system was removed after euthanasia and the oviduct and ovary or testes were weighed (±0.001 g). 46 Chapter Two 2.2.4 Hormone assays Corticosterone, testosterone, oestradiol, and luteinising hormone (LH) concentrations in plasma were determined by radioimmunoassay. 2.2.4.1 Corticosterone Corticosterone concentrations in extracted quail plasma were measured by radioimmunoassay, using a modification of the method described by (Wingfield et al., 1997a) . For extraction, 1 ml of plasma was thawed and spun in a 1 .5 ml Eppendorf tube at 14 000 g for 5 min (IEC Micromax ventilated microcentrifuge OM3590) to separate clear plasma from lipid. A 20 J.Ll aliquot of plasma was added to a screw top glass extraction tube (13 mm x 100 mm), and 1 ml of double-distilled dichloromethane (dichloromethane "AnalaR", BDH) was added using a Merck bottle-top dispenser. The plasma and dichloromethane were vortexed for 10 s, shaken together for 1 h on a Chiltern Scientific 5570 orbital shaker, and centrifuged at 4 500 g for 10 min in a Heraeus Christ 50005 refrigerated centrifuge to separate the organic and aqueous phases. An 800 .ul aliquot of the organic phase was removed from each tube and dried under a stream of air in a heating block at 37°C. The dried extract was then reconstituted in 450 .u1 of phosphate-buffered saline with gelatine (PBSG; O.lM, pH 7.0), vortexed, then shaken for 1 h at room temperature and then stored overnight a t 4T. The percentage recovery of corticosterone following the extraction process was determined by mixing tritiated corticosterone with 12 different plasma samples in triplicate . A 100 .ul aliquot of the reconstituted extract containing label was taken for determination of the percentage recovery of the sample. This yielded a mean recovery for corticosterone of 99.7 ± 2.0% (n = 12) for quail plasma, with this mean value used to calculate corticosterone concentrations in samples. All samples subsequently extracted were extracted without the addition of tritiated corticosterone. Samples were assayed in duplicate and were randomly distributed throughout two assays. Reconstituted extracts were incubated with 100 .ul of antibody (gift of Dr. R. J. Etches, University of Guelph, Ontario, Canada; 1 :18 000 final dilution) and 100 J.LI of tritiated corticosterone (approximately 5 000 cpm; 1 ,2,6,7-3H-corticosterone Chapter Two 47 TRK.406 Amersham, UK) at 4°C overnight. Bound and free components were separated by the addition of 500 J.LI dextran-coated charcoal (2.5 gIl charcoal, 0.25 gIl dextran (Dextran T70, Amersham Pharmacia) in PBSG) for 15 min at 4°C. Tubes were then centrifuged at 4 500 g for 15 minutes at 4°C and the supernatant poured off. 3 ml of scintillant (5 gIl PPO (2,5-diphenyl-oxazole, Sigma), 0 .3 gIl dimethyl POPOP (l ,4-bis- [4-methyl-5-phenyl-2-oxazoly]-benzene, Sigma) in toluene) was added, the samples were shaken for 1 h, left for 1h at room temperature, and then each sample was counted for five minutes in a Wallac 1409-41 1 liquid scintillation counter. The limit of sensitivity of the radioimmunoassay, defined as the hormone concentration at the mean minus two standard devia tions from the zero hormone point on the standard curve and expressed as corticosterone concentration in plasma, was 0.61 nglml. Serial dilutions of extracted quail plasma in assay buffer (PBSG) were parallel to the corticosterone standard curve (n=3) . Recovery of corticosterone added to quail plasma was 93.9 ± 4.9%, 98.6 ± 7.3%, and 88.9 ± 5.5% for three samples. Solutions of corticosterone in PBSG were prepared at concentrations that gave approximately 20, 50, and 80% binding on the standard curve. These solutions were used as high, medium and low quality control samples in every assay. The intra-assay coefficients of varia tion for these samples were 13.2, 14.8, and 13.4% (n=20) and inter-assay coefficients of variation in all 12 assays were 4.9, 6. 1 , and 4.3%. The cross-reactions of the antibody with other steroids tested by Etches (1976) were deoxycorticosterone (27.9%), cortisol (6.9%), progesterone (37.6%), l l�­ hydroxyprogesterone (21 .3%), oestradiol, testosterone, 1 7a-hydroxyprogesterone, l l-deoxycortisol, aldosterone and pregnenalone (all < 5%). 2.2.4.2 Testosterone Testosterone concentrations in extracted quail plasma were measured by radioimmunoassay, using a modification of the method described by (Wingfield et al., 1997) . For extraction, 1 ml of plasma was thawed and centrifuged in a 1 .5 ml Eppendorf tube at 14 000 g for 5 min (IEC Micromax ventilated microcentrifuge 48 Chapter Two OM3590) to separate clear plasma from lipid within the plasma sample. A 20 J.LI aliquot of plasma was added to a screw top glass extraction tube (13 mm x 100 mm), and 1 ml of double-distilled dichloromethane (dichloromethane 11 AnalaR", BDH) was added using a Merck bottle-top dispenser. The plasma and dichloromethane were vortexed for 10 s, shaken together for 1 h on a Chiltern Scientific SS70 orbital shaker, and centrifuged at 4 500 g for 1 0 min in a Heraeus Christ 5000S refrigerated centrifuge to separate the organic and aqueous phases. An 800 J.LI aliquot of the organic phase was removed from each tube and dried under a stream of air in a heating block at 37°C. The dried extract was then reconstituted in 450 J.LI of phosphate-buffered saline with gelatine (PBSG; O.lM, pH 7.0), vortexed, then shaken for 1 h a t room temperature and then left overnight at 4°C. The percentage recovery of testosterone following the extraction process was determined by mixing tritiated testosterone with six different plasma samples in triplicate. A 100 J.LI aliquot of the reconstituted extract containing label was taken for determination of the percentage recovery of the sample. This yielded a mean recovery for testosterone of 96.6 ± 4.0% (n= 18) for quail plasma, with this mean value used to calculate testosterone concentrations in samples. All samples subsequently extracted were extracted without the addition of tritiated testosterone. Samples were assayed in duplicate and were randomly distributed throughout three assays. Reconstituted extracts were incubated with 100 J.LI of antibody (Endocrine Sciences CA, US; testosterone antiserum T3-125 raised in rabbit) and 100 J.LI of tritiated testosterone (approximately 5 000 cpm; 3H-testosterone TRK.406 Amersham, UK) at 4°C overnight. Bound and free components were separated by the addition of 500 J.LI dextran-coated charcoal (2.5 gIl charcoal, 0.25 gIl dextran (Dextran T70, Amersham Pharmacia) in PBSG) for 15 min at 4°C. Tubes were then centrifuged at 4 500 g for 15 minutes at 4°C and the supernatant poured off. Three ml of scintillant (5 gIl PPO (2,5-diphenyl-oxazole, Sigma), 0.3 gIl dimethyl POPOP (1,4-bis- [4-methyl-5-phenyl-2-oxazoly]-benzene, Sigma) in toluene) was added, the samples were shaken for 1 h, left for 1 h at room temperature, and then each sample was counted for five min in a Wallac 1409-411 liquid scintillation counter. Chapter Two 49 The limit of sensitivity of the radioimmunoassay, defined as the hormone concentration at the mean minus two standard deviations from the zero hormone point on the standard curve and expressed as testosterone concentration in plasma, was 0.36 nglml. Serial dilutions of extracted quail plasma in assay buffer (PBSG) were parallel to the testosterone standard curve (n=3) . Recovery of testosterone added to quail plasma was 97.4 ± 1 .7%, 99.1 ± 8.1 %, and 102.0 ± 2.7% for three samples. Solutions of testosterone in PBSG were prepared at concentrations that gave approximately 20, 50, and 80% binding on the standard curve. These solutions were used as high, medium and low quality control samples in every assay. The intra-assay coefficients of variation for these three samples were 19.3, 7.8, and 5.5% (n = 20). The inter-assay coefficients of variation for three assays were 9.5, 8.3, and 11 .7% for high, medium and low quality controls respectively. The cross-reactions of the antibody with other steroids tested by Endocrine Sciences were dihydrotestosterone (20%), corticosterone « 0.01 %), oestradiol (0.14%), �-1 - testosterone (52%), 4-androsten-313-1713-diol (3%), 5a-androstan-313-1 713-diol (1 .8%), �-4-androstenedione (0.5%), others « 0.5%). 2.2.4.3 Oestradiol Oestradiol concentrations in quail plasma were measured by radioimmunoassay using an ICN 1mmunoChem n. Double Antibody 1713-estradiol 1251 RIA kit (lCN Biomedicals, Costa Mesa, CA). Samples were assayed in duplicate and were run in one assay. For assay, plasma was thawed and centrifuged in a 1 .5 ml Eppendorf tube at 14 000 g for 5 min (lEC Micromax ventilated microcentrifuge OM3590) to separate clear plasma from lipid within the plasma sample. A 10 /-LI aliquot of plasma was incubated with 100 ILl of antibody (anti-1713-oestradiol Cat # 07-1381 13) and 100 /-LI of iodinated oestradiol (approximately 5 000 cpm; 1251-oestradiol Cat # 07-138121) at 37°C for 90 minutes. After incubation, 100 /-LI of precipitant solution (Cat # 07-166624) was added to all tubes and mixed using a Chiltern Scientific SS70 orbital shaker for approximately 5 s. Tubes were then centrifuged at 4 500 g for 15 min at 4°C and the supernatant aspirated . The precipitates were then counted in a LKB Wallac 1261 Multigamma Gamma Counter. 50 Chapter Two The limit of sensitivity of the radioimmunoassay, defined as the hormone concentration at the mean minus two standard deviations from the zero hormone point on the standard curve and expressed as oestradiol concentration in plasma, was 7.4 pg/m!. Serial dilutions of quail plasma in diluent buffer were parallel to the oestradiol standard curve (n =3) . Recovery of oestradiol added to quail plasma was 97.4 ± 13.0%, 97.4 ± 12.3%, and 92.1 ± 1 1 . 1% for three samples. Solutions of oestradiol in PBSG were prepared at concentrations that gave approximately 20, 50, and 80% binding on the standard curve. These solutions were used as high, medium and low quality control samples in every assay. Intra-assay coefficients of variation have yet to be determined. No inter-assay coefficients of variation were calculated as all samples were analysed in one assay. The cross-reactions of the antibody with steroids tested by ICN Biomedicals were oestradiol-17� (100.00%), oestrone (20.00%), oestriol (1 .51 %), oestradiol-17a (0.68%), ethinyl oestradiol, androstenedione, dehydroepiandrosterone (DHEA), 5a­ dihydrotestosterone, 20a-dihydroprogesterone, deoxycorticosterone, progesterone, testosterone, pregnenolone, 17-hydroxypregnenolone, DHEA-sulphate, aldosterone, cortisol, 1 1-desoxycortisol, 1 7a-hydroxyprogesterone and cholesteronol (all < 0.01 %) . 2.2.4.4 Luteinising hormone (LH) Luteinizing hormone (LH) concentrations in plasma were measured directly by Dr M. Kikuchi using a modification of the double- antibody radioimmunoassay for chicken LH described by (Hattori and Wakabayashi, 1979) . Purified chicken LH (CANOMS12442B; Kikuchi and Ishii, 1 989) was used for radioiodination, with a crude chicken LH fraction (AGC112B; Kikuchi and Ishii, 1 989) for the reference standard (Silverin et al., 1999), and an anti-avian LH antiserum (AL-MH#l) (Hattori and Wakabayashi, 1 979) . Plasma was thawed and centrifuged in a 1 .5 ml Eppendorf tube at 14 000 g for 5 min (lEC Micromax ventilated microcentrifuge OM3590) to separate clear plasma from lipid within the plasma sample. Samples were assayed in duplicate within a single assay. Chapter Two 51 Twenty five fLl of plasma or standard was incubated overnight a t 4°C with 400 fLl of 0.1 % gelatin in PBS (PBSG), 1 00 fLl of diluted antiserum (1 :100,000) in 1 % normal rabbit serum (0.05M EDTA-PBS) (Hattori and Wakabayashi, 1979) . One hundred fLl of 131I_LH in 0.1% PBSG was added the day after incubation and incubated overnight at 4°C. A goat anti-rabbit y-globulin serum (H-4) was added to all tubes and then mixed before the supernatant was aspirated and the pellet counted (Hattori and Wakabayashi, 1979) . The LH content in each sample was expressed in terms of the weight of CANOMS12442B. The limit of sensitivity of the radioimmunoassay defined as the concentration of LH that corresponded to the percentage binding two standard deviations from the mean % bound of zero control tubes, was 0.06 nglml. The intra-assay coefficient of variation provided by Dr M. Kikuchi was 6.2% . Quail plasma produced dilution curves parallel to chicken LH standard curves. 2.2 .5 Calcu lation of the area under the corti costerone c u rve The area under the curve for plasma corticosterone concentrations versus time was calculated for each bird, using the method described by Cockrem and Silverin (2002a). The 'total integrated response' (Cockrem and Silverin, 2002a) is a measure of the amount of corticosterone secreted during the sampling period and is expressed as nglmhnin. A second measure called the 'corrected integrated response' was also calculated by multiplying the corticosterone concentration at time 0 by the duration of experiment and subtracting this from the total integrated response area. The area of the curves for testosterone, oestradiol, and LH which had negative responses were also calculated for each bird. Areas under the curve were calculated using the trapezoid rule using GraphPad Prism 4.0 (GraphPad Software Inc.). 2.2.6 Statisti cal analyses All results are presented as the mean ± SE (standard error) or individual data where appropriate. All the data were examined for normality using Shapiro-Wilks tests 52 Chapter Two and for equality of variance using Levene's homogeneity of variance test. Hormone data were logIo transformed to avoid heteroscedasticity. If variances were homogenous then groups were compared with repeated measures two-way analyses of variance (ANOY A) with time and treatment as the grouping factors. Comparisons between times within each treatment and between treatments for each time were examined with post-hoc repeated measures contrasts. Data not complying with normality or variance criteria were compared using a Kruskal-Wallis non-parametric test followed by Mann-Whitney U independent group comparisons tests, or Friedman's test for repeat measures data. All comparisons of means were conducting using SPSS ™ 1 1 .0 (SPSS Inc., Illinois) or Systat ™ 8.0 (SPSS Inc., Illinois). 2.3 Resu lts 2.3 .1 Experiment 1 - Acute response to a s i ng le subcutaneous cort icosterone i njection in 8 week o ld male Japanese quai l 2.3.1 . 1 Gross morphology The seven week old male quail weighed 187.9 - 248.1 g (mean 214.4 ± 4.0 g; Figure 2.1a) when collected, with no differences between the three randomly assigned treatment groups (F2,18 = 0.385, P = 0.686). There was a slight but significant increase (FI,I8 = 41 .485, P < 0.001) in body weight across all groups by the end of the experiment ranging from 200.0 - 249.1 g (mean 226.2 ± 3.3 g) . Cloacal gland areas were large in all groups, with no significant increases in area during the experiment and no effect of treatment (F2,I8 = 1 .362, P = 0.281; Figure 2. 1b). There were no differences in testis weight between groups at the end of the experiment (F2,I8 = 0.374, P = 0.694; Figure 2.1c) . The testes were the same size as testes of adult birds held on long days. 2.3.1 .2 P lasma corticosterone Corticosterone declined after the first sample in all the untreated birds and then Chapter Two 53 remained relatively constant or increased, whereas there were no consistent changes in oil treated birds (Figure 2.2) . Corticosterone rose to a peak (221 .56 - 382.71 nglml) at 0.5 h in all birds injected with corticosterone, declined until 6 h after the injection and then remained constant. There was no relationship between body weight of the injected birds and peak corticosterone concentrations (r 0.059, p = 0.600). Corticosterone varied between treatments and times, with a significant interaction between treatment and time (see Table 2.1 for all statistics for this analysis). Corticosterone concentrations were initially low in the three groups of birds (Figure 2.3a). Mean corticosterone concentrations in the untreated birds decreased 0.5 h after the initial sample, rose significantly from 1 .5 to 3.0 h and remained constant and low thereafter, whereas there were no significant changes in corticosterone in the oil treated birds. Corticosterone in the birds injected with 1 .2 mg corticosterone increased to high concentrations at 0.5 h (289.10 ± 20.56 nglml), declined to low concentrations at 6 h and remained low at 12 h. Corticosterone concentrations at 6 and 12 h did not differ from initial concentrations. There was a significant difference between groups in the total integrated corticosterone response (F2,20 = 623.632, P < 0.001; Figure 2.5a), with the corticosterone treated group having a significantly greater response than the untreated and oil control groups (post-hoc Bonferroni test, p < 0.001 for both groups) . It was not possible to calculate a corrected integrated corticosterone response for either control group as corticosterone concentrations dropped below 0 h values during the course of the experiment. 54 250 -0) - 1 50 400 -N E E - ro Q.) L-ro "0 300 c:: ro Ol � ro 0 (3 200 8 -0) - -..c:: 0) .Ci) 3 en 6 � en Q.) - "0 Q.) L- ·co 0.. 4 Untreated Oil Corticosterone Untreated Oil Corticosterone I I I Untreated Oil Corticosterone Group L-------,I One week before treatment L-------,I One day after treatment Chapter Two (a) (b) (c) Figure 2.1 Body weight (a), c10acal gland area (b), and paired testis weights (c) in untreated male quail and in quail treated with oil or 1 .2 mg corticosterone. Results are plotted as mean ± standard error. "'C Q,) - nl Q,) � - c :J '0 0126 _35 Al40 "v/l� j f� , .. ........ .. .. ..... :;; . . . .. . . . . . .. .. .. l . . ... . . .. .. . _ . . . . .. ......... . , . . .. ... .. _ .. . . . ... . . . . ...... . .... . Al43 Al46 � 1 �\r--:------ .. .... .... -.. . .. _ . . . . .... . . .. _ --_ .... .. .. . .. .. .. . .. _71 � '- /' '-.. ..... .. _ .. .. .. .. .. . .. . . . . .. .. . . . . '0 I I _3 I � �. � -" /- -"-.. � ••• "!."".� . . .. .. .. ... - - • • - . .. -� -i Cl) E '01 � � '0 "2 1' �', 2 "0 "2 1' '�c � '0 '2 1' '�' � "0 "2 1' � ' , � "0 "2 1' � " � "0 "2 1 �" � "0 ,'2 r '0 S __ 0127 _304 .36 Al41 _ Al48 Al49 � � e e � � 0 ::l Cl) Q) c o o Q; �lr� \r- l� �----------- � � �. � ..... (/) 8 ..... L-o Q,) U 400 C o � Q,) - (f) o .� 1:: o (,) 300 200 '00 .. . . . .... . . .. -... . .. - ..... -._ .. ......... .. • • cr .. ... .. . ... .. - .. . ... .. .... . . . .. . . -.... .. . .. .. ... .. . . .. .. -. . . . ... .. .. .. .. .. .. ... . ... . . '" ,.' _ .... .. . ... . . .. .. .. .. .. .. .. .. .. . . ao .. ... .. _ .. .. .. .. -0- __ .. .. .. .. .. __ • •• r I 0 --c,--,-�-�-,-�--,-J", , ; I ! . • 1 1. . " , I , I ', • I " , . i . ::J 10 12 0 2 10 12 0 2 10 12 0 2 10 12 0 2 10 12 0 2 10 12 0 2 10 12 (Q 2 0128 .. .. - . . 10 12 #29 1 r �o 1 r I �9 1 r 11'42 1 r t45 1 r 11'47 r '0 3" .. .. .. .... .. ....... . - - - _ .. .., 10 12 .. . .. .. . 0 10 1 2 00 _ _ _ · ·-- - .. .. .. • .. - · · .... 10 12 Time ( hour) -Corticosterone Testosterone . -. LH 10 1 2 1 0 1 2 1 0 1 2 e ...... Figure 2.2 Plasma corticosterone, testosterone and LH in untreated male quail and in quail given a single injection of oil or 1 .2 mg corticosterone. Note that the scale for the corticosterone axis differs between treatments. Results are plotted as raw data. n ::r tIl "'d ..... (1) >-t >-3 � 0 U1 U1 56 E -- 300 200 1 00 g> 50 Q) § 20 ..... Q) U5 1 0 8 :;:::; ..... o () 5 2 .00- . Untreated Oi l - Corticosterone I , - - .. ,.['; - . - ,.. - - - - � - - - - - - - - - - - - - - _ .... \>-- -.,... 1�._----._----._----._----._----,_----._-- E --O'l c:: '--'" Q) c:: 0 ..... Q) ...... (Jl 0 ...... (Jl Q) I- E --O'l c:: '--'" I ...J 6 5 4 3 2 ,t.. ..t ··· . . . . . . . . . , \ Y ,,' , , , .. , .. , .... , .. , .. .. .. .. .. .. .. .. .. o�.-----.-----.-----�-----.-----.----��- 2 o o 2 - - - - - - - - - - - - - - - - - - - 4 6 8 lime (hour) 1 0 1 2 Chapter Two (a) (b) (c) Figure 2.3 Plasma corticosterone (a), testosterone (b), and LH (c) in untreated male quail and in quail treated with oil or 1 .2 mg corticosterone. Corticosterone concen tration is plotted against a loglo vertical axis. Results are plotted as mean ± standard error. Chapter Two 57 6 5 E -- 4 O'l c a> 3 c 0 .... a> 1il 2 0 1il a> I- 0 6 5 E -- 4 O'l c --- a> 3 c 0 .... a> 1il 2 0 1il a> I- 0 Birds with < 1 ng / ml testosterone at 0 h " ,0. .. .. .. __ ... ... , ... ... ... ... ... .. -... ... ... .. .. " . p' " " . f ... ... 0.. ,,' . . . · . . · . . · . · . o " \ , , ' .-..,..... . \/ ------� -�. • 0 2 4 6 8 Time (hour) --- 1 0 1 2 (a) ..0- - . Untreated (n = 1 ) O i l ( n = 2 ) - Co rticosterone ( n =2) Birds with � 1 ng / ml testosterone at 0 h (b) 0 2 ,/ _ .. ... - . . . . . . 4 6 8 Time (hour) 1 0 1 2 • • 0-• . Untreated ( n=6) Oil ( n =5) - Corticosterone ( n=5) Figure 2.4 Plasma testosterone in male quail with initial concentrations < 1 nglml (a) or � 1 nglml (b). Quail were untreated or given oil or corticosterone injections. Results are plotted as mean ± standard error. 58 Chapter Two 500 I Corticosterone (a) 400 300 200 ...- 1 00 C E E 0 - 50 0> (b) c Testosterone -- ID 40 Cl) c I 0 I Cl. Cl) 30 ID I- I ID c 20 0 E I- 0 1 0 ..c "'C ID ....... 0 co I- 20 0> ID LH (c) ....... C 1 5 I 1 0 I I 5 o L-____________________________________ __ Untreated Oil Corticosterone Group Figure 2.5 Total integrated responses for corticosterone (a), testosterone (b) and LH (c) in untreated male quail and in quail treated with oil or 1 .2 mg corticosterone. Results are plotted as mean ± standard error. Chapter Two 59 Table 2.1 Two-way repeated measures ANOVA for corticosterone concentrations in untreated male quail and in quail treated with oil or 1 .2 mg corticosterone. Plasma corticosterone Effect F df p Treatment 189.851 2,1 8 < 0.001 • • Time 81 .796 5,90 < 0.001 • • Interaction of treatment and time 122.280 10,90 < 0.001 • • Comparisons within treatments for each treatment o vs 0.5 h Untreated 12.685 1,18 0.002 • • Oil 0.702 1,18 0.413 Corticosterone 1 1 50.135 1,18 < 0.001 • • 0.5 vs 1 .5 h Untreated 0.346 1,18 0.564 Oil 1 .1 75 1,18 0.293 Corticosterone 46.003 1,18 < 0.001 • • 1 .5 vs 3 h Untreated 5.122 1,18 0.036 • Oil 0.102 1,18 0 .753 Corticosterone 194.658 1,18 < 0.001 • • 3 vs 6 h Untreated 0.367 1,18 0.552 Oil 1 .721 1 ,18 0.206 Corticosterone 172.807 1,18 < 0.001 • • 6 vs 1 2 h Untreated 1 .084 1,18 0.31 1 Oil 0.300 1,18 0.590 Corticosterone 4.039 1,18 0.060 o vs 6 h Untreated 3 .295 1,18 0.086 Oil 0.003 1,18 0.954 Corticosterone 0.358 1,18 0.557 o vs 12 h Untreated 0.823 1,18 0.376 Oil 0.438 1,18 0.516 Corticosterone 3 .750 1,18 0.069 Note. The first three rows show the results of the two-way repeated measures ANOVA for the main effects and their interaction. The remaining rows show the results of post-hoc comparisons of times for each treatment and comparisons of treatments at each time. Significant differences (p < 0.05) are marked with a single asterisk, while highly significant differences (p < 0.01 ) are marked with a double asterisk. 60 Chapter Two Table 2.1 cont Two-way repeated measures ANOVA for corticosterone concen tra tions in untreated male quail and in quail treated with oil or 1 .2 mg corticosterone. Plasma corticosterone Comparisons between treatments for each time F df P O h Untreated vs Oil 1 .513 1,18 0.235 Untreated vs Corticosterone 1 .080 1,18 0.312 Oil vs Corticosterone 0.036 1,18 0.851 0.5 h Untreated vs Oil 0.468 1,18 0.502 Untreated vs Corticosterone 808.564 1,18 < 0.001 ** Oil vs Corticosterone 770.1 88 1,18 < 0.001 ** 1 .5 h Untreated vs Oil 0.225 1,18 0.641 Untreated vs Corticosterone 1 069.055 1,18 < 0.001 ** Oil vs Corticosterone 1038.240 1,18 < 0.001 * * 3 h Untreated vs Oil 0.920 1,18 0.350 Untreated vs Corticosterone 92.683 1,18 < 0.001 * * Oil v s Corticosterone 1 12.074 1,18 < 0.001 * * 6 h Untreated vs Oil 0.256 1,18 0.619 Untreated vs Corticosterone 2.699 1,18 0.1 18 Oil vs Corticosterone 1 .294 1 ,18 0.270 12 h Untreated vs Oil 0.01 1 1,18 0.918 Untreated vs Corticosterone 2.322 1,18 0.145 Oil vs Corticosterone 2.014 1,18 0.173 Chapter Two 61 2.3.1 .3 Plasma testosterone Testosterone declined after the first sample in all but one of the birds when the initial plasma concentrations were greater than 1 nglml (Figure 2.2) . Testosterone declined until 3 h, then increased until 6 h in all groups and then remained constant or continued to increase. Testosterone was less than 1 nglml in two birds in the oil group and two in the corticosterone treated group for the first three hours of the experiment and remained low (Figure 2.4a). There was no effect of treatment on testosterone, but it did vary between time and there was a significant interaction between treatment and time (see Table 2.2 for all statistics for this analysis). Testosterone concentrations varied from 0.36 to 6.38 nglml in birds at 0 h, but there were no significant differences between the groups (Figure 2.3b) . Mean testosterone concentrations in the oil and corticosterone treated birds decreased 0.5 h after the initial sample, and continued to decrease from 0.5 to 1 .5 h in the oil group. Mean testosterone concentrations in untreated birds decreased from 0.5 to 3 h, and testosterone increased to initial concentrations in both control groups from 3 to 6 h. Testosterone in the birds injected with 1 .2 mg corticosterone remained low between 0.5 and 3 h, then increased from 3 h and did not differ from initial concentrations at 12 h. There was no difference between the groups in the total integrated testosterone response (F2,2o = 1 . 126, P = 0.346; Figure 2.5b), and no differences between birds treated with 1 .2 mg corticosterone and the untreated and oil groups (post-hoc Bonferroni test, p = 0.458 and p = 1 .000 respectively). It was not possible to calculate a corrected integrated testosterone response for any of the groups as testosterone concentrations dropped below 0 h values during the course of the experiment. 62 Chapter Two Table 2.2 Two-way repeated measures ANOVA for testosterone concentrations in untreated male quail and in quail treated with oil or 1 .2 mg corticosterone. Plasma testosterone Effect F df p Treatment 1 .866 2,12 0.183 Time 30.781 5,90 < 0.001 • • Interaction of treatment and time 2.995 1 0,90 0.006 •• Comparisons within treatments for each treatment o vs 0.5 h Untreated 0.181 1,18 0.675 Oil 24.474 1,18 < 0.001 • • Corticosterone 51.572 1,18 < 0.001 • • 0.5 vs 1 .5 h Untreated 9.728 1,18 0.006 • • Oil 10.761 1,18 0.004 • • Corticosterone 2.322 1,18 0.145 1 .5 vs 3 h Untreated 12.753 1,18 0.002 • • Oil 0.216 1,18 0.647 Corticosterone 0.232 1,18 0.636 3 vs 6 h Untreated 20.093 1,18 < 0.001 • • Oil 9.528 1,18 0.006 • • Corticosterone 3.678 1,18 0.071 6 vs 12 h Untreated 0.766 1,18 0.393 Oil 0.022 1,18 0 .884 Corticosterone 16.733 1,18 0.001 • • o vs 3 h Untreated 28.188 1,18 < 0.001 • • Oil 19.983 1,18 < 0.001 • • Corticosterone 25.340 1,18 < 0.001 • • o vs 6 h Untreated 0.011 1 , 18 0.917 Oil 0.530 1,18 0.476 Corticosterone 0.143 1,18 0.036 • o vs 12 h Untreated 1 .820 1,18 0.194 Oil 2.353 1,18 0.142 Corticosterone 2.712 1,18 0.105 Note. The first three rows show the results of the two-way repeated measures ANOVA for the main effects and their interaction. The remaining rows show the results of post-hoc comparisons of times for each treatment and comparisons of treatments at each time. Significant differences (p < 0.05) are marked with a single asterisk, while highly significant d ifferences (p < 0.01) are marked with a double asterisk. Chapter Two 63 Table 2.2 cont Two-way repeated measures ANOVA for testosterone concentrations in untreated male quail and in quail treated with oil or 1 .2 mg corticosterone. Plasma testosterone Comparisons between treatments for each time F df P O h Untreated vs Oil 0 .124 1 ,18 0.729 Untreated vs Corticosterone 1 .286 1 ,18 0.272 Oil vs Corticosterone 0.61 1 1 ,18 0.445 0.5 h Untreated vs Oil 2.680 1 ,18 0.119 Untreated vs Corticosterone 9.834 1,18 0.006 • • Oil vs Corticosterone 1 .246 1 ,18 0.151 1 .5 h Untreated vs Oil 3.475 1,18 0.079 Untrea ted vs Corticosterone 7.422 1 ,18 0.014 • Oil vs Corticosterone 0.740 1 ,18 0.401 3 h Untreated vs Oil 0.003 1,18 0.956 Untreated vs Corticosterone 0.929 1,18 0.348 Oil vs Corticosterone 1 .039 1,18 0.321 6 h Untreated vs Oil 0.377 1,18 0.547 Untreated vs Corticosterone 4.233 1,18 0.054 Oil vs Corticosterone 2.084 1,18 0.166 12 h Untreated vs Oil 2.537 1 ,18 0.129 Untreated vs Corticosterone 0.338 1,18 0.568 Oil vs Corticosterone 1 .023 1,18 0.325 2.3.1 .4 Plasma LH LH concentrations remained relatively constant for the duration of the experiment in all birds (Figure 2.2). LH did not vary between the treatments or between time, and there was no significant interaction between treatment and time (see Table 2.3 for all statistics for this analysis) . Initial plasma LH concentrations were low and ranged from 0.49 - 1 .67 ng/ml. LH concentrations in the untreated control group were higher than those of the other groups at 0 and 0.5 h, but not thereafter. There was no significant difference in the total in tegrated LH response between the groups (F2,20 = 1 .023, P = 0.379; Figure 2.5c) . It was not possible to calculate a corrected integrated testosterone response for any of the groups as LH concentrations dropped below 0 h values during the course of the experiment. 64 Chapter Two Table 2.3 Two-way repeated measures ANOVA for LH concentrations in untreated male quail and in quail treated with oil or 1 .2 mg corticosterone. Plasma LH Effect F df P Treatment 2.297 2,18 0.129 Time 1 .001 5,90 0.406 Interaction of treatment and time 1 .091 10,90 0.380 Comparisons within treatments for each treatment o vs 0.5 h Untreated 0.288 1,18 0.598 Oil 0.737 1,18 0.402 Corticosterone 1 .004 1,18 0.330 0.5 vs 1 .5 h Untreated 6.327 1,18 0.022 • Oil 2.278 1,18 0.149 Corticosterone 5.355 1,18 0.033 • 1 .5 vs 3 h Untreated 0.001 1 ,18 0.981 Oil 0.889 1,18 0.358 Corticosterone 1 .742 1 ,18 0.203 3 vs 6 h Untreated 1 .386 1 ,18 0.254 Oil 0.175 1,18 0.680 Corticosterone 3.451 1 ,18 0.080 6 vs 12 h Untreated 0.365 1,18 0.553 Oil 3.049 1,18 0.098 Corticosterone 0.001 1 ,18 0.979 o vs 6 h Untreated 0.038 1 ,18 0.847 Oil 1 .211 1,18 0.286 Corticosterone 0.343 1,18 0.565 o vs 12 h Untreated 0.350 1,18 0.562 Oil 1 .440 1,18 0.246 Corticosterone 0.350 1,18 0.562 Note. The first three rows show the results of the two-way repeated measures ANOVA for the main effects and their interaction. The remaining rows show the results of post-hoc comparisons of times for each treatment and comparisons of treatments a t each time. Significant differences (p < 0.05) are marked with a single asterisk, while highly significant differences (p < 0.01 ) are marked with a double asterisk. Chapter Two 65 Table 2.3 cont Two-way repeated measures ANOVA for LH concentrations in untreated male quail and in quail treated with oil or 1 .2 mg corticosterone. Plasma LH Comparisons between treatments for each time F df P O h Untreated vs Oil 6.121 1,18 0.024 • Untreated vs Corticosterone 4.539 1,18 0.047 • Oil vs Corticosterone 0.1 18 1,18 0.735 0.5 h Untreated vs Oil 10.350 1,18 0.005 • • Untreated vs Corticosterone 8.859 1,18 0.008 •• Oil vs Corticosterone 0.058 1,18 0.812 1 .5 h Untreated vs Oil 1 .780 1,18 0.199 Untreated vs Corticosterone 0.634 1,18 0.436 Oil vs Corticosterone 0.289 1,18 0.597 3 h Untreated vs Oil 0.181 1,18 0.676 Untreated vs Corticosterone 2.170 1,18 0.158 Oil vs Corticosterone 1 .098 1,18 0.309 6 h Untreated vs Oil 0.492 1,18 0.492 Untreated vs Corticosterone 0.763 1,18 0.394 Oil vs Corticosterone 0.030 1,18 0.865 12 h Untreated vs Oil 2.832 1,18 0.110 Untreated vs Corticosterone 0.072 1,18 0.791 Oil vs Corticosterone 2.000 1,18 0.174 2.3.2 Experiment 2 - Acute response to a s ing le subcutaneous corticosterone injection in 8 week old female Japanese quai l 2.3.2.1 Gross morphology The seven week old female quail weighed 204.7 - 277.6 g (mean 236.94 ± 4.72 g; Figure 2.6a) when collected, with no differences between the two randomly assigned treatment groups (F),)9 = 0.012, P = 0.914) and no change (F),)8 = 0.452, P = 0.510) in body weight in the two groups over the course of the experiment where body weights were 207.5 - 280.7 g (mean 238.3 ± 4.33 g) . All females had a palpable egg in the oviduct when they were collected from the supplier and continued to lay up to one egg a day until the end of the experiment. Ovary weight did not differ between the groups one day after the treatment (8.38 ± 2.65 in oil treated compared with 8.75 ± 2.77 g in corticosterone treated; 66 Chapter Two F},}9 = 0.213, P = 0.650; Figure 2.6b). All ovaries had normal yellow follicular hierarchies and were the same size as those of adult birds held on long days. 2 .3.2.2 Plasma corticosterone Corticosterone declined after the first sample in 7 out of 10 birds treated with oil and then generally increased between 0.5 and 3 h (Figure 2.7). Corticosterone rose to a peak (75.84 - 229.91 nglml) at 0.5 h in all birds injected with corticosterone, declined until 12 h after the injection and then remained constant. Corticosterone varied between treatment and between time, with a significant interaction between treatment and time (see Table 2 .4 for all statistics for this analysis) . Corticosterone concentrations were initially low in both groups of birds (Figure 2.8a). Mean corticosterone concentrations in the oil treated birds decreased 0.5 h after the initial sample, then increased and remained elevated from 1 .5 to 6 h before decreasing and remaining constant and low after 12 h. Corticosterone in the birds injected with 1 .2 mg corticosterone increased to high concentrations at 0.5 h (158.69 ± 9.63 nglml) and then declined to initial concentrations at 24 h. Corticosterone concentrations at 12 h were significantly higher than initial concentrations, but did not differ at 24 h. Birds treated with 1 .2 mg corticosterone had a significantly greater total integrated corticosterone response than birds in the oil treated group (F},}9 = 52.673, P < 0.001; Figure 2.9a) . It was not possible to calculate a corrected integrated corticosterone response for the oil treated group as corticosterone concentrations dropped below 0 h values during the course of the experiment. Chapter Two 250 240 ..- 0> -- 1: 230 0> ·w � >. 220 "0 o CO ..- 0> '-"' ...... .r::. 0> 2 1 0 200 1 0 .� 8 � � o 6 - - - - - - I I I I Untreated Corticosterone I I Untreated Corticosterone Group c=:::J One week before treatment c=:::J After treatment 67 (a) (b) Figure 2.6 Body weight (a) and ovary weight (b) in untreated female quail and in quail treated with 1 .2 mg corticosterone. Results are plotted as mean ± standard error. Oi l group 10 _59 1122 _24 2 1:./,----__ _ .�u ..-.. E _ 10 Ol C '-' ID C o 12 16 20 24 0 -.u 12 16 20 24 0 12 16 20 24 0 500 1IIl7 W33 400 300 � ID (j) o (.) 6; , � .. 41 , A. " ' , . 2 1, � ,.. I� : :: ..... � o U 300 200 100 12 16 20 24 0 � 12 16 20 2" 0 12 16 20 24 0 1IIl3 ". 12 16 20 24 0 �4 12 16 20 24 12 16 20 2" �7 .58 _71 _34 _31 " , , � � � 12 16 20 24 0 12 16 20 24 0 12 16 20 24 0 12 16 20 24 300 _32 - \ _70 ( ' 200 '\. 100 oil ,=: ..." 1 11 ', '::-:; i 1 .Jl.ll_-;=::::;:==;:="'T=""I 12 16 20 24 0 12 16 20 24 0 12 16 20 24 �8 �6 .72 500 400 300 200 100 500 400 300 200 100 500 400 300 200 100 ;::: ,I. 12 16 20 24 0 12 16 20 24 0 12 16 20 24 0 12 16 20 24 Time (hour) Corticosterone g roup -Corticosterone • Oestradiol 0 CD en .-., ru Cl. 0 ..-.. "0 (Cl - 3 '-' Figure 2.7 Plasma corticosterone and oestradiol responses in female quail treated with oil or 1 .2 mg corticosterone. Note that the scale for the corticosterone axis differs between treatments. Results are plotted as raw data. Q'\ 00 n ::r' III � .... rtI '"i ...., � o Chapter Two 200 1 00 ....-.. E 50 -O'l C 0) 20 c 0 L- 0) 1 0 -U) 0 0 - 5 L- 0 () 2 1 400 300 E - O'l Cl. '-'" o 200 "'C ro L-- U) ID o 1 00 - � · Oil - Corticosterone ,� ... ? - - - -9- ... ... ... ' - - . . � - - � - - - - - - - - - - - - - - - - - - - 0 2 4 6 8 1 0 1 2 1 4 1 6 .. .. ... ... ... ... 1 .. .. .. .. ... ... ... T .. .. .. 1 8 20 .. .. .. .. .. .. 22 24 .. .. .. .. .. .. o 2 4 6 8 1 0 1 2 1 4 1 6 1 8 20 22 24 Time (hour) 69 (a) (b) Figure 2.8 Plasma corticosterone (a), oestradiol (b) in female quail treated with oil or 1 .2 mg corticosterone. Corticosterone concentration is plotted on a log1o axis. Results are plotted as mean ± standard error. 70 m Cl) c o Cl. Cl) m � m c ­o c � .- 800 600 m E en --..: 400 o E u __ � O'l � c 8 - -0 200 m ..... co � Cl m ..... c 0 m Cl) c 8000 � 6000 m � - '2 o ._ � .s 4000 ..... E CI) __ m O'l o C -0 - 2 2000 co � Cl m ..... c o Chapter Two (a) I (b) I I L-________ �--------------------�--------- Oil Corticosterone Groups Figure 2.9 Total integrated responses for corticosterone (a) and oestradiol (b) in female quail treated with oil or 1 .2 mg corticosterone. Results are plotted as mean ± standard error. Chapter Two 71 Table 2.4 Two-way repeated measures ANOVA for corticosterone concentrations in quail treated with oil or 1 .2 mg corticosterone. Plasma corticosterone Effect F df P Treatment 483.720 1,18 < 0.001 •• Time 95.208 6,108 < 0.001 •• Interaction of treatment and time 83.501 6,108 < 0.001 •• Comparisons within treatments for each treatment o vs 0.5 h Oil 2.213 1,18 0.154 Corticosterone 641 .039 1,18 < 0.001 • • 0.5 vs 1 .5 h Oil 5.149 1,18 0.036 • Corticosterone 2.723 1 ,18 0 .116 1 .5 vs 3 h Oil 0.1 1 5 1,18 0.698 Corticosterone 19.348 1,18 < 0.001 • • 3 v s 6 h Oil 0.490 1,18 0.493 Corticosterone 66.776 1 ,18 < 0.001 •• 6 vs 12 h Oil 15.331 1,18 0.001 • • Corticosterone 59.141 1,18 < 0.001 • • 12 vs 24 h Oil 0.002 1 ,18 0.965 Corticosterone 26.378 1 ,18 < 0.001 • • o vs 12 h Oil 2.292 1,18 0.147 Corticosterone 79.646 1 ,18 < 0.001 • • o vs 24 h Oil 1 .363 1,18 0.258 Corticosterone 3.252 1 ,18 0.088 Comparisons between treatments for each time F df P Oil vs corticosterone O h 0.284 1,18 0.600 0.5 h 785.190 1,18 < 0.001 • • 1 .5 h 349.392 1 ,18 < 0.001 • • 3 h 247.606 1,18 < 0.001 • • 6 h 1 53.785 1,18 < 0.001 • • 12 h 64.554 1,18 < 0.001 • • 24 h 3.602 1,18 0.074 Note. The first three rows show the results of the two-way repeated measures ANOVA for the main effects and their interaction. The remaining rows show the results of post-hoc comparisons of times for each treatment and comparisons of treatments at each time. Significant differences (p < 0.05) are marked with a single asterisk, while highly Significant differences (p < 0.01 ) are marked with a double asterisk. 72 Chapter Two 2.3.2.3 Plasma oestradiol Oestradiol declined after the first sample in 9 out of 10 birds treated with oil, and 7 out of 10 birds treated with 1 .2 mg corticosterone (Figure 2.7) . Oestradiol increased between 3 and 24 h in all corticosterone treated birds, while there was no consistent pattern in oestradiol concentrations after 0.5 h in the oil treated group. Oestradiol varied in time but not between treatment, and there was a significant interaction between treatment and time (see Table 2.5 for all statistics for this analysis). Oestradiol concentrations were initially lower in the group treated with corticosterone (Figure 2.8b). Mean oestradiol concentrations in birds given an oil injection declined between 0 and 3 h, but did not differ from initial oestradiol concentrations after 6, 12 and 24 h. Mean oestradiol concentrations in birds treated with 1 .2 mg corticosterone remained rela tively constant between 0.5 and 3 h, rose significantly from 3 to 12 h and remained constant and high thereafter. Concentrations at 12 and 24 h were significantly higher than initial oestradiol concentrations. There was no significant difference in the total integrated oestradiol response between the groups (F],]9 = 2.498, P = 0.131 ; Figure 2.9b). It was not possible to calculate a corrected integrated oestradiol response for either group as corticosterone concentrations dropped below 0 h values during the course of the experiment. Chapter Two 73 Table 2.5 Two-way repeated measures ANOVA for oestradiol concentrations in quail treated with oil or 1 .2 mg corticosterone. Plasma oestradiol Effect F df p Treatment 3.570 1,18 0.075 Time 6.688 6,108 < 0.001 • • Interaction of treatment and time 3.714 6,108 0.008 •• Comparisons within trea tments for each treatment o vs 0.5 h Oil 1 .620 1,18 0.219 Corticosterone 3.492 1,18 0.078 0.5 vs 1 .5 h Oil 0.234 1,18 0.634 Corticosterone 1 .226 1,18 0.283 1 .5 vs 3 h Oil 2.083 1,18 0.166 Corticosterone 0.286 1,18 0.600 3 vs 6 h Oil 2.007 1,18 0.174 Corticosterone 4.472 1 ,18 0.049 • 6 vs 12 h Oil 2.190 1,18 0.1 56 Corticosterone 5.131 1,18 0.036 • 12 vs 24 h Oil 12.812 1 ,18 0.002 .. Corticosterone 0.127 1,18 0.726 o vs 6 h Oil 0.691 1,18 0.417 Corticosterone 0.396 1,18 0.537 o vs 12 h Oil 3.794 1,18 0.067 Corticosterone 5.928 1,18 0.026 • o vs 24 h Oil 0.067 1,18 0.799 Corticosterone 7.099 1,18 0.016 • Comparisons between treatments for each time F df P Oil vs corticosterone O h 3.607 1,18 0.074 0.5 h 4.774 1,18 0.042 • 1 .5 h 5.552 1 ,18 0.030 • 3 h 2.021 1 ,18 0.172 6 h 0.472 1 ,18 0.501 1 2 h 8.496 1 ,18 0.009 • • 24 h 0.535 1 ,18 0.474 Note. The first three rows show the results of the two-way repeated measures ANOVA for the main effects and their interaction. The remaining rows show the results of post-hoc comparisons of times for each treatment and comparisons of treatments at each time. Significant differences (p < 0.05) are marked with a single asterisk, while highly significant d ifferences (p < 0.01) are marked with a double asterisk. 74 Cha pter Two 2.4 D iscussion The administration of a single 1 .2 mg corticosterone injection to male quail caused a rapid increase in plasma corticosterone concentrations. Plasma corticosterone remained elevated 3 hours later and had returned to initial concentrations after 6 hours. Testosterone concentrations declined over 3 hours in both treated and control birds then returned to initial concentrations after 6 hours in control birds and after 12 hours in treated birds. LH did not change in any of the groups. Oestradiol declined after treatment then returned to initial concentrations after 6 hours in oil and corticosterone treated females. Oestradiol in both groups of treated females returned to initial concentrations whilst corticosterone remained elevated . 2.4.1 Ma le quai l 2.4. 1 . 1 Corticosterone In the current experiment corticosterone concentrations remained elevated for a t least 3 hours whereas Boyd (2000) reported that plasma corticosterone in males was elevated for at least 6 hours after a single 1.2 mg corticosterone injection. Plasma corticosterone concentrations 3 hours after treatment ranged between 70 and 220 ng/ml in the Boyd (2000) s tudy, while they were 15 to 54 ng/ml in the current experiment. The corticosterone was delivered as a suspension in oil with stirring necessary to maintain the suspension's uniformity (Boyd, 2000). The differen t corticosterone profiles in the two studies could be due, at least in part, to differences in the actual amount of corticosterone delivered. Alternatively, the two groups of birds may have differed in their rates of clearance of corticosterone. The first blood sample after the corticosterone injection was collected at 0.5 hours in the present study, compared with 3 hours by Boyd (2000). Beuving and Yonder (1978) and Davison et al. (1980) showed that plasma corticosterone increased 3.5 minutes after an injection of ACTH or corticosterone in chickens. Corticosterone in quail was elevated 0.5 hours after the injection, and it is likely that sampling closer to the injection time would reveal an earlier and higher corticosterone peak than observed in this study. Chapter Two 75 Despite all birds being similar in weight and receiving the same amount of corticosterone, there were a range of peak concentrations measured at 30 minutes. A number of factors are likely to have influenced and caused this variation between the birds such as, body weight, metabolic rate, the amount of corticosterone delivered, and the concentration of corticosterone binding globulin (CBG). All are factors that contribute to different rates of corticosterone clearance from the plasma. Plasma corticosterone remained low in both control groups and there was no pattern of secretion during the experiment. Corticosterone secretion in quail follows a circadian rhythm, it remains low during hours of light, increases at the onset of darkness remaining elevated before decreasing at the onset of light (Boissin and Assenmacher, 1970). However, plasma corticosterone concentrations have also been reported to peak in late morning and then decline in other quail (Kovacs and Peczely, 1983; Kovacs et al., 1983) . In the current study, plasma corticosterone decreased in the few hours after the lights went on, and then remained relatively constant for the duration of the experiment. Corticosterone concentrations did not differ between the untreated and oil treated birds, showing the injection alone did not elicit a corticosterone response 0.5 hours later. 2.4. 1 .2 Testosterone Mean testosterone concentrations in all groups decreased after 30 minutes and further decreased or remained low at 1 .5 and 3 hours before rising at 6 hours. Testosterone concentrations in the corticosterone treatment group were lower between 0.5 and 3 hours, although this is attributed to this group having lower initial corticosterone concentrations. The results of the current study differ from Boyd (2000) who showed that testosterone concentrations decreased 3 hours after a 1 .2 mg corticosterone injection, but did not change in oil treated quail. In contrast, Heiblurn et al. (2000) showed testosterone concentrations increased within 30 minutes of an injection of ACTH in male chickens, which conflicts with previous studies showing declines in plasma testosterone during acute and chronic stressors (reviewed by Deviche, 1 983). The decrease in testosterone concentration in the current experiment occurred over 1 .5 - 3 hours and was unlikely to be circadian in nature. Ottinger and Brinkley (1979) showed that testosterone in quail is highest in 76 Chapter Two the early morning and slowly decreases over 9 hours to a minimum in the late a fternoon before rapidly increasing in the evening. Birds in the current study were bled at 0, 0.5 and 3 hours, whereas Boyd (2000) bled birds at 0 and 3 hours. The extra handling in the current study may have caused the decrease in plasma testosterone concentrations in all groups as seen in hens handled and blood sampled every 15 minutes (Wilson et al., 1979) . An effect of repeated handling and sampling by wing vein venipuncture on plasma testosterone has also been shown in other studies of chickens (Eskeland and Blom, 1 979; Johnson, 1981) . The authors of all three studies considered handling stress to be the cause of the decline in testosterone secretion without defining stress or considering a mechanism for stressors to inhibit testosterone secretion. A decrease in testosterone secretion following a handling stressor is said to be caused by an increase in corticosterone or cortisol with handling or restraint (Orr and Mann, 1 990; Orr and Mann, 1992) . Corticosterone in rats inhibits the conversion of 17a­ OH progesterone to androstenedione, an androgen that is then converted into testosterone (Orr and Mann, 1990; Orr and Mann, 1992), and Deviche et al. (1982) showed that corticosterone injections in Japanese quail reduced the production of androstenedione which led to a decrease in testosterone. Satterlee et al. (2002) showed that genetically selecting quail for reduced corticosterone responses to mechanical restraint accelerated puberty. This was assessed by measuring the area and volume of the androgen-dependent cloacal gland, which was larger in birds with a low corticosterone response compared with quail with a high response. The decline in testosterone in the control birds means that it is not possible to determine from this data whether corticosterone inhibited testosterone. However, a transient corticosterone increase due to handling might have contributed to the decline in testosterone in the control birds. The pattern of testosterone secretion varied considerably between birds. A few birds had low initial testosterone concentrations and testosterone remained low for the duration of the study. All of these birds had large testes, large cloacal glands and were producing cloacal foam. Chapter Two 77 2.4.1 .3 LH There was no change in plasma LH concentrations after birds were treated with corticosterone or in control birds, as found by Boyd (2000). Deviche et al. (1980) also showed that plasma LH and FSH remained unchanged whilst testosterone decreased in male ducks given a corticosterone injection. Handling did not affect LH in the present study, although Wilson and Sharp (1975) found that plasma LH decreased in hens blood sampling at 10 and 15 minute intervals. The prolonged elevation of plasma corticosterone using osmotic pumps has been shown to cause ovarian regression and reduce plasma LH and oestradiol concentrations (Etches et al., 1984; Williams et al., 1985; Petitte and Etches, 1989) . The decreases in plasma LH and oestradiol could be reversed by the administration of GnRH and PMSG (pregnant mare serum gonadotrophin) respectively (Etches et al., 1984; Petitte and Etches, 1989) . The responsiveness of the pituitary and gonad to exogenous GnRH and a gonadotrophin show that prolonged elevation of corticosterone acts in some part at the hypothalamus and pituitary gland to inhibit the reproductive axis in chickens. Corticosterone can inhibit LH secretion from quail pituitary cells in culture (Connolly and Callard, 1987). However, corticosterone has many influences in the avian reproductive system and injections can either cause a decrease or increase in plasma LH concentrations in laying hens depending on the timing of the injection within the ovulatory cycle (Etches and Croze, 1983). Elevated corticosterone in the treated birds may have altered the pulsatile secretion of LH without a change in mean LH concentrations. LH secretion is pulsatiJe in the male Japanese quail with 6 - 10 pulses per day (Gledhill and Folletl, 1976) that are 90 - 120 minutes in duration (Wilson and Sharp, 1 975) . Acute changes in cortisol and ACTH in mammals can decrease LH pulse frequency without affecting mean LH concentrations (Daley et al., 1999; Van Lier et al., 1999; Smith and Dobson, 2002), and this could also occur in birds. The secretion of LH from the pituitary is in part controlled by negative feedback by testosterone (Gibson et al., 1975; Davies et al., 1976; Deviche et al., 1979; Deviche 78 Chapter Two et al., 1980; Knight et al., 1983). The permanent depression of testosterone following gonadectomy results in an increase in LH secretion in birds (Gibson et al., 1975; Wilson and Sharp, 1975; Deviche et al., 1980) . The decline in testosterone concentration in both the current study and that of Boyd (2000) did not however lead to an increase in LH secretion. 2.4.2 Female quai l 2.4.2. 1 Corticosterone Administration of a 1 .2 mg corticosterone injection to female Japanese quail resulted in a significant increase in corticosterone concentrations after 30 minutes followed by a gradual decline, with corticosterone still elevated 12 hours after the injection. The period of elevated corticosterone was longer than in the male quail in the previous experiment and found by Boyd (2000) . However, the current study was conducted after the male treatment study and can not be d irectly compared. Differences in peak corticosterone concentrations between the sexes are likely to be due to slight differences in treatment, and the fact that the corticosterone doses for either sex were identical can not be assumed. 2.4.2.2 Oestradiol Plasma oestradiol concentrations gradually decreased in both oil and corticosterone-treatment birds after the injection, then increased after 3 hours. As was the case in the male study, it is likely that the handling associated with blood sampling caused this decreased, where oestradiol concentrations are lowest in both groups after 3 hours. A decrease in oestradiol following a pre-ovulatary surge may also be a possible cause of the decline in oestradiol concentrations during the first 1 .5 hours after the injection. Oviposition occurred between 1400 - 1900 in all females while ovulation occurs within 15-30 minutes (Woodard and Mather, 1964) . However, this cause i s unlikely given that the time of oviposition varied greatly between females while the decrease in oestradiol after the injection was so synchronous within and between both groups. Chapter Two 79 The role of corticosterone in the ovulatory cycle in female birds makes studying the inhibitory effects of an acute increase in corticosterone difficult as the effect of corticosterone can be stimulatory or inhibitory to the reproductive axis depending on the stage of the ovulatory cycle that an injection is administered (Etches and Croze, 1983) . In contrast, an increase in corticosterone concentrations by 1 - 2 nglml for 14 days in chickens implanted with osmotic pumps resulted in a decrease in plasma oestradiol and LH, and a cessation in lay (Etches et al., 1984). The same dose of corticosterone administered for 14 days in immature hens prevented the increase in plasma LH and oestradiol that follow photostimulation (Petitte and Etches, 1 988). 2.4.3 Summary The aim of this study was to determine the effects of an acute increase in corticosterone on testosterone and LH in male and oestradiol in female quail. The steroids decreased in corticosterone-treated birds but also in birds that only received saline, whereas LH did not change. Gonadotrophin secretion (Etches et al., 1984; Williams et al., 1985; Petitte and Etches, 1989) has been previously shown to decrease in response to corticosterone treatment, although in all such cases corticosterone was delivered constantly over a period of days. Prolonged exposure to stressors over hours and days can have negative effect on breeding behaviours and the reproductive axis (Wingfield, 1994). Factors such as the species of bird, latitude, and type of stressor influence how the different levels of the reproductive axis respond to stressors, as reported in wild passerines (Astheimer et al., 2000). Astheimer et al. (2000) proposed that the reproductive axis can become unresponsive to the inhibitory effects of stressors in Arctic birds so that breeding is not disrupted in species with a short period available for breeding. The natural breeding season of Japanese quail in their original range is not known, but the absence of an LH response to a stressor in quail could indicate the presence of a similar mechanism to tha t proposed for Arctic birds. The aim of the study was not met because sex steroids fell in control as well as corticosterone-treated birds. This effect of a single handling event was unexpected, as Boyd (2000) showed no effect of handling on testosterone after 3 hours, and 80 Chapter Two testosterone in chickens declined only when blood sampling was repeated at < 5 minute intervals (Wilson et al., 1979; Eskeland and Blom, 1979; Johnson, 1981). An alternative method for the delivery of corticosterone to quail is the use of jugular cannulae. This method would allow hormone solution to be delivered and blood samples withdrawn remotely, thereby avoiding any effect of handling. However, cannulation is used only in larger birds such as chickens and turkeys and was not considered feasible for quail in our laboratory. Instead, the novel finding that a single handling event would reduce sex steroids was explored in detail in another study. The effects on corticosterone and testosterone of different periods of handling and of exposure to other stressors were quantified to determine relationships between corticosterone and testosterone responses. Cha pter Three Chapter 3 Adrenocortical and testosterone responses of Japanese quail to a range of novel st imuli 3 . 1 I ntroduction 81 Exposure to a stressor can have a negative effect on reproduction in vertebrates (reviewed by Rivier and Rivest, 1991; Tilbrook et al., 2000; Tilbrook et al., 2002; Wingfield and Sapolsky, 2003) . In the previous chapter, repeated blood sampling in Japanese quail was shown to cause a decrease in plasma testosterone without affecting LH secretion. The decline in plasma testosterone following serial blood sampling has been well documented in chickens (Wilson et al., 1979; Eskeland and Blom, 1979; Johnson, 1981) . The duration of manual restraint or other stressors needed to initiate a corticosterone response and decline in testosterone secretion had not been reported in quail and was therefore determined in the study described in this chapter. The corticosterone response to handling or mechanical restraint has been widely studied in a wide range of species such as quail (Boyd, 2000), chickens (Beuving and Vonder, 1978; Littin and Cockrem, 2001) and a variety of other free-living, wild species (SiIverin, 1998). Handling can cause an increase in corticosterone concentrations within two minutes in chickens (Beuving and Vonder, 1978) and one minute in ducks (Harvey et al., 1 980), and corticosterone increases within one minute of restraint in quail (Satterlee and Johnson, 1988). Plasma corticosterone has been shown to increase 45 seconds after the start of a stressor and increase rapidly for the first 10 minutes in chickens restrained for 60 minutes (Beuving and Vonder, 82 Chapter Three 1978). In Japanese quail, corticosterone increased to a peak four minutes after the start of restraint and remained relatively constant during 20 minutes of mechanical restraint (Satterlee and Johnson, 1988). However, the relationship between the length of the handling procedure and changes in plasma corticosterone have not been quantified in the quail. In the first part of the study the relationships between the duration of manual restraint, corticosterone and testosterone were determined . As testosterone was affected by the blood sampling procedure used in the previous chapter, separate groups of birds were sampled at each collection time, as described by Boyd (2000) . Another factor that can influence the size of the corticosterone response is the type of stressor. Handling (Boyd, 2000), mechanical restraint (Satterlee and Johnson, 1988), or social disruption (Satterlee et al., 1983) can initiate corticosterone responses in Japanese quail whereas testosterone responses to different stressors in Japanese quail have not been described. In the second part of the present study corticosterone and testosterone responses to manual restraint, mechanical restraint, and social stimuli were characterised. In this study, as in those described by Satterlee and Johnson (1988), mechanical restraint covers a series of events in which quail were captured from their home cage, transported to the mechanical restraint area, blood sampled, then mechanically restrained before being returned to their cage. 3.2 Methods and materials 3.2.1 Animals and housing 3.2. 1 . 1 Source and type of birds Seven week-old male Japanese quail (Coturnix coturnix japonica) were purchased from our supplier (Rangitikei Game Birds Ltd ., Bulls) . The. birds had been raised under a long-day photoperiod (15L:9D) at air temperatures of 20 to 25°C in mixed sex groups. The 7 week-old birds were sexually mature at time of purchase with all birds having large cloacal glands and cloacal foam present. Chapter Three 83 3.2.1 .2 H o u s i n g co n d itions Each bird was identified with a numbered leg band and housed in an individual cage measuring 20 cm (W) x 35 cm (D) x 24 cm (H) in animal rooms in the Veterinary Science building. Quail were held on a long day photoperiod (16L:8D; lights on from 0900 - 0100 h). An extractor fan provided ventilation for each room, and only one sex was held in a room during each experiment. A temperature control unit enabled air temperature to be maintained at 20°C. Light in each room was provided by two 75W incandescent light bulbs controlled by a 24 hour!7 day time switch (HPM Excel Light Switch and Timer, Cat XL770T) . The light intensity in the quail rooms was 130 lux in cages at the top, and 1 5 lux in cages near the floor of the room (MinoJta Illuminance meter) . 3.2.1 .3 Food and water Quail were provided with fresh water and food (Harvey Farms quail layer mash) ab libitum. The food was the same as that used by Rangitikei Game Birds. 3 . 2 . 2 Experimental design 3.2.2.1 Experi ment 1 - Q u a ntification o f the effect o f m a n u al restra i n t o n p l a s m a testoste rone a n d corti costero n e i n the J a panese q u a i l Seventy 7 week old male Japanese quail were held with ad libitum access to food and water under long days (16L:8D) at 20°C for 1 0 days. The birds were assigned into 2 groups; birds in one group (n=35) were bled at time 0 and handled for approximately 2 minutes (called 'untreated group'), the time taken to collect a single blood sample. Birds in the second group (n=35) were bled at time 0 and manually restrained until 5 min total handling time had elapsed from when they were first removed from their cages, birds were returned to their cage after this time had past. Manual restraint consisted of a bird being held in one hand by the experimenter. Birds were removed from their cages and taken to another room where they were held firmly to prevent movement. Blood samples were collected from each group of birds (n=7) at 15, 3D, 60, 120, or 240 min after the time 0 sample. Birds were returned to their cages between samples. 84 Chapter Three Individual sample groups for each sample time were required to provide a clear picture of what effect manual restraint has on plasma testosterone (as we had previously shown in Chapter 3) as repeated blood sampling in individual birds decreases plasma testosterone. Both groups of birds were housed for another 10 days under the same conditions. After 10 days, the blood sampling procedure described above was repeated with manual restraint for 10 or 15 minutes in the place of 2 and 5 min. Birds were euthanased by stunning and decapitation at the final sample time (240 min) so that the testes could be weighed and a terminal blood sample collected. Body weight and the area of the cloacal protuberance were measured before the treatment. Plasma concentrations of corticosterone and testosterone were measured in all blood samples. 3.2.2.2 Experiment 2 - Adrenocortical and testosterone responses to manual restra int, mechanical restra int and social stimul i in Japanese quai l 3.2 .2 .2 . 1 The effect of d ifferent stressors on the adrenocortical response Fifty 7 week old male Japanese quail were obtained from our normal supplier (Rangitikei Game Birds) and held with ad libitum access to food and water under long days (16L:80) a t 20°C for 10 days. All quail were housed in individual cages. After 10 days, the quail were assigned into five groups (n=10 per group). Birds in each group were given one of the following treatments, (a) 2 min of manual restraint by an experimenter (also called 'untreated group'), (b) 15 min of manual restraint by the same experimenter in the previous group, (c) 15 min of mechanical restraint that prevented movement, (d) 15 min in a cage with an unfamiliar quail, or (e) 15 min in a cage with an unfamiliar quail with birds separated by a wire mesh partition. Blood samples were collected from each bird at 0, 15, 30, 60, 120, and 240 minutes. Birds were returned to their cages between samples. Plasma corticosterone concentrations were subsequently measured by radioimmunoassay. Chapter Three 85 3.2 .2 .2 .2 The effects of manual and mechan ica l restra int on testosterone and corticosterone In the previous experiment (Section 3.2.2.2.1 ) 15 minutes of mechanical restraint caused a smaller and more variable corticosterone response than 15 minutes of manual restraint. The goal of the previous study was to identify a stressor that stimulated a similar but smaller response to that caused by manual restraint. All 50 birds from the previous study were housed for another 10 days under the same conditions then assigned into five groups (n= 10) . All birds were blood sampled at time 0 prior to the treatment, and then each group was sampled once more at IS, 30, 60, 120 or 240 minutes. Half of the birds received 15 min of manual restraint (n =25) while the other half received 15 min of mechanical restraint. This protocol was repeated 10 days later to increase the sample size to 10 for each group. Thirty five birds were held for another 10 days, then assigned into five groups (n = 7) and blood sampled at time 0 then each group was sampled once more at IS, 30, 60, 120 or 240 minutes without manual or mechanical restraint. These birds were untreated controls. Birds were euthanased at the end of the study, and terminal blood samples collected. Plasma concentrations of corticosterone and testosterone were measured in all blood samples. The experiments were conducted under a protocol approved by the Massey University Animal Ethics Committee. 3.2 .3 Data col lection 3.2.3.1 Administration of stressors 3 .2 .3 . 1 . 1 Manual restraint Birds were removed from their cage and the 0 min blood sample collected within 2 min. The 2 min manual restraint group were then returned to their cage. Birds in the other groups were then held by a handler for up to 5, 10 or 15 min of manual restraint then returned to their cages. 86 Chapter Three 3.2.3 . 1 .2 Mechanical restraint Individual quail were placed into a firm well ventilated plastic bag that prevented the bird from flapping its wings but still allowed the bird free movement of its head outside the bag. The bird was then placed inside a metal cage 6 cm (W) x 15 cm (D) x 9 cm (H) that was larger than the quail. Quail were left in an upright dorso-ventral position for 15 min. 3.2 .3 . 1 . 3 Exposure to an unfam il iar qua i l Birds were removed from their cage and a blood sample collected. The bird was then returned to it's home cage which measured 20 cm (W) x 35 cm (D) x 24 cm (H). A quail that had not been housed nearby was then either (a) introduced into the adjacent cage with a wire mesh partition physically separating them or (b) introduced into the cage with the quail that had been blood sampled. In both treatments, birds were exposed to each other for up to 15 min after the initial blood sample and the stocking density remained 1 bird / 0.07 m2• After the 15 min, the unfamiliar quail was removed and returned to it's home cage. 3.2.3.2 Tissue and blood samples Blood samples were collected by venipuncture of the brachial vein in the wing with a 25 g needle, with 200 ILl of blood collected into heparinised capillary tubes. All samples were collected within 2 minutes from the time the bird was removed from the cage. The final blood sample was collected when the birds were euthanased by stunning followed by decapitation. The blood was stored on ice until centrifuged at 2 000 g for 15 minutes at 4°C (Heraeus Christ 5000S refrigerated centrifuge). The plasma was removed and stored at -20°C until assayed. The width and height of the cloacal protuberance was measured (±0.1 mm) in males and body weight was measured (±0.1 g) in all birds on the day before the experimental treatment. The reproductive system was removed after euthanasia and the testes weighed (±0.001 g). Chapter Three 87 3.2.4 Hormone assays Corticosterone and testosterone concentrations in plasma were determined by radioimm unoassay. 3.2.4 . 1 Corticosterone Plasma corticosterone concentrations were determined by radioimmunoassay using the methodology outlined in Section 2.2.4. 1 . The inter-assay coefficients of variation for 9 assays were 9.7, 1 0.3, and 7.1 % for high, medium and low quality controls respectively. 3.2.4.2 Testosterone Plasma testosterone concentrations were determined by radioimmunoassay using the methodology outlined in Section 2.2.4.2. The inter-assay coefficients of variation for three assays were 9.1 ., 6 .1, and 5.7% for high, medium and low quality controls respectively. 3.2.5 Calcu lation of the area under the corticosterone curve The area under the curve for plasma corticosterone concentrations versus time was calculated for each bird that was repeat sampled, using the method described in Chapter 2. 3.2.6 Statistical analyses All results are presented as the mean ± SE (standard error) or individual data where appropriate. All the data were examined for normality using Shapiro-Wilks tests and for equality of variance using Levene's homogeneity of variance test. Hormone data were IOglO transformed to avoid heteroscedasticity. If variances were homogenous then groups were compared with repeated measures two-way analyses of variance (ANOV A) with time and treatment as the grouping factors. Comparisons between times within each treatment and between treatments for each time were examined with post-hoc repeated measures contrasts. Data not complying with normality or variance criteria were compared 88 Chapter Three using a Kruskal-Wallis non-parametric test followed by Mann-Whitney U independent group comparisons tests, or Friedman's test for repeat measures data. All comparisons of means were conducting using SPSS™ 1 1 .0 (SPSS Inc., Illinois) or Systat™ 8.0 (SPSS Inc., Illinois). Graphs were generated using GraphPad Prism 4.0 (GraphPad Software Inc.) and SigmaPlot 2001 (SPSS Inc. , Il linois). 3.3 Resu lts 3 .3 . 1 Experiment 1 - Quantificat ion of the effect of manual restraint on plasma testosterone and corticosterone in the Japanese qua i l 3.3.1 . 1 Corticosterone Corticosterone varied between treatments and between times, with a significant interaction between treatment and time (see Table 3 .1 for al l statistics for this analysis). Initial corticosterone concentrations were low in all four groups of birds, with differences between groups (F3,136 = 5.752, P = 0.001; Figures 3. 1a) . Corticosterone remained low in untreated birds manually restrained for the minimum 2 min (Figures 3.1a) . Plasma corticosterone in birds manually restrained for 5 min increased after 15 min, then decreased to similar concentrations to untreated birds by 30 min and thereafter. Corticosterone rose to a peak at 15 min in birds manually restrained for 10 min (6.84 - 18.13 nglml) or 15 min (9.23 - 17.78 nglml) and did not differ significantly from each other. Corticosterone decreased between 15 and 30 min in birds manually restrained for 10 min, and did not differ from initial concentrations after 30 min and was similar to untreated birds thereafter. Plasma corticosterone decreased between 15 and 30 min in birds manually restrained for 1 5 min and was still elevated at 30 min. Corticosterone continued to decrease and was similar to initial concentrations at 60 min, although it subsequently increased between 60 and 120 min before remaining constant. Overall, plasma corticosterone concentrations at 15 min appeared to be related to the duration of manual restraint. Chapter Three 89 3.3.1 .2 Testosterone Testosterone did not vary between treatments and there was no interaction between treatment and time, although there was an effect of time (see Table 3.2 for all statistics for this analysis) . Testosterone concentrations were high and did not differ between the groups at the beginning of the experiment (F3,136 = 0.521, P = 0.668; Figure 3 .1b). Testosterone remained relatively constant in untreated birds. Plasma testosterone concentrations in birds manually restrained for 5 min decreased between 30 and 60 min, then returned to initial concentrations by 120 min. Testosterone remained unchanged between 0 and 30 min in birds manually restrained for 1 0 min then decreased after 30 min, was significantly lower than initial concentrations after 60 min then increased between 60 and 120 min and remained relatively constant. Testosterone initially (0-30 min) remained unchanged in birds manually restrained for 15 min, then dramatically decreased between 30 and 60 min and were significantly less than initial concentrations. Testosterone then increased back to pretreatment concentrations between 60 and 120 min and did not differ from untreated birds or initial testostosterone concentrations. Overall, the magnitude of the decrease of plasma testosterone concentrations by 60 min appeared to be related to the duration of manual restraint. Mean plasma testosterone concentrations were significantly lower at 60 min in birds manually restrained for 15 min than in any other birds. Testosterone concentrations did not differ between treatments at any other time. 90 ...- E - 0) c --- a> c 0 L- a> ....... en 0 Cl ....... L- 0 () E - 0) c --- a> c o L- a> ....... en o ....... en a> r- Chapter Three 1 5 (a) - ' 2 min 5 m in 1 0 - -0- - ' 1 0 min - 1 5 m i n . , 5 • ...,........ • . , - :., -' - - - - - .". -::-::"� - - - - - ..,, - _ .. .... -' 0 4 (b) 3 - - - - - - - - - - - - - - - - - - - - - - - - - �- - - . - - - - - - - - -- -- 2 1 � Manual restraint begins O ����--�--------�------------------�-- o 1 5 30 60 1 20 Time (min) 240 Figure 3.1 Plasma corticosterone (a) and testosterone (b) in male quail blood sampled at 0 min (2 min group) then manually restrained up to 5, 1 0 or 15 min after the initial blood sample, then blood sampled at 15, 30, 60, 1 20 or 240 min. Resul ts are plotted as mean ± standard error. Chapter Three 91 Table 3.1 Two-way repeated measures ANOVA of corticosterone concentrations in male quail that were manually restrained for 2, 5, 10 or 15 minutes. Effect Treahnent Time Interaction of treatment and time Comparisons within treatments for each treatment o vs 15 min 15 vs 30 min 30 vs 60 min 60 vs 120 min 120 vs 240 min o vs 30 min o vs 60 min o vs 120 min o vs 240 min 2 min manual restraint 5 min manual restraint 10 min manual restraint 15 min manual restraint 2 min manual restraint 5 min manual restraint ID min manual restraint 15 min manual restraint 2 min manual restraint 5 min manual restraint 10 min manual restraint 15 min manual restraint 2 min manual restraint 5 min manual restraint 10 min manual restraint 15 min manual restraint 2 min manual restraint 5 min manual restraint 10 min manual restraint 15 min manual restraint 2 min manual restraint 5 min manual restraint 10 min manual restraint 15 min manual restraint 2 min manual restraint 5 min manual restraint ID min manual restraint 15 min manual restraint 2 nun manual restraint 5 min manual restraint 10 min manual restraint 15 min manual restraint 2 nUn manua l restraint 5 min manual restraint 10 min manual restraint 15 min manual restraint Plasma corticosterone F 1 8.298 28.243 6.282 0.064 9.637 51 .227 92.188 7.101 4.531 36.845 8.553 0.172 3.247 4.926 26.883 2.283 0.493 4.709 9.993 0.442 0.433 0.034 2.342 1 0.164 0.127 0.461 33.941 7.035 3.880 12.563 0.753 0.492 1 .131 0.522 1 0.325 0.024 0.046 0.255 1 .531 df 3,256 5,256 1 5,256 1 ,256 1 ,256 1 ,256 1 ,256 1 ,256 1,256 1 ,256 1,256 1,256 1 ,256 1 ,256 1 ,256 1 ,256 1 ,256 1,256 1 ,256 1,256 1,256 1 ,256 1,256 1 ,256 1,256 1,256 1,256 1 ,256 1 ,256 1 ,256 1 ,256 1 ,256 1 ,256 1,256 1 ,256 1 ,256 1 ,256 1 ,256 1 ,256 p < 0.001 .,. < 0.001 .,. < 0.001 .,. 0.801 0.002 * * < 0.001 * * < 0.001 * * 0.008 * * 0.034 • < 0.001 • • 0.004 • • 0.679 0.073 0.027 • < 0.001 • • 0.132 0.483 0.031 • 0.002 • • 0.507 0.511 0.854 0.127 0.002 • • 0.722 0.498 < 0.001 • • 0.008 • • 0.051 < 0.001 • • 0.386 0.483 0.289 0.458 0.001 • • 0.876 0.831 0.614 0.217 Note. The first three rows show the results of the two-way ANOVA for the main effects and their interaction. The remaining rows show the results of post-hoc comparisons of times for each treatment and comparisons of treatments at each time. Significant differences (p < 0.05) are marked with a single asterisk, while highly significant differences (p < 0.01) are marked with a double asterisk. 92 Chapter Three Table 3.1 cont Two-way repeated measures ANOVA of corticosterone concentrations in male quail that were manually restrained for 2, 5, 1 0 or 15 minutes. Comparisons between trea tments for each time O min 15 min 30 min 60 min 120 mm 240 min 2 min vs 5 min 2 min vs 10 min 2 min vs 15 min 5 vs l0 min 5 vs 15 min 10 vs 15 min 2 min vs 5 min 2 min vs 10 min 2 min vs 15 min 5 vs 10 min 5 vs 15 min 10 vs 15 min 2 min vs 5 min 2 min vs 10 min 2 min vs 15 min 5 vs 10 min 5 vs 15 min 10 vs 15 min 2 mm vs 5 min 2 nun vs 10 mm 2 min vs 15 min 5 vs 10 min 5 vs 15 min 10 vs 15 min 2 mm vs 5 min 2 min vs 10 mm 2 min vs 15 min 5 vs 10 min 5 vs 15 min 10 vs 15 min 2 min vs 5 min 2 min vs 10 min 2 mm vs 15 min 5 vs 10 min 5 vs 15 min 10 vs 15 min Plasma corticosterone F 10.672 0.317 7.356 7.309 0.308 4.617 0.560 25.978 36.350 18.909 27.886 0.869 1 .650 2.861 33.285 0.166 20.1 13 16.628 0.869 0.889 0.029 0.000 1 .214 1 .237 3.031 0.081 3.31 1 2.123 12.679 4.425 3.054 0.584 0.141 0.967 1 .883 0.151 df 1 ,256 1 ,256 1 ,256 1 ,256 1,256 1,256 1 ,256 1 ,256 1 ,256 1,256 1 ,256 1 ,256 1 ,256 1 ,256 1 ,256 1 ,256 1 ,256 1 ,256 1 ,256 1 ,256 1 ,256 1,256 1 ,256 1 ,256 1,256 1 ,256 1 ,256 1 ,256 1,256 1 ,256 1 ,256 1 ,256 1 ,256 1 ,256 1 ,256 1 ,256 p 0.001 * * 0.574 0.007 * * 0.007 * * 0.580 0.033 • 0.455 < 0.001 • • < 0.001 • • < 0.001 • • < 0.001 • • 0.352 0.200 0.092 < 0.001 • • 0.684 < 0.001 • • < 0.001 • • 0.352 0.347 0.866 0.992 0.272 0.267 0.083 0.777 0.070 0.146 < 0.001 • • 0.036 • 0.082 0.445 0.708 0.326 0.171 0.698 Chapter Three 93 Table 3.2 Two-way repeated measures ANOVA of testosterone concentrations in male quail that were manually restrained for 2, 5, 10 or 15 minutes. Plasma testosterone Effect F df p Treatment 0.396 3,256 0.756 Time 2.653 5,256 0.023 .. Interaction of treatment and time 0.841 1 5,256 0.631 Com.earisons within treatments for each treatment o vs 15 min 2 min manual restraint 0.166 1 ,256 0.684 5 min manual restraint 0.022 1 ,256 0.882 10 min manual restraint 0.135 1 ,256 0.714 15 min manual restraint 0.007 1 ,256 0.935 15 vs 30 min 2 min manual restraint 0.356 1 ,256 0.551 5 min manual restraint 0.000 1 ,256 0.997 10 min manual restraint 1 .016 1 ,256 0.314 15 min manual restraint 0.057 1 ,256 0.811 30 vs 60 min 2 min manual restraint 0.004 1 ,256 0.950 5 min manual restraint 1 .985 1 ,256 0.160 10 min manual restraint 1 .135 1 ,256 0.288 15 min manual restraint 6.750 1 ,256 0.010 .. 60 vs 1 20 min 2 min manual restraint 0.001 1 ,256 0.973 5 min manual restraint 0.155 1 ,256 0.694 10 min manual restraint 4.109 1 ,256 0.044 .. 15 min manual restraint 4.346 1 ,256 0.038 .. 120 vs 240 min 2 min manual restraint 0.076 1 ,256 0.782 5 min manual restraint 0.151 1 ,256 0.698 10 min manual restraint 0.086 1 ,256 0.770 1 5 min manual restraint 0.485 1 ,256 0.487 o vs 30 min 2 min manual restraint 0.132 1,256 0.717 5 min manual restraint 0.021 1 ,256 0.886 10 min manual restraint 0.873 1 ,256 0.351 15 min manual restraint 0.052 1,256 0.820 o vs 60 min 2 min manual restraint 0.197 1 ,256 0.657 5 min manual restraint 2.806 1 ,256 0.095 10 min manual restraint 5.333 1 ,256 0.022 .. 1 5 min manual restraint 12.829 1 ,256 < 0.001 .. * o vs 120 min 2 min manual restraint 0.238 1 ,256 0.626 5 min manual restraint 1 .362 1 ,256 0.244 10 min manual restraint 0.095 1 ,256 0.759 J 5 min manual restraint 0.793 1 ,256 0.374 o vs 240 min 2 min manual restraint 0.017 1 ,256 0.896 5 min manual restraint 0.443 1 ,256 0.506 10 min manual restraint 0.005 1 ,256 0.944 15 min manual restraint 0.000 1 ,256 0.993 Note. The first three rows show the results of the two-way ANOVA for the main effects and their interaction. The remaining rows show the results of post-hoc comparisons of times for each treatment and comparisons of treatments at each time. Significant differences (p < 0.05) are marked with a single asterisk, while highly significant differences (p < 0.01 ) are marked with a double asterisk. 94 Chapter Three Table 3.2 cont Two-way repeated measures ANOVA of testosterone concentrations in male quail that were manually restrained for 2, 5, 10 or 15 minutes. Plasma testosterone Comparisons between treatments for each time F df p O min 2 min vs 5 mjn 1 .319 1 ,256 0.252 2 mjn vs 10 mjn 1 .408 1 ,256 0.236 2 min vs 15 mm 0.882 1 ,256 0.365 5 vs 10 nun 0.001 1 ,256 0.969 5 vs 15 min 0.058 1 ,256 0.809 10 vs 15 min 0.078 1 ,256 0.780 15 min 2 min vs 5 mm 0.891 1 ,256 0.346 2 min vs 10 min 1 .278 1 ,256 0 .259 2 min vs 15 min 0.615 1 ,256 0.434 5 vs 10 min 0.035 1 ,256 0.852 5 vs 15 min 0.026 1 ,256 0.873 10 vs 15 nun 0.120 1 ,256 0.729 30 min 2 min vs 5 min 0.1 1 8 1 ,256 0.732 2 mjn vs 10 min 0.225 1 ,256 0.636 2 min vs 15 mjn 0.003 1 ,256 0.958 5 vs l0 min 0.669 1 ,256 0.414 5 vs 1 5 min 0.156 1 ,256 0.693 10 vs 15 min 0.178 1 ,256 0.673 60 min 2 nun vs 5 min 1 .273 1 ,256 0.260 2 min vs 10 min 2.567 1 ,256 0.1 10 2 min vs 15 mjn 7.361 1 ,256 0.007 • • 5 vs 10 mjn 0.225 1 ,256 0.636 5 vs 1 5 min 2.512 1 ,256 0 .114 1 0 vs 1 5 mjn 0.159 1 ,256 0.686 120 mm 2 min vs 5 nUn 0.590 1 ,256 0.443 2 mm vs 10 min 0.153 1 ,256 0.696 2 mm vs 15 nUn 0.438 1 ,256 0.509 5 vs 1 0 nUn 1 .344 1 ,256 0.247 5 vs 1 5 min 0.01 1 1 ,256 0.915 10 vs 15 min 1 . 109 1 ,256 0.293 240 mjn 2 min vs 5 min 0.01 1 1 ,256 0.918 2 mjn vs 10 mjn 0.140 1 ,256 0.708 2 mjn vs 15 mjn 0.097 1 ,256 0.756 5 vs 10 mm 0.228 1 ,256 0.633 5 vs 15 min 0.171 1 ,256 0.679 10 vs 15 mjn 0.004 1 ,256 0.949 Chapter Three 3.3 .2 Experiment 2 - Corticosterone and testosterone responses to manual restraint, mechan ical restraint and social stimu l i in Japanese quai l 95 3.3.2 . 1 The effect of different stimul i on the corticosterone response Corticosterone did not change significantly after the first sample in untreated birds (Figure 3.2). In birds that were exposed to an unfamiliar quail that was kept physically separated, corticosterone increased at 15 or 30 min in some birds while there was no response in others. The corticosterone concentrations of birds that were kept in the same cage as an unfamiliar quail did not differ from untreated birds, with one exception. Corticosterone increased at 15 min following 15 min of mechanical restraint, then decreased between 15 and 60 min and remained relatively thereafter. The magnitude of the increase in corticosterone varied between birds and concentrations ranged from 1 .24 to 6.06 nglml at 15 min. There was a strong corticosterone response in all birds manually restrained for 15 min with a peak in corticosterone at 15 min followed by a decrease to initial concentrations at 30 min. Initial corticosterone concentrations differed between groups (F4,45 = 2 .671, P = 0.044; Figure 3.3a). The changes in corticosterone from initial concentrations were therefore calculated for all birds (Figure 3.3b), with a value of 5 nglml added to each calculated value to enable them to be log1o transformed for statistical analysis. Overall, mean changes in corticosterone did not vary between treatment but did between time, and there was a significant interaction between treatment and time (see Table 3.3 for all statistics for this analysis). Corticosterone concentrations remained low and relatively constant in birds that were untreated and those in contact with an unfamiliar quail. Corticosterone was elevated at 15 min in birds that were exposed to an unfamiliar quail but kept separate from them, then decreased and did not differ from untreated birds. Mean corticosterone concentrations in mechanically restrained birds increased 15 min after the initial sample and were higher than in birds kept separated from an unfamiliar quail. Corticosterone in the birds manually restrained for 15 min 96 Chapter Three increased to a peak at 15 min (6. 1 1 ± 1 .93 nglml), declined to low concentrations at 30 min and remained below initial concentrations at 60 and 120 min. Concentrations at 30, 60 and 120 min generally did not differ greatly between treatments. There was a significant difference in the total integrated corticosterone response between the groups (F4,45 = 7.305, P < 0.001; Figure 3.4a), although differences in baseline corticosterone between the groups confounded differences in the integrated response. Corrected corticosterone responses were calculated from 0 to 30 min. Corticosterone concentrations between 0 and 30 min varied between groups (Kruskal-Wallis t = 21 .376, P < 0.001; Figure 3.4b). Untreated birds and those in direction contact with an unfamiliar bird showed little response over this time, whilst there was a significant response in birds that were mechanically restrained and those that were exposed to an unfamiliar bird but kept separated from it. These two responses did not differ (post-hoc Bonferroni test, p = 0.356). Birds that were manually restrained for 15 min showed the largest corrected corticosterone response between 0 and 30 min, and this was significantly greater than responses to either mechanical restraint or visual exposure to an unfamiliar quail (post-hoc Bonferroni test, p = 0.043 and p = 0.022 respectively) . Chapter Three --- 2 min 8 6 4 • .0 .' .' 2j I m������0iii� 8� I °i , E - 0) 8 Unfamiliar quail (separated) 8 Unfamiliar quail (contact) C --- Q) c e Q) +-' Cl) o u t o <-) "'� :: ����:�i:;��i� ./ " o IL-,-_-._-. ___ -. _______ ---._ 6 4 2 ° LI� __ '-__ -r ______ � ________________ '-__ Manual restraint 8 Mechanical restraint 8 6 6 4 4 2 ! - ·-'·--:_.L �?;;:�� 2 . . o IL.,----r--.----.-------...,- °LI�-�-�--��------�- o 15 30 60 120 o 15 30 60 120 Time (min) 97 Figure 3.2 Plasma corticosterone in male quail that were blood sampled (2 min), or in addition were manually or mechanically restrained for 15 min, or exposed to an unfamiliar quail that was visible but separated by a wire mesh or in direct physical contact. Results are plotted as raw data. 98 Chapter Three - -0- - ' 2 min (a) 6 -----d---- Unfami l iar quai l (separated) ......... --- Unfami l iar quai l (contact) E - Mechanical restraint -0) c - Manual restraint .......... (l) 4 c a L- ........... (l) -'-...... .. ...... Cl) a ..... () --- .......... _- .. _-...... 2 ---------- -_ ... _---------L- a U + 0 6 0 1 5 30 60 1 20 ......... (b) E 5 -0) c .......... 4 (l) c 0 3 L- (l) ...... Cl) 0 2 () ...... L- a 1 () c (l) 0 0) - - - - -c ro - 1 .c U -2 0 1 5 30 60 1 20 Time (min) Figure 3.3 Plasma corticosterone (a) and change in plasma corticosterone relative to 0 min (b) in male quail that were blood sampled (2 min), or in addition were manually or mechanically restrained for 15 min, or exposed to an unfamiliar quail that was visible but separated by a wire mesh or in direct physical contact. Results are plotted as mean ± standard error. Chapter Three ...--.. c E E -0> C ......... ID IF) C 0 c-IF) � 0 ID c 0 1 00 � ID ...... IF) 0 (J 80 t 0 (J "'0 60 ID ...... co � 0> ID 40 ...... c 20 0 Total response (a) Corrected response (0 - 30 min) ( b ) c:=J 2 min CJ Unfamil iar quai l (separated) c:=J Unfamil iar quai l (contact) c:=J Mechanical restra int _ Manual restra int 99 Figure 3.4 Total (a; all groups n=10) and corrected (b) integrated corticosterone responses (0-30 min) in male quail that were blood sampled (2 min, n=8), or in addition were mechanically restrained (n=10) or manually restrained for 15 min (n= 1 0), or exposed to an unfamiliar quail that was visible but separated by a wire mesh (n =9) or in direct physical contact (n= 6) . Results are plotted as mean ± standard error. 100 Cha pter Three Table 3.3 Two-way repeated measures ANOVA of corticosterone concentrations in male quail exposed to different stimuli. Plasma corticosterone Effect F df p Treatment 1 .081 4,45 0.377 Time 54.050 3,135 < 0.001 • • Interaction of treatment and time 12.632 12,135 < 0.001 • • Comparisons within treatments for each treatment 15 vs 30 min 2 min 0.295 1,45 0.590 Separated 0.989 1,45 0.325 Contact 4.190 1,45 0.047 • Mechanical restraint 17.573 1,45 < 0.001 • • Manual restraint 128.820 1,45 < 0.001 • • 30 v s 60 min 2 min 7.539 1 ,45 0.009 • • Separated 12.320 1,45 0.001 • • Contact 4.956 1,45 0.031 • Mechanical restraint 4.613 1,45 0.037 • Manual restraint 3.040 1,45 0.088 60 vs 120 min 2 min 3.677 1,45 0.062 Separated 0.328 1,45 0.570 Contact 0.127 1,45 0.723 Mechanical restraint 0.105 1,45 0.748 Manual restraint 0.262 1,45 0.611 Note. The first three rows show the results of the two-way repeated measures ANOVA for the main effects and their interaction. The remaining rows show the results of post-hoc comparisons of times for each treatment and comparisons of treatments at each time. Significant differences (p < 0.05) are marked with a single asterisk, while highly significant differences (p < 0.01 ) are marked with a double asterisk. Cha pter Three 101 Table 3.3 cont Two-way repeated measures ANOV A of corticosterone concentrations in male quail exposed to different stimuli. Plasma corticosterone Comparisons between treatments for each time 15 min 30 min 60 min 120 min 2 min vs Separated 2 min vs Contact 2 min vs Mechanical restraint 2 min vs Manual restraint Separated vs Contact Separated vs Mechanical restraint Separated vs Manual restraint Contact vs Mechanical restraint Contact vs Manual restraint Mechanical vs Manual restraint 2 min vs Separated 2 min vs Contact 2 min vs Mechanical restraint 2 min vs Manual restraint Separated vs Contact Separated vs Mechanical restraint Separated vs Manual restraint Contact vs Mechanical restraint Contact vs Manual restraint Mechanical vs Manual restraint 2 min vs Separated 2 min vs Contact 2 min vs Mechanical restraint 2 min vs Manual restraint Separated vs Contact Separated vs Mechanical restraint Separa ted vs Manual restraint Contact vs Mechanical restraint Contact vs Manual restraint Mechanical vs Manual restraint 2 min vs Separated 2 min vs Contact 2 min vs Mechanical restraint 2 min vs Manual restraint Separated vs Contact Separated vs Mechanical restraint Separated vs Manual restraint Contact vs Mechanical restraint Contact vs Manual restraint Mechanical vs Manual restraint F 6.177 1 .064 21 .783 63.456 12.368 4.760 30.036 32.474 80.951 10.881 0.826 0.034 0.260 2.557 0.526 0.159 6.291 0.106 3.179 4.448 0.1 89 0.276 0.829 0.973 0.008 0.227 2.019 0 .148 2.286 3.599 1 .646 1 .082 0.387 7.279 0.059 0.437 2.002 0.175 2.749 4.309 df 1 ,45 1 ,45 1,45 1 ,45 1 ,45 1 ,45 1 ,45 1,45 1 ,45 1,45 1,45 1,45 1 ,45 1 ,45 1,45 1 ,45 1,45 1 ,45 1 ,45 1 ,45 1 ,45 1 ,45 1 ,45 1,45 1 ,45 1 ,45 1 ,45 1 ,45 1 ,45 1,45 1 ,45 1 ,45 1,45 1 ,45 1 ,45 1 ,45 1 ,45 1 ,45 1 ,45 1,45 p 0.017 • 0.308 < 0.001 • • < 0.001 • • 0.001 • • 0.034 • < 0.001 • • < 0.001 * * < 0.001 • • 0.002 • • 0.368 0.855 0.613 0.1 17 0.472 0.692 0.016 • 0.746 0.081 0.041 • 0.666 0.602 0.367 0.329 0.928 0.636 0.162 0.702 0.137 0.064 0.206 0.304 0.537 0.010 • 0.809 0.512 0.164 0.678 0.104 0.044 .. 102 Chapter Three 3.3.2.2 The effect of mechanical and manual restra int on corticosterone and testosterone concentrations 3.3 .2 .2 . 1 Plasma corticosterone Corticosterone varied between treatment and between time, with a significant interaction between treatment and time (see Table 3.4 for all statistics for this analysis). Half of each treatment group (n = 10) were sampled on one day and the other half 10 days later. Corticosterone did not differ between the different sample days (see Table 3.6 for statistics for this analysis), thus the birds were combined for subsequent analyses. Initial corticosterone concentrations were low « 4 ng/ml) and remained constant in untreated birds (Figure 3.5a), and initial corticosterone concentrations did not differ between the three groups (F2,132 = 0.631, P = 0.533) . Mean corticosterone concentrations increased to a peak (5.34 ± 1 . 75 ng/ml) at 15 min in birds that were mechanically restrained, declined from this peak until 60 min and then remained constant. Concentrations at 60, 120 and 240 min did not differ from initial corticosterone concentrations or those of untreated birds. Manual restraint for 15 min elicited a larger increase in plasma corticosterone than mechanical restraint after 1 5 min (7.95 ± 2.51 ng/ml), with corticosterone then declining by 30 min, and were lower than initial values at 60 min and remaining relatively constant thereafter. Corticosterone at 60, 120 and 240 min did not differ from initial concen tra tions. 3.3 .2 .2 .2 Plasma testosterone Testosterone varied between treatment and between time, with a significant interaction between treatment and time (see Table 3.5 for all statistics for this analysis), while testosterone did not differ between the two sample days (Table 3.6). Initial testosterone concentrations did not differ between the groups (F2,132 = 0.218, P = 0.804; Figure 3.5b), and did not change with time in untreated birds. Testosterone declined between 0 and 30 min in birds that were mechanically restrained, was significantly lower than in untreated birds after 60 min and remained low until 120 min, before increasing to a concentration similar to Cha pter Three 103 untreated birds at 240 min. 15 min manual restraint caused testosterone concentrations to decline until 60 min and remain low until 120 min before increasing between 120 and 240 min. The mean concentration at 240 min did not differ from the initial concentrations or from that of untreated birds at 240 min. Mean testosterone concentrations in mechanically restrained birds were most different from untreated birds at 60 min with manual restraint producing the most effect. 104 1 0 ............. E 8 - Cl C --- ID 6 c 0 L- ID 4 +-' Cl) 0 () +-' 2 L-0 0 0 6 ............. 5 E - Cl 4 c --- ID C 3 0 L- ID +-' 2 Cl) 0 +-' Cl) ID 1 I- 0 Chapter Three . - - - - - - - � -- - -� - - - -o- - - - - - - - - - - - - - � � Restraint begins o 1 5 30 60 1 20 Time (min) 2 min manual restra int - -0- - ' Mechanical restra int - Manual restraint 240 (a) (b) Figure 3.5 Plasma corticosterone (a) and testosterone (b) male quail blood sampled at 0 min (2 min manual restraint) then manually or mechanically restrained for 1 5 min, then blood sampled at 15, 30, 60, 120 o r 240 min. Results are plotted a s mean ± standard error. Chapter Three 105 Table 3.4 Two-way repeated measures ANOVA of corticosterone concentrations in male quail manually or mechanically restrained for 15 min. Plasma corticosterone Effect F df p Treatment 9.929 2,252 < 0.001 • • Time 17.434 5,252 < 0.001 •• Interaction of treatment and time 3.861 1 0,252 < 0.001 • • Comparisons within treatments for each treatment o vs 15 min 2 min 0.234 1 ,252 0.629 Mechanical restraint 34.848 1 ,252 < 0.001 • • Manual restraint 69.095 1 ,252 < 0.001 • • 15 v s 30 min 2 min 0.107 1 ,252 0.744 Mechanical restraint 12.433 1 ,252 0.001 . .. Manual restraint 1 1 .913 1 ,252 0.001 . .. 30 v s 60 min 2 min 0.402 1 ,252 0.527 Mechanical restraint 5.034 1 ,252 0.026 .. Manual restraint 20.381 1 ,252 < 0.001 . .. 60 vs 1 20 min 2 min 0.1 15 1 ,252 0.735 Mechanical restraint 0.037 1 ,252 0.847 Manual restraint 3.278 1 ,252 0.071 120 vs 240 min 2 min 1 .517 1 ,252 0.219 Mechanical restraint 0.844 1 ,252 0.359 Manual restraint 0 .265 1 ,252 0.607 o vs 30 min 2 min 0.004 1 ,252 0.950 Mechanical restraint 1 .825 1 ,252 0.178 Manual restraint 14.872 ] ,252 < 0.001 • • o vs 60 min 2 min 0.571 1 ,252 0.450 Mechanical restraint 2.389 1 ,252 0.123 Manual restraint 1 .825 1 ,252 0.178 o vs 1 20 min 2 min 0.101 1 ,252 0.751 Mechanical restraint 3.222 ],252 0.074 Manual restraint 2.389 1 ,252 0.123 o vs 240 min 2 min 1 .618 1 ,252 0.205 Mechanical restraint 0.371 1 ,252 0.543 Manual restraint 3 .222 1 ,252 0.074 Note. The first three rows show the results of the two-way ANOVA for the main effects and their interaction. The remaining rows show the results of post-hoc comparisons of times for each treatment and comparisons of treatments at each time. Significant differences (p < 0.05) are marked with a single asterisk, while highly significant differences (p < 0.01 ) are marked with a double asterisk. 106 Chapter Three Table 3.4 cont Two-way repeated measures ANOV A of corticosterone concentrations in male quail manually or mechanically restrained for 15 min. Plasma corticosterone Comparisons between treatments for each time F df p O min 2 min vs Mechanical restraint 0.196 1 ,252 0.658 2 min vs Manual restraint 1 .240 1 ,252 0.267 Mechanical vs Manual restraint 0.546 1,252 0.461 15 min 2 min vs Mechanical restraint 15 .333 1 ,252 < 0.001 >1->1- 2 min vs Manual restraint 35.218 1 ,252 < 0.001 >1- >1- Mechanical vs Manual restraint 4.825 1 ,252 0.029 >I- 30 min 2 rnin vs Mechanical restraint 1 .200 1 ,252 0.274 2 min vs Manual restraint 9.962 1,252 0.002 >1- >1- Mechanical vs Manual restraint 5.128 1 ,252 0.024 >I- 60 min 2 min vs Mechanical restraint 0.064 1 ,252 0.801 2 min vs Manual restraint 0.000 1,252 1 .000 Mechanical vs Manual restraint 0.000 1 ,252 1 .000 120 min 2 min vs Mechanical restraint 0.635 1,252 0.426 2 min vs Manual restraint 1 .044 1,252 0.308 Mechanical vs Manual restraint 4.016 1,252 0.046 >I- 240 min 2 min vs Mechanical restraint 1 .687 1 ,252 0.195 2 min vs Manual restraint 0.610 1 ,252 0.436 Mechanical vs Manual restraint 0.326 1 ,252 0.569 Chapter Three 107 Table 3.5 Two-way repeated measures ANOV A of testosterone concentrations in male quail manually or mechanically restrained for 15 min. Plasma testosterone Effect F df p Treatment 14.219 2,252 < 0.001 •• Time 1 1 .071 5,252 < 0.001 • • Interaction o f treatment and time 4.207 10,252 < 0.001 •• Comparisons within treatments for each treatment o vs 15 min 2 min 0.011 1 ,252 0.917 Mechanical restraint 3.899 1 ,252 0.049 • Manual restraint 4.304 1 ,252 0.039 • 15 vs 30 min 2 min 0.000 1 ,252 0.991 Mechanical restraint 0.149 1 ,252 0.700 Manual restraint 0.416 1 ,252 0.519 30 vs 60 min 2 min 0.006 1 ,252 0.940 Mechanical restraint 0.000 1 ,252 0.984 Manual restraint 17.837 1 ,252 < 0.001 • • 60 vs 120 mm 2 min 0.161 1,252 0.689 Mechanical restraint 1 . 152 1 ,252 0.284 Manual restraint 2.825 1 ,252 0.094 120 vs 240 min 2 min 0.136 1 ,252 0.713 Mechanical restraint 1 1 .723 1 ,252 0.001 . . Manual restraint 18.980 1 ,252 < 0.001 • • o vs 30 min 2 min 0.014 1 ,252 0.905 Mechanical restraint 6 .114 1 ,252 0.014 • Manual restraint 6.318 1 ,252 0.013 • o vs 60 min 2 min 0.000 1 ,252 0 .982 Mechanical restraint 6.240 1 ,252 0.013 • Manual restraint 63.457 1 ,252 < 0.001 . . o vs 1 20 min 2 min 0.292 1 ,252 0.590 Mechanical restraint 1 5.080 1 ,252 < 0.001 -- Manual restraint 27.959 1 ,252 < 0.001 • • o vs 240 min 2 min 1 .031 1 ,252 0.311 Mechanical restraint 0.288 1,252 0.592 Manual restraint 0 .113 1 ,252 0.737 Note. The first three rows show the results of the two-way ANOVA for the main effects and their interaction. The remaining rows show the results of post-hoc comparisons of times for each treatment and comparisons of treatments a t each time. Significant differences (p < 0.05) are marked with a single asterisk, while highly significant differences (p < 0.01 ) are marked with a double asterisk. 108 Chapter Three Table 3.5 cont Two-way repeated measures ANOV A of testosterone concentrations in male quail manually or mechanically restrained for 15 min. Plasma testosterone Comparisons between treatments for each time F df p O min 2 min vs Mechanical restraint 0.005 1 ,252 0.941 2 min vs Manual restraint 0.320 1,252 0.572 Mechanical vs Manual restraint 0.295 1 ,252 0.588 15 min 2 min vs Mechanical restraint 2.081 1,252 0.150 2 min vs Manual restraint 1 .032 1 ,252 0.311 Mechanical vs Manual restraint 0.221 1 ,252 0.639 30 min 2 min vs Mechanical restraint 3.259 1 ,252 0.072 2 min vs Manual restraint 2.605 1 ,252 0.108 Mechanical vs Manual restraint 0.044 1 ,252 0.833 60 min 2 min vs Mechanical restraint 3.033 1 ,252 0.083 2 min vs Manual restraint 28.784 1,252 < 0.001 * * Mechanical vs Manual restraint 1 5.943 1 ,252 < 0.001 * * 120 min 2 min vs Mechanical restraint 9.926 1 ,252 0.002 * * 2 min vs Manual restraint 15.346 1 ,252 < 0.001 ** Mechanical vs Manual restraint 0.714 1 ,252 0.399 240 min 2 min vs Mechanical restraint 0.196 1 ,252 0.658 2 min vs Manual restraint 0.132 1 ,252 0.717 Mechanical vs Manual restraint 0.008 1 ,252 0.930 Table 3.6 One-way ANOVA corticosterone and testosterone concentrations in quail sampled on Day 0 versus 10. Day 0 versus 1 0 Effect F df p Corticosterone Manual restraint 0.041 1 ,50 0.841 Mechanical restraint 0.008 1,50 0.927 Testosterone Manual restraint 2.526 1,50 0.120 Mechanical restraint 0.062 1,50 0.804 3.4 D iscussion Manual restraint o f male Japanese quail for 10 minutes o r greater resulted in a significant rise in plasma corticosterone concentrations, with corticosterone remaining elevated for up to 30 minutes. The increase in corticosterone was Cha pter Three 109 followed by a decline in testosterone. Testosterone was lowest after 60 minutes and remained low for up to two hours. The adrenocortical response varied with the type of the stressor. Manual restraint consistently initiated a large corticosterone increase in every bird, while the response to mechanical restraint varied from a small increase to a strong response seen in handled birds. The introduction of an unfamiliar quail as a social stimulus resulted in no change in corticosterone when quail were placed in the same cage as the unfamiliar quail despite both birds displaying agonistic behaviour towards each other. In contrast, unfamiliar quail separated by a wire mesh showed a significant adrenocortical response, although there was no clear pattern to corticosterone secretion. Mechanical restraint causing immobilisation resulted in a significant decline in plasma testosterone for up to 2 hours, manual restraint was more effective and caused a larger decrease in testosterone at 60 minutes. 3.4. 1 Quantification of the effect of manual restraint on plasma testosterone and corticosterone 3.4.1 . 1 Effect of manual restraint time on plasma corticosterone This experiment showed that 5 or more minutes o f immobilisation by manual restraint were necessary to significantly increase plasma corticosterone concentrations at 15 minutes in male Japanese quail . The response of quail to 15 min of manual restraint was similar to the response of handling reported previously in our laboratory (Boyd, 2000). All birds were blood sampled twice only, a t 0 minutes and one further time to prevent the additive effect of serial blood sampling on corticosterone and testosterone as reported in previous studies (Wilson et al., 1979; Eskeland and Blom, 1979; Johnson, 1981). All blood samples were collected within 2 min to ensure that measured corticosterone values did not reflect a response to the bleeding procedure. Corticosterone can increase after 2 minutes in quail used in our lab (Boyd, 2000) . Five minutes of manual restraint resulted in a small increase in corticosterone after 15 minutes. However, corticosterone may have increased during the 5 min period of restraint then decreased by the time of the 15 min sample. Plasma corticosterone 110 Cha pter Three rises rapidly in response to manual restraint, increasing within two minutes in chickens (Beuving and Yonder, 1978), and one minute in ducks (Harvey et al., 1980) and Japanese quail (Satterlee and Johnson, 1988) . Plasma corticosterone concentrations in Japanese quail restrained for 20 min peak after four minutes and were no greater at 10 or 20 min (Satterlee and Johnson, 1988) . Jones et al. (1994a) reported corticosterone increased significantly after 5.5 minutes of manual restraint, so the five minutes of manual restraint in the current study is likely to have caused a transient increase in corticosterone that was not detected after 15 minutes. The half-life of corticosterone in male Japanese quail is 10 minutes (Kovacs and Peczely, 1983) thus an increase could have occurred and diSSipated within 15 min. Therefore peak corticosterone concentrations after 5 min of manual restraint are likely to have been lower than after 10 and 15 minutes of manual restraint. Ten minutes of manual restraint resulted in a five-fold increase in mean corticosterone concentrations after 15 minutes, followed by a decline to initial concentrations by 30 minutes. Ten minutes of manual restraint clearly initiated a strong adrenocortical response in our strain of Japanese quail, whereas halving the manual restraint time had substantially less effect on the corticosterone response. Peak corticosterone concen trations at 15 minutes in birds manually restrained for 10 minutes appeared less than birds manually restrained for 15 minutes but results were not statistically significant. There was a seven-fold increase in plasma corticosterone in birds manually restrained for 15 minutes, with corticosterone still elevated at 30 minutes. Boyd (2000) reported a three-fold increase at 15 minutes after handling, and a return to initial concentrations by 30 minutes. The differences in peak corticosterone concentration between the current study and that reported by Boyd (2000) may indicate different responses to continuous immobilisation by hand (current study) compared with repeatedly removing and returning a bird from a box. The higher peak and prolonged elevation in corticosterone concentrations in the current study indicate that continuous manual restraint is a more potent stressor than picking up and putting down. Chapter Three 1 1 1 3.4. 1 .2 Effect of duration of manual restra int on plasma testosterone Manual restraint for 5 minutes or greater resulted in a significant decline in plasma testosterone after 60 minutes. Decreases in testosterone concentration following manual restraint and serial blood sampling have been found in quail (previous chapter) and chickens (Wilson et al., 1979; Eskeland and Blom, 1979; Johnson, 1 981). A different group of birds was blood sampled at each time to avoid measuring the decline in testosterone associated with serial blood collection from the same bird, This resulted in testosterone concentrations remaining unaffected for up to 30 minutes after the manual restraint in all groups. Collection of the first blood sample had no effect on subsequent testosterone concentrations in the untreated group. Mean testosterone in birds manually restrained for five minutes decreased, but not significantly, between 30 and 60 minutes. Manual restraint for 10 or 15 minutes resulted in plasma testosterone concentrations falling significantly below initial concentrations 60 minutes after the onset of manual restraint. The delay in the decline of testosterone secretion until 30 minutes after the quail were first handled conflicts with what was found in the previous chapter. In Chapter 2, testosterone had decreased 30 minutes after the collection of the blood sample and injection of oil or corticosterone in oil. Blood sampling followed by the added injection procedure is the most likely cause of the initial decline in testosterone secretion by 30 minutes, as testosterone had not significantly decreased at 30 minutes in birds that were only blood sampled. In the current study, the first manipulation was restricted to a short period of blood collection taking less than 2 minutes, followed by manual restraint until 5, 10 or 15 minutes after birds were first removed from the cage. Testosterone did not decline for the 15 minutes after birds were first returned to their cages. Heiblum et al. (2000) reported that testosterone increased in chickens 30 minutes after 10 minutes of immobilisation, returning to initial concentrations by 60 minutes. However, testosterone declined in response to mechanical restraint in one quarter of the chickens indicating individual variation in the testosterone response to mechanical restraint. 112 Chapter Three 3.4.2 Adrenocortica l and testosterone responses to d ifferent stressors 3.4.2 . 1 Adrenocortical responses to manual restraint, mechanical restra int and socia l stressors The results of the experiment described in Section 3.3.1 clearly demonstrated that 10 or 15 minutes of immobilisation by manual restraint resulted in a decrease in testosterone by 60 minutes. The current experiment showed that the adrenocortical response varies depending on the type of stressor. As the aim of the study was to examine the adrenocortical response to different stressors, repeated sampling of the same bird was appropriate in this study. Social disruption has previously been shown to initiate a corticosterone response in quail. Satterlee et al. (1983) reported that corticosterone was elevated at two hours in wild-type quail after the introduction of an albino quail. In the current study, corticosterone did not change when an unfamiliar male quail was introduced into the cage for 15 minutes. This was despite vigorous pecking and other agonistic behaviours being exhibited by both birds towards each other. Littin and Cockrem (2001) also reported no effect of mixing unfamiliar chickens. In contrast, the introduction of an unfamiliar quail that was physically separated by a wire partition resulted in a strong adrenocortical response after 1 5 minutes in 30% of quail. Both quail attempted pecking the other through the wire mesh during the 15 minutes, a behaviour that was infrequent between individuals that were normally housed next to each other. Manual restraint for 15 minutes resulted in a significant increase in corticosterone in all birds. The magnitude of the response was similar for all birds indicating that the stressor elicited a maximum adrenocortical response. Corticosterone concentrations had declined to initial concentrations by 30 minutes, as reported for female quail handled for 15 minutes (Boyd, 2000) . The adrenocortical responses varied between birds restrained and immobilised in a small metal cage for 15 minutes, with responses ranging from small increases in corticosterone to increases identical to those after 15 minutes of manual restraint. Mechanical restraint is a weaker stressor than manual restraint, so is preferable to manual restraint for investigating Chapter Three 1 13 differences between birds in their corticosterone responses. The response of quail to mechanical restraint in the current study is not unlike that reported by Satterlee and Johnson (1988), where mechanical restraint was used to discriminate between birds with low or high corticosterone responses to immobilisa tion. 3.4.3 Adrenocortical and testosterone response to manual restraint and mechanical restra int 3.4.3.1 Plasma corticosterone Initial corticosterone concentrations in quail in the previous two experiments and in this experiment were about 2 ng/ml, higher than the 1 ng/ml reported in quail by Satterlee and Johnson (1988), but similar to concentrations reported for chickens (Beuving and Yonder, 1978; Littin and Cockrem, 2001) . Fifteen minutes of manual restraint initiated a corticosterone response at 15 minutes, a result similar to that reported in Section 3.3 .1 , where corticosterone increased to a peak of 12.53 ± 4.74 ng/ml whilst it was 7.95 ± 2.51 ng/ml in the current study. The peaks in corticosterone are comparable with those reported previously for quail (4 to 18 ng/ml; Satterlee and Johnson, 1988) and chickens (4 to 1 6 ng/ml; Beuving and Yonder, 1978; Beuving and Yonder, 1986; Littin and Cockrem, 2001), whilst significantly lower than those reported in free-living, wild birds subjected to a similar stressor (Silverin, 1998) . In the current study, manual restraint initiated a strong response of similar magnitude in all birds. Previous studies showed a rapid decrease in corticosterone after the end of the stressor, returning to basal after 30 minutes in quail (Boyd, 2000), and 40 minutes in chickens (Littin and Cockrem, 2001) . Mechanical restraint initiated a significant corticosterone response, although as discussed in the previous experiment (Section 3.3.2), the response varied between birds and the mean response was less than that of manually restrained birds at 5.54 ± 1 .75 ng/ml at 15 minutes. This highlights the individual variation of the corticosterone response to mechanical restraint in the quail. Boyd (2000) also reported variation in the corticosterone response to handling after 15 minutes. Handling was a weaker stressor than manual restraint in the current experiment, whilst mechanical restraint in the current study initiated a varying adrenocortical 114 Chapter Three response resulting in varying responses as reported in the previous study and by Satterlee and Johnson (1988) and Boyd (2000). 3.4.3.2 Plasma testosterone Manual and mechanical restraint resulted in a significant decrease in plasma testosterone concentrations in both groups after 15 minutes. Testosterone remained low for up to 120 minutes in the mechanical restraint group while it continued to decline until 60 minutes in the manual restraint group. In the previous chapter, the decrease in testosterone was more rapid following serial blood sampling of the same bird, while Heiblum et al. (2000) reported testosterone increased 10 minutes after mechanical restraint in 75% of chickens. Testosterone declined more rapidly in this comparison of manual and mechanical restraint than that described in Section 3.4. 1 .2. In the current study, plasma testosterone concentrations started higher at 4.5 - 5 nglml compared with 2.7 - 3 nglml in the earlier study. I t is possible that the inhibitory effect of manual and mechanical restraint on testosterone is greater for birds with higher basal testosterone. Testosterone concentrations were their lowest at 1 - 1 .5 nglml, 60 minutes after birds were first taken from their cages and manually restrained for 1 5 minutes in both the current study and in Section 3.4.1 .2. 15 minutes of manual and mechanical restraint clearly initiated a decline in testosterone concentrations. Testosterone was lowest at 30 and 60 minutes in mechanical and manual restraint groups respectively, while it was unchanged in untreated birds. In contrast, the results in the previous chapter showed that serial blood sampling resulted in a decline in testosterone concentrations in untreated as well as treatment birds. It is clear that blood sampling separate groups of birds at each sampling time is necessary when the effect of a single stressor on plasma testosterone is being examined. Manual and mechanical restraint both initiated an increase in plasma corticosterone concentrations which was associated with a decline in plasma testosterone. However, it cannot be concluded from these results alone that elevated Chapter Three 1 15 corticosterone caused the decrease in testosterone secretion. A previous study showed that a single injection of corticosterone (1 .2 mglbird) causes a significant decline in plasma testosterone, still measurable after 3 hours (Boyd, 2000). Unlike the study in Chapter 3, the first blood sample after the injection was collected at 3 hours meaning repeated handling was unlikely to have caused the decline. The decline in testosterone in response to mechanical restraint was of the same duration as that in manually restrained birds, but of a lesser magnitude which may reflect the smaller corticosterone response. However, as discussed earlier manual restraint initiated a similar corticosterone response in each bird, whilst mechanical restraint resulted in a range of responses. This difference could not be attributed to differences in gonad size as all birds had large testes, although Boyd (2000) reported a larger corticosterone response to manual restraint in female quail with larger gonads compared with birds with small gonads. This result indicates that the size of the corticosterone response may be related to the testosterone concentration of the individual. Previous studies have reported a positive correlation between high testosterone and a large corticosterone response (Boyd, 2000; Heiblum et al., 2000). 3.4.4 Summary The aims of this study were to determine the relationships between the duration of manual restraint and the magnitude of changes in plasma corticosterone and testosterone concentrations in quail, and to examine responses of quail to other stressors. Two minutes of manual restraint had no effect whereas 5, 10 or 15 minutes of restraint led to increased corticosterone and decreased testosterone. Decreases in testosterone in quail and chickens have previously been shown only in birds that were handled on more than one occasion (Wilson et al., 1979; Eskeland and Blom, 1 979; Johnson, 1981; Chapter 2) . The duration of reduced testosterone concentrations was greater than the duration of increased corticosterone, and the rate of testosterone decline was inversely related to initial testosterone concen tra tions. 116 Chapter Three The results are consistent with a transient inhibitory action of corticosterone on the control of testosterone secretion. If the testosterone response to restraint is indeed mediated by corticosterone then this may be directly at the gonads. A reduction in testosterone with no change in LH was shown in the previous chapter, and corticosterone has a direct inhibitory effect on testosterone synthesis in rat leydig cell in vitro (Mann and Orr, 1990) and on steroidogenesis in the quail cloacal gland (Deviche et al., 1982) . Corticosterone responses to stressors in birds are already measured by restraining birds in bags (Wingfield et al., 1992; Wingfield, 1994) or sometimes in a mechanical device (Satterlee and Johnson, 1988). The responses are therefore to artificial stressors, and little is known about the acute corticosterone responses of birds to natural stressors. Responses of quail to situations that might impose social stressors were therefore tested in the current study. SurpriSingly, the introduction of an unfamiliar quail into a bird's home cage was not a stressor, whereas the introduction of a quail into an adjacent cage did induce corticosterone responses in some birds. Previous studies of quail have shown that the introduction of an unfamiliar albino quail into the cage did cause a corticosterone response (Satterlee et al., 1983). The quail used by Satterlee are generally more responsive to stressors than the quail used in the current study G . F. Cockrem pers. comm.), which may account for the different results. The absence of consistent corticosterone responses to social stimuli meant that they would not be used for more detailed studies of responses of quail to natural stressors. Corticosterone responses to manual restraint in which birds were held by the experimenter for 15 minutes were consistently high, whereas responses to mechanical restraint varied markedly between birds. Manual restraint was therefore perceived by the birds to be a stronger emotional stressor than mechanical restraint. The range of responses was similar to that in chickens (Littin and Cockrem, 2001) and great tits (Cockrem and Silverin, 2002a) . This variation in quail corticosterone responses was investigated in detail and quantified in a study reported in the next chapter. Chapter Four Chapter 4 Variation within and between birds in corticosterone responses of Japanese quail 4.1 Introduction 1 17 Corticosterone is the major adrenal glucocorticoid hormone in birds (Carsia and Harvey, 2000). Exposure to a stressor initiates an increase in plasma corticosterone concentrations. In Japanese quail, immobilisation caused corticosterone concentrations to increase within one minute, reach a peak after four minutes and generally remain elevated during 20 minutes of mechanical restraint (Satterlee and Johnson, 1988) . In the previous chapter it was shown that 15 minutes of handling or mechanical restraint resulted in a corticosterone peak at 15 minutes, followed by a return to basal concentrations between 15 and 60 minutes. The results of the previous experiment illustrated that significant variation in the corticosterone response to stressors exists between birds from the same population as reported previously in quail (Satterlee and Johnson, 1988) and willow tits (Silverin, 1997) . Differences in the adrenocortical response have been reported in a wide variety of free-living and domesticated birds. The majority of studies have concentrated on ecological factors such as differences between species (Wingfield et al., 1 992), geography (Silverin, 1997), or breeding or life stages (Wada et al., 1999; Romero et al., 1997; Romero and Remage-Healey, 2000). An area that has received little attention are the differences in hypothalamic-pituitary-adrenal (HPA) axis sensitivity to stressors within and between individual birds under constant conditions. Studies where birds are selected for divergent adrenocortical responses 118 Chapter Four to stressors such as low temperatures (Brown and Nestor, 1973), social stress (Gross and Siegel, 1985), or immobilisation (Satterlee and Johnson, 1988) are evidence of significant differences in the corticosterone response to stress between birds, irrespective of external influencing factors. There is likely to be a genetic basis to this variation in birds, as in other vertebrates (reviewed by Mormede et al., 2003) . In almost all previous studies of corticosterone responses to handling, individual birds were bled on only one occasion. I t has been presumed that the corticosterone response of a bird on a single occasion indicated its sensitivity to handling. However, recent studies in our laboratory on chickens (Littin and Cockrem, 2001) and grea t tits (Cockrem and Silverin, 2002a) were the first to show that individual responses are indeed repeatable and to quantify variation in responses. In addition, these two studies showed that the corticosterone response is generally repeatable within birds. In the previous chapter, the results showed that the size of the corticosterone response varied between quail from the same population. The aim of the present study was to define variation in corticosterone responses within and between Japanese quail, and at the same time determine the repeatability of the response. 4.2 Methods and materials 4.2.1 Animals and housing 4.2. 1 . 1 Source and type of b i rds Eight week-old male Japanese quail (Coturnix coturnix japonica) were purchased from our supplier (Rangitikei Game Birds Ltd., Bulls). The birds had been raised under a long-day photoperiod (15L:90) at a ir temperatures of 20 to 25°C in mixed sex groups. The birds were sexually mature at time of purchase with all birds having large cloacal glands and cloacal foam present. Chapter Four 119 4.2 . 1 .2 Housing conditions Each bird was identified with a numbered leg band and housed in an individual cage measuring 20 cm (W) x 35 cm (D) x 24 cm (H) in animal rooms in the Veterinary Science building. Quail were held on a long day photoperiod (16L :8D; lights on from 0900 - 0100 h). An extractor fan provided ventilation for each room during each experiment. A temperature control unit enabled the air temperature to be maintained at 20°C. Light in each room was provided by two 75W incandescent light bulbs controlled by a 24 hour/7 day time switch (HPM Excel Light Switch and Timer, Cat XL770T). The light intensity in the quail rooms was 130 lux in cages at the top, and 15 lux in cages near the floor of the room (Minolta Illuminance meter). 4.2 . 1 .3 Food and water Quail were provided with fresh water and food (Harvey Farms quail layer mash) ab libitum. The food was the same as that used by Rangitikei Game Birds. 4.2.2 Experimental design Twenty 8 week old male Japanese quail were held with ad libitum access to food and water under long days (16L:8D) a t 20°C for 10 days. The birds were assigned into two groups; birds in the untreated group (n=10) were blood sampled at 0, 15, 30, 60 and 120 min. Birds in the treated group (n=10) were blood sampled at 0 min and handled, handling comprised picking up and putting down the bird continuously for 15 min (Boyd, 2000). Blood samples in treated birds were then collected at 15, 30, 60, and 120 min. Birds were returned to their cage after time 0 in the untreated group, or 15 min in the handled group, and following all subsequent blood samples. Blood sampling and handling was repeated 10 and 20 days after the first instance. Birds were euthanased at the final sample time on day 20 and a terminal blood sample was collected. 4.2 .3 Data collection Blood samples were collected by venipuncture of the brachial vein in the wing with a 25 g needle and 200 J.LI of blood was collected into heparinised capillary tubes. All 120 Chapter Four samples were collected within 1 .5 min from the time the bird was removed from the cage. The final blood sample was collected when the birds were euthanased by stunning followed by decapitation. The blood was stored on ice until centrifuged at 2 000 g for 15 minutes at 4°C (Heraeus Christ 5000S refrigerated centrifuge). The plasma was removed and stored at -20°C until assayed. The width and height of the cloacal protuberance was measured (±0.1 mm) and body weight was measured (±0.1 g) in all birds on the day before the experimental treatment. The reproductive system was removed after euthanasia and the testes weighed (±0.001 g) . The experiment was conducted under a protocol approved by the Massey University Animal Ethics Committee. 4.2.4 Hormone assays Plasma corticosterone concentrations were determined by radioimmunoassay using the methodology outlined in Section 3.2.4. 1 . The inter-assay coefficients of variation for five assays were 7.4, 6.2, and 6.6% for high, medium and low quality controls respectively. 4.2.5 Calculation of i nteg rated corti costerone responses The area under the curve for plasma corticosterone concentrations versus time were calcula ted for each bleed for each bird, using the method described by Cockrem and Silverin (2002a) . The ' total integrated response' (Cockrem and Silverin, 2002a) is a measure of the amount of corticosterone secreted during the sampling period and is expressed as nglml·min. The 'corrected integrated response' was also calculated by multiplying the corticosterone concentration at time 0 by the duration of experiment and subtracting this from the total integrated response area. Given that corticosterone peaks at 15 min and concentrations have largely returned to basal by 30 min, the corrected integrated response was calculated between 0 and 30 min and was not calculated for birds where subsequent corticosterone concentrations declined below the initial concentration. Areas under the curve were calculated using the trapezoid rule using GraphPad Prism 4.0 (GraphPad Software Inc.). Chapter Four 121 4.2 .6 Statistical analyses All results are presented as the mean ± SE (standard error) or individual data where appropriate. Plasma corticosterone concentrations were loglo transformed to minimise heteroscedasticity. All the data were examined for normality using Shapiro-Wilks tests and for equality of variance using Levene's homogeneity of variance test. The changes in plasma corticosterone in birds sampled on three occasions were analysed using repeated measures two way ANOVA with time (0, 15, 30, 60 and 120 min) and bleed (first, second and third) as the grouping factors. Post-hoc comparisons were made between times for each bleed and between bleeds for each time using univariate F tests. Differences between bleeds in integrated corticosterone responses were examined by one way ANOV A followed by Bonferroni pairwise comparisons between pairs of bleeds. Differences between birds in corticosterone concentrations at each time and in integrated corticosterone responses were examined by one way ANOV A. Data not complying with normality or variance criteria were compared using a Kruskal-Wallis non-parametric test followed by Mann-Whitney U independent group comparisons tests, or Friedman's test for repeat measures data. All comparisons of means were conducting using SPSS ™ 1 1 .0 (SPSS Inc., Illinois) or SystaCM B.O (SPSS Inc., Illinois). 4.3 Resu lts 4.3 .1 Cort icosterone responses The corticosterone response to handling was largely repeatable on three occasions within individual quail (Figure 4.1), with only two birds out of the 10 (#3 and 7) exhibiting marked variation between bleeds. Plasma corticosterone concentrations were low in the majority of birds when they were first sampled, although four birds had higher initial concentrations on the second sampling occasion. Plasma 122 Chapter Four corticosterone increased 15 min later in 90% of birds in each of the three blood sampling periods (Figure 4.1 and 4.2), then decreased between 15 and 30 min in all birds except for one (third bleed for bird 4; Figure 4.1). Corticosterone concentrations then remained relatively constant in all birds between 30 and 120 min (Figure 4.2). The corticosterone response to handling at 15 min was markedly larger for the second bleed in three birds (Nos. 2, 3, 8) and one bird for the third bleed (No. 2). Corticosterone was significantly higher at 15 min for the second bleed in two other birds (Nos. 5 and 7), but the size of the increase was no different from the first bleed. Initial corticosterone concentrations in bird no. 4 were high on a ll three occasions and declined after 1 5 min on the first and third bleed. On the second bleed, corticosterone remained high before declining by 30 min and remaining relatively constant. Mean corticosterone concentrations in individual birds ranged from 1 .01 ± 0.64 to 7.57 ± 0.28 ng/ml at 0 min, 4.57 ± 1 .76 to 9.63 ± 1 .67 ng/ml at 15 min, 1 .85 ± 0.66 to 3.76 ± 1 .84 ng/ml at 30 min, 1 .09 ± 0.38 to 2.42 ± 0.68 ng/ml at 60 min, and from 0.91 ± 0.12 to 2.71 ± 1 .25 ng/ml at 120 min (Figure 4.2) . In all but bird no. 4, mean corticosterone concentrations increased after 15 min handling, then decreased to initial concentrations by 30 min and remained relatively constant. Mean corticosterone concentrations remained low and relatively unchanged in birds that returned to their cages between samples (untreated; Figure 4.3a). In untreated birds corticosterone varied between bleeds and between times, but there was no significant interaction between bleed and time (Table 4.1) . In handled birds, corticosterone varied between bleeds and between time, and there was a significant interaction between bleed and time (see Table 4.2 for a ll statistics for this analysis) . Mean corticosterone concentrations at 0 min did not differ between bleeds in handled birds (Figure 4.3b) and increased after 15 min for al l three bleeds. Corticosterone concentrations were higher at 15 min in the second bleed than in either of the other two bleeds. Plasma corticosterone declined after 15 min in all bleeds, and for the first bleed declined below initial concentrations and those of the other two bleeds at 30, 60 and 120 min. Elevated corticosterone concentrations at Chapter Four 123 15 min for all bleeds were significantly higher than in untreated birds for the same bleed, although corticosterone concentrations in the second bleed were significantly higher in handled birds versus untreated birds throughout the sampling period (Table 4.3). The total integrated corticosterone responses varied between bleeds in handled birds (F2,29 = 34.159, P < 0.001; Figure 4.4b), with the response for the second bleed greater than the first and third bleeds (post-hoc Tukey tests, p < 0.001 and p = 0.028), and the third greater than than the first bleed (post-hoc Tukey tests, p < 0.001). Some of the birds had to be excluded from the calculation of the corrected integrated corticosterone response between 0 and 30 min due to corticosterone falling below initial concentrations. There were no differences in corrected corticosterone responses from 0 to 30 min between the three bleeds (F2,13 = 1 .029, P = 0.389). The total integrated corticosterone responses also varied in untreated birds (F2,29 = 15.746, P < 0.001; Figure 4.4a), with the response in birds for the third bleed greater than the responses for the first and second bleeds (post-hoc Tukey tests, p = 0.001 and p < 0.001 ). Corrected responses could be calculated for only a few birds, but did not appear to differ between bleeds. 4.3.2 Quantification of variation in corticosterone responses Mean corticosterone concentrations for each time and the mean integrated corticosterone responses were calculated for each quail (Figure 4.5 and 4.6). There were close to significant differences between birds in corticosterone concentrations at the 0 and 15 min sampling times (see Table 4.4 for statistics for this analysis). The initial corticosterone concentrations in bird no. 4 was higher than most of the o ther birds. Using the method described by Cockrem and Silverin (2002a), the ratio of the largest to smallest value for corticosterone concentrations at each time and for integrated corticosterone responses were calculated (Table 4.5). The ratio was largest in the first sample, and was similar for all subsequent sampling times. The 124 Chapter Four ratio for the corrected integrated response was twice that for the total integrated response. Using the method described by Cockrem and Silverin (2002a), variation within and between birds was quantified by calculating the coefficients of variation (CV) for corticosterone concentrations and integrated responses for each bird at each time (Table 4.6 and 4.7). Variation between birds, measured as the CVs of the mean of individual bird means, was greatest for corticosterone at 0 min (65.5%), and smaller at IS, 30, 60 and 120 min (24.5, 27.7, 24.4 and 24.5%). Variation within birds, measured as the means of the CVs of individual birds, was lower for corticosterone at 15 min (38.0%) than at other times. Variation between birds was lower for total integrated corticosterone responses (14.8%) than for corrected responses (36.0%), while variation within birds was higher for total integrated responses (45.0%) than for corrected responses (28.6%). Chapter Four E - 0> C -.- ID C e ID -+-' (j) o u t o U 1 5 1 1 0 o �-.--.----.--------�r 15 3 15 1 5 7 1 5 9 1 0 ,', iR.\. 5 ! \\ '�'" n" \\ 4 '.�.: ..... <... . .. . .. , . ,., .. . • • . ....... . .... ..... ............ :;w o r o 1 5 30 60 1 20 2 A .. ; \ " .. " .. .... " ..... . f \ . . ... ...... . . ...... . , b .... • .... · ... • ..... o._._·_·_·_·_·_·_ .. .a 4 .. . ......... .• . � ... �:::.-:�::-:.:-:.�:�.:::.�::-:.��:.� 6 1 0 o 1 5 30 60 1 20 Time (min) - First bleed _ .-& • • Second bleed --0- Third bleed 125 Figure 4.1 Plasma corticosterone in individual male quail handled for 15 min on three separate occasions at 1 0 day intervals. Results are plotted as raw data, 126 15 1 10 5 0 15 3 10 5 ......- E - Cl 15 C 5 -- ID C 10 e ID ........ 5 en 0 0 t:: 0 0 () 15 7 10 5 0 15 9 10 , /\ 0 0 15 30 60 1 20 2 4 6 8 1 0 !\ 0 15 30 Time (min) Chapter Four 60 120 Figure 4.2 Mean plasma corticosterone concentrations in individual male quail handled for 15 min on three separate occasions at 1 0 day intervals. Results are plotted as means ± standard error. Chapter Four 1 2 1 0 8 6 4 E - 0) C 2 -- Q) C (a ) Untreated - Bleed 1 . _-- _ . B leed 2 ..... - B leed 3 J.... -- --1-- ... :_----1 '---._ __-- --- - -----� - - - .... - - - -- - - - - - - -+ - -f- - - - - - -.� - - - - - - - - - - .�.- - - - . - - .. - . . - . . . - . . . . - - . . - . . . - . e o���--�--�--------�------------------�--- Q) 1 2 ... en o C,) t:: 1 0 o o 8 6 4 2 ( b) J, . , . , .' " . , : \ . , : ' . , . , " " , , , , 't.- . Hand l i ng - Bleed 1 . _-- _ . Bleed 2 ..... - B leed 3 �\'*- .... . .. ... . . . . .1... . _ . .. . . -.=.":. �: :: :.:.: :.: : :.-:.:::.: 1 T- ·- -- - - -- ·- '1"- t o�--�--�--�--------�------------------�--- o 1 5 30 60 1 20 Time (min) 127 Figure 4.3 Mean plasma corticosterone concentrations in individual male quail that were returned to their cages between samples (untreated; a) or handled for 15 min (b) on three separate occasions at 10 day intervals. Results are plotted as means ± standard error. 128 300 (a) 200 ...- c E E - 1 00 0) C -- Q.) Cl) C 0 Cl. 0 Cl) Q.) � Q.) c 0 � Q.) 600 ...... Cl) 0 (b) u ...... 500 � 0 U "0 400 Q.) ...... CO � 300 0) Q.) ...... C 200 1 00 0 - A1l n = 1 0 n = 6 Total All n = 1 0 n = 4 Total Chapter Four Untreated - Bleed 1 1 " ,"',,"" '1 B leed 2 c:::::::::J B le ed 3 n = 1 n = 3 Corrected Handl ing - Bleed 1 1-,,,, 1 B leed 2 c:::::::::J B leed 3 n = 6 n = 4 Corrected Figure 4.4 Total and corrected corticosterone responses of male quail that were (a) returned to their cages between samples (untreated); or (b) handled for 15 min on three separate occasions at 10 day intervals. Total integrated corticosterone responses were calculated from the areas under the corticosterone vs time curves for each bird. Corrected integrated corticosterone responses were calculated by subtracting the area under the corticosterone vs time (0-30 min) curve due to basal corticosterone concentrations from the total area. Results are plotted as means ± standard error for 10 birds. Chapter Four 1 5 o min 1 0 : • • • 1 • • • • 1 5 1 0 o � 1 5 0> C --- ID 1 0 C e 1 5 min L-____________________________ _ 30 min ID 1 : • • • • • • • • • • o U 1 5 60 min 10 5 1 5 1 20 min 10 5 2 3 4 5 6 7 8 9 1 0 Bird 129 Figure 4.5 Mean corticosterone concentrations at 0, 15, 30, 60, and 120 min in individual male quail handled for 15 min on three separate occasions at 10 day intervals. Results are plotted as means ± standard error. 130 Chapter Four Total integrated response 600 400 a.> Cl) 200 C 0 0- Cl) a.> L- a.> 0 c ....- 0 C 2 3 4 5 6 7 8 9 1 0 L- a.> E Bird ........ Cl) - o E u _ .- 0> t:::: c Corrected integrated response 0 '-'"" U "'0 a.> ........ ro 200 L- Ol a.> ........ c 1 00 0 I 2 3 4 5 6 7 8 9 10 Bird Figure 4.6 Mean total and corrected (0-30 min) integrated corticosterone responses in individual male quail handled for 15 min on three separate occasions at 10 day intervals. The number of corrected integrated responses per bird are shown inset in the columns. Results are plotted as means ± standard error. Chapter Four 131 Table 4.1 Statistical analysis for variation in the corticosterone response in birds that were returned to their cages between samples (untreated) on three separate occasions at 1 0 day intervals. Effect F d[ r. Bleed 8.755 2,27 0.001 � O' Time 7.333 4,108 < 0.001 � O' Interaction of bleed and time 1 .614 8,108 0.129 Com�arisons within treatments for each bleed o vs 15 min Bleed 1 1 .481 1,27 0.234 Bleed 2 0.195 1,27 0.662 Bleed 3 0.335 1 ,27 0.567 15 vs 30 min Bleed 1 0.224 1 ,27 0.640 Bleed 2 0.053 1,27 0.819 Bleed 3 0.084 1 ,27 0.774 30 vs 60 min Bleed 1 22.515 1 ,27 < 0.001 � O' Bleed 2 6.797 1 ,27 0.015 � Bleed 3 2.399 1 ,27 0.133 60 vs 120 min Bleed 1 1 1 .079 1,27 0.003 * O' Bleed 2 0.273 1,27 0.606 Bleed 3 2.092 1,27 0.160 o vs 30 min Bleed 1 5.932 1 ,27 0.022 * Bleed 2 0.113 1,27 0.739 Bleed 3 1 .555 1,27 0.223 o vs 60 min Bleed 1 7.395 1,27 0.011 � Bleed 2 7.741 1,27 O.OlD � Bleed 3 0.292 1,27 0.593 o vs 120 min Bleed 1 1 .238 1,27 0.276 Bleed 2 6.866 1,27 0.014 � Bleed 3 1 .549 1,27 0.224 Comparisons between bleeds for each time F df p O min Bleed 1 vs Bleed 2 0.194 1,27 0.663 Bleed 1 vs Bleed 3 3.013 1,27 0.094 Bleed 2 vs Bleed 3 1 .677 1,27 0.206 15 min Bleed 1 vs Bleed 2 0.898 1,27 0.352 Bleed 1 vs Bleed 3 1 .812 1,27 0.189 Bleed 2 vs Bleed 3 5.262 1,27 0.030 * 30 min Bleed 1 vs Bleed 2 2.280 1 ,27 0.143 Bleed 1 vs Bleed 3 2.527 1,27 0.124 Bleed 2 vs Bleed 3 9.608 1 ,27 0 .004 O' * 60 min Bleed 1 vs Bleed 2 0.195 1,27 0.662 Bleed 1 vs Bleed 3 13 .092 1,27 0.001 * * Bleed 2 vs Bleed 3 10.092 1,27 0.004 . * 120 min Bleed 1 vs Bleed 2 4.888 1,27 0.036 • Bleed 1 vs Bleed 3 5.251 1,27 0.030 • Bleed 2 vs Bleed 3 20.270 1,27 < 0.001 . * Note. The first three rows show the results o f the two-way repeat measures ANOVA for the main effects and their interaction. The remaining rows show the results of post-hoc comparisons of times for each bleed and comparisons of bleed at each time. Significant differences (p < 0.05) are marked with a single asterisk, while highly significant differences (p < 0.01 ) are marked with a double asterisk. 132 Chapter Four Table 4.2 Statistical analysis for variation in the corticosterone response in birds that were handled for 15 min on three separate occasions at 10 day intervals. Effect F dt r Bleed 38.735 2,27 < 0.001 * * Time 30.958 4,108 < 0.001 ** Interaction of bleed and time 4.281 8,108 0.001 ** Comearisons within treatments for each bleed o vs 15 min Bleed 1 19.463 1 ,27 < 0.001 * * Bleed 2 13.824 1 ,27 0.001 ** Bleed 3 5.678 1,27 0.024 * 15 vs 30 min Bleed 1 1 10.052 1 ,27 < 0.001 * * Bleed 2 23.395 1 ,27 < 0.001 * * Bleed 3 23.589 1,27 < 0.001 * * 30 vs 60 min Bleed 1 7.199 1,27 0.012 * Bleed 2 9.550 1,27 0.005 * * Bleed 3 0.000 1 ,27 0.984 60 vs 120 min Bleed 1 1 .002 1 ,27 0.326 Bleed 2 1 .232 1 ,27 0.277 Bleed 3 0.505 1 ,27 0.483 o vs 30 min Bleed 1 1 .294 1 ,27 0.265 Bleed 2 1 .664 1 ,27 0.208 Bleed 3 0.026 1 ,27 0.874 o vs 60 min Bleed 1 8.056 1 ,27 0.009 ** Bleed 2 0.896 1,27 0.352 Bleed 3 0.025 1 ,27 0.877 o vs 120 min Bleed 1 9.013 1 ,27 0.006 * * Bleed 2 0.026 1 ,27 0.874 Bleed 3 0.079 1 ,27 0.781 Comparisons between bleeds for each time F df p O min Bleed 1 vs Bleed 2 2.678 1 ,27 0.11 3 Bleed 1 vs Bleed 3 1 .165 1 ,27 0.290 Bleed 2 vs Bleed 3 0.310 1,27 0.582 15 min Bleed 1 vs Bleed 2 6.825 1,27 0.015 * Bleed 1 vs Bleed 3 1 .771 1 ,27 0.194 Bleed 2 vs Bleed 3 15.550 1,27 0.001 * * 30 min Bleed 1 vs Bleed 2 43.459 1,27 < 0.001 * * Bleed 1 v s Bleed 3 1 1 .968 1 ,27 0.002 * * Bleed 2 v s Bleed 3 9.815 1 ,27 0.004 * * 60 min Bleed 1 vs Bleed 2 25.019 1,27 < 0.001 * * Bleed 1 v s Bleed 3 26.636 1 ,27 < 0.001 * * Bleed 2 v s Bleed 3 0.025 1 ,27 0.875 120 min Bleed 1 vs Bleed 2 33.523 1 ,27 < 0.001 * * Bleed 1 v s Bleed 3 31 .645 1 ,27 < 0.001 * * Bleed 2 v s Bleed 3 0.027 1 ,27 0.871 Note. The first three rows show the results of the two-way repeat measures ANOV A for the main effects and their interaction. The remaining rows show the results of post-hoc comparisons of times for each bleed and comparisons of bleed at each time. Significant differences (p < 0.05) are marked with a single asterisk, while highly significant differences (p < 0.01) are marked with a double asterisk. Chapter Four 133 Table 4.3 One-way ANOVA analyses for comparison of untreated and handled corticosterone concentrations for each bleed at each time. Comparisons between treatments for each bleed F df p Bleed 1 O min 0.1 91 1,18 0.667 15 min 126.210 1,18 < 0.001 • • 30 min 3.621 1,18 0.073 60 min 0.597 1,18 0.450 120 min 1 1 .667 1,18 0.003 • • Bleed 2 O min 5.641 1,18 0.029 • 15 min 69.961 1,18 < 0.001 * * 30 min 84.544 1,18 < 0.001 • • 60 min 20.781 1,18 < 0.001 • • 120 min 16.582 1,18 0.001 • • Bleed 3 O min 1 .012 1,18 0.328 15 min 13.482 1,18 0.002 • • 30 min 0.038 1,18 0.848 60 min 1 .964 1,18 0.178 120 min 0.540 1,18 0.472 Note. One-way ANOVA for the main effects of treatment (untreated vs handled) for each bleed at each time. Significan t differences (p < 0.05) are marked with a single asterisk, while highly significant differences (p < 0 .01 ) are marked with a double asterisk. Table 4.4 One-way ANOVA analyses for corticosterone concentrations for each time and integrated corticosterone responses in birds that were handled for 15 min on three separate occasions at 10 day intervals. Variable F df P Plasma corticosterone concentrations Time O min 2.386 9,29 0.051 15 min 1 .211 9,29 0.342 30 min 0.555 9,29 0.817 60 min 0.209 9,29 0.990 120 min 0.260 9,29 0.979 Integrated corticosterone responses Total 0.266 9,29 0.977 Corrected 3.467 7,13 0.075 134 Chapter Four Table 4.5 Smallest and largest mean values of corticosterone concentrations and integrated corticosterone responses for individual quail. O min Smallest value Bird number 9 Mean 1 .01 Largest value Bird number 4 Mean 7.57 Ratio of smallest to largest value Mean 7.5 Corticosterone 1 5 min 4 4.57 2 9.63 2. 1 (nglml) 30 min 8 1 .85 2 3.76 2.0 60 min 1 20 min 9 9 1 .09 0.91 4 1 2.42 2.71 2.2 3.0 Integrated response (nglml'min) Total Corrected 9 7 274.66 52.55 2 2 408.50 1 70.55 1 .5 3.2 Table 4.6 Coefficients of variation of mean corticosterone concentrations and integrated corticosterone responses within and between quail. Corticosterone Integrated response O min 15 min 30 min 60 min 1 20 min Total Corrected All birds Mean 2.81 7.13 2.51 1 .72 2.05 323.29 1 10.76 SE 0.41 0.52 0.29 0.23 0.27 25.79 1 2.32 CV 79.47 40.05 63.73 72.86 73.11 43.70 41 .62 n 30 30 30 30 30 30 14 Variation between birds (means of the individual bird means) Mean 2.81 7.13 2.51 1 .72 2.05 323.29 1 10.76 SE 0.58 0.55 0.22 0.13 0.16 15 .15 14.09 CV 65.49 24.53 27.51 24.42 24.50 14.82 35.98 n 10 10 10 1 0 1 0 10 8 Variation within birds (means of the individual bird CVs) Mean 60.71 37.95 66.29 75.21 72.61 45.04 28.55 SE 12.57 5.17 7.25 7.77 7.38 5.60 5.49 n 10 10 10 1 0 1 0 10 5 Chapter Four 135 Table 4.7 Coefficients of variation of mean corticosterone concentrations and integrated corticosterone responses within and between quail. Corticosterone at 0 min Corticosterone a t 15 min Corticosterone at 30 min Mean SE CV n Mean SE CV n Mean SE CV n Bird (nglml) (%) (nglml) (%) (nglml) (%) 1 2.23 0.58 45.19 3 5.33 0.73 23.60 3 3.04 0.45 25.86 3 2 1 .57 0.12 13.03 3 9.63 1 .67 30.02 3 3.76 1 .84 84.61 3 3 1 .46 1 .03 122.41 3 6.27 1 .56 43.02 3 2.84 0.97 59.12 3 4 7.57 0.28 6.52 3 4.57 1 .76 66.82 3 2.1 9 0.99 78.75 3 5 3.54 2.02 99.05 3 9.46 2.37 43.40 3 3.35 0.62 31 .88 3 6 2.38 0.97 70.87 3 6.56 0.88 23.17 3 2.04 0.73 61 .71 3 7 2.82 1 .00 61 .31 3 6.15 2.23 62.78 3 2.29 1 .04 79.01 3 8 2.97 0.85 49.79 3 9.14 1 .60 30.38 3 1 .85 0.66 61 .65 3 9 1 .01 0.64 108.90 3 7.21 0.79 19 .04 3 1 .87 0.94 86.75 3 10 2.53 0.44 29.94 3 7.01 1 .51 37.32 3 1 .86 1 .01 93.59 3 Corticosterone at 60 min Corticosterone a t 120 min Mean SE CV n Mean SE CV n Bird (nglml) (%) (nglml) (%) 1 1 .70 0.76 77.18 3 2.71 1 .25 79.83 3 2 2.22 1 .45 1 12.71 3 2.26 0.97 74.74 3 3 1 .79 0.35 34.20 3 1 .88 0.60 54.96 3 4 2.42 0.68 48.96 3 1 .9 1 0.94 85.57 3 5 1 .79 0.72 69.77 3 2.59 1 .38 92.22 3 6 1 .47 0.74 86.75 3 2.08 0.91 75.50 3 7 1 .23 0.55 77.24 3 2.1 1 1 .04 85.55 3 8 1 .09 0.38 60.75 3 2.29 1 .36 103.17 3 9 1 .99 1 .27 1 10.86 3 0.91 0.12 23.63 3 10 1 .47 0.63 73.66 3 1 .74 0.51 50.93 3 Total integrated response Corrected integrated response (0 - 30 min) Mean SE CV n Mean SE CV n Bird (nglml) (%) (nglml) (%) 1 323.00 73.73 39.54 3 52.55 7.45 24.55 3 2 408.50 155.91 66.10 3 170.55 17.45 14.47 2 3 306.03 16.92 9.58 3 1 19.65 39.55 46.75 2 4 340.73 68.83 34.99 3 5 402.36 1 19.02 51 .23 3 93.69 15.31 23.1 1 2 6 290.80 95.91 57.12 3 72.23 1 7 283.56 108.88 66.50 3 8 318.83 90.01 48.90 3 158.30 1 9 274.66 77.76 49.04 3 1 13.92 27.29 33.87 2 1 0 284.40 44.94 27.37 3 105.20 1 136 Chapter Four 4.4 D iscussion 4.4.1 Variation in corticosterone responses The results of this study contribute to our understanding of individual variation of the adrenocortical response from work previously conducted on chickens and great tits in our laboratory. Plasma corticosterone in quail increased after handling and the response was largely repeatable in individual quail, although the profile of the corticosterone response and basal corticosterone varied between birds. Variation in basal corticosterone concentrations between birds has been reported previously (Brown and Nestor, 1973; Schwabl, 1999) . Earlier studies have concentrated on the influence of environmental factors such as season, geography or breeding state on the temporal modulations of the adrenocortical responses in birds (Wingfield et al., 1992; Astheimer et al., 1995; Romero et al., 1997; Remage-Healey and Romero, 2000). These studies have noted marked variation of the corticosterone response between birds but have not determined if the observed corticosterone responses were repeatable in the same bird. Under the same conditions, different individuals can have a range of corticosterone responses to an identical s tressor. Vleck et al. (2000) reported that individual Adelie penguins (Pygoscelis adeliae) maintained distinct differences in basal corticosterone concentrations throughout the breeding season. The current study confirms that individual differences in corticosterone are repeatable within individuals (e.g. different quail can have low or high corticosterone concentrations in response to mechanical restraint (Satterlee and Johnson, 1988; Jones et al., 2000)) . In the current study, plasma corticosterone concentrations increased in all but one quail after 15 minutes of handling. Similar results were reported for chickens (Littin and Cockrem, 2001) . Initial and basal corticosterone concentrations were low and consistent with those reported in the previous chapter and by Satterlee and Johnson (1988) . One bird had high initial corticosterone concentrations on all three blood sampling sessions, which declined after the initial sample in contrast to all other birds. This bird was sampled within one minute of being removed from a Chapter Four 137 cage, making it unlikely that the blood sampling procedure was the cause of such high basal corticosterone concentrations. In the remainder of birds, corticosterone generally declined to initial concentrations between 15 and 30 minutes after the onset of handling and remained low between 30 and 120 minutes, as reported in the previous chapter and by Boyd (2000). Selecting birds for divergent corticosterone concentrations has been used in several studies to produce strains with low or high basal or adrenocortical responses to a stressor. Selection for low or high basal corticosterone concentrations or responses has been made in chickens (Edens and Siegel, 1975), turkeys (Brown and Nestor, 1973) and quail (Satterlee and Johnson, 1 988) . Such studies emphasize the genetic origin of variation of the adrenocortical response and differences in hypothalamic­ pituitary adrenal (HPA) axis sensitivity to stressors between individuals. In the current study all birds were held and handled under identical conditions, with the observed variation especially that of bird no. 4 indicating inherent genetic differences in adrenocortical responsiveness within the birds. Variation of the adrenocortical response between individuals has been documented in a wide range of vertebrates, and the genetic factors controlling it are currently being studied in rodents (reviewed by Mormede et al., 2003) . Cockrem and Silverin (2002a) reported that the corticosterone response decreased over three repeat bleed occasions in great tits, indicating possible habituation to handling. In that study, great tits were captured from a free-living situation. Repeated handling of chickens on successive days, a domesticated species, resulted in no change in the corticosterone response to handling (Beuving and Vonder, 1978; Littin and Cockrem, 2001) . Quail in the current study had similar initial corticosterone concentrations for all three bleeds as reported for chickens handled on four occasions (Littin and Cockrem, 2001) and great tits on three occasions (Cockrem and Silverin, 2002a). In contrast, previous studies in chickens have also reported a reduction in basal corticosterone concentrations in chickens after one week of twice daily handling (Webb and Marshaly, 1984) and a reduction in the corticosterone response over three weeks with handling repeated ten times per week (Freeman and Manning, 1979). 138 Chapter Four 4.4.2 Quantification of variation i n corticosterone responses Variation in the corticosterone response to handling between and within individual quail was quantified using the approach applied previously in our laboratory in chickens (Littin and Cockrem, 2001) and great tits (Cockrem and Silverin, 2002a). Mean corticosterone concentrations did not differ between quail at any of the sampling times, although differences between birds at 0 minutes were almost significant with high concentrations in one bird. This indicates that corticosterone concentrations were more consistent between birds during the corticosterone response than were basal concentrations, whereas peak concentrations were the most consistent. In chickens, corticosterone varied significantly at 15 and 40 minutes but not at 0 minutes (Littin and Cockrem, 2001) . The ratios of the largest to the smallest mean corticosterone concentrations of individual birds were also more consistent between 15 and 120 minutes than at 0 min. Neither the total or corrected integrated corticosterone responses differed between birds, whereas both varied in chickens (Littin and Cockrem, 2001) and the total but not the corrected response varied between great tits (Cockrem and Silverin, 2002a) . The results of the current experiment indicate that the corticosterone responses were more consistent between individual quail than great tits and chickens. Variation between and within birds was also quantified for corticosterone concentrations and integrated responses using the method described by Cockrem and Silverin (2002a). The coefficients of variation between birds for corticosterone concentrations between 15 and 120 minutes were relatively consistent and all were lower than for concentrations at 0 minutes. These results are similar to those reported in great tits (Cockrem and Silverin, 2002a), and support our earlier findings that the corticosterone concentrations during the adrenocortical response are more consistent than initial corticosterone concentrations. The coefficient of variation within birds for corticosterone at 15 minutes was smaller than for concentrations before and after the peak, whereas in great tits initial concentrations varied most within birds (Cockrem and Silverin, 2002a), and in chickens the coefficients were largest at 0 and 15 min (Littin and Cockrem, 2001). Cha pter Four 139 Calculating the coefficients of variation for all variables of the corticosterone response is a useful tool for quantifying the variation of the corticosterone responses between and within birds, and allows comparisons between species. These results showed that despite significant variation in the response within birds, variation in the response between birds is relatively consistent. There was considerably more variation in initial corticosterone concentrations, than in the response, as found for great tits (Cockrem and Silverin, 2002a) but not chickens (Littin and Cockrem, 2001) . 4.4.3 Summary This study was conducted to quantify variation within and between quail in their corticosterone responses to handling. Responses of individ ual birds were generally repeatable, as previously found in chickens (Littin and Cockrem, 2001) and great tits (Cockrem and Silverin, 2002a). The selection of lines of quail for low or high corticosterone responses (Satterlee et al., 1988) indicates a genetic basis for variation in corticosterone responses. Heritable variation in corticosterone responses between birds could be due to variation in hypothalamic responses to a stimulus, variation in pituitary gland responses to hypothalamic releasing hormones (CRF, AVT and MT in birds), variation in adrenal gland responses to ACTH, or to variable responses at more than one level of the HPA-axis. Variation in pituitary and adrenal responses have been reported in other species (Carsia et al., 1988; Astheimer et al., 1992; Romero et al., 1998a), but have not been investigated in quail. The study reported in the next chapter was therefore conducted to examine the sources of variation in corticosterone responses in quail. Chapter Five Chapter 5 Adrenal and pituitary responsiveness in quail with differing corticosterone responses to mechanical restraint 5 . 1 Introduction 141 In Chapter 3 and 4, the size of the corticosterone response to mechanical restraint or handling was shown to vary significantly between individuals. Individual variation of the corticosterone response is thought to be partly due to differences at some level of the HPA-axis (Astheimer et al., 1994; Romero et al., 1998). In the present study, quail with low or high corticosterone responses to mechanical restraint were used to determine if the adrenal and pituitary levels of the HPA-axis differed in quail with divergent corticosterone responses. In many free-living passerines, the corticosterone response to stressors change markedly during different stages of the life cycle (Romero et al., 1997; Romero et al., 1998a; Romero et al., 1998b; Romero and Wingfield, 1999) . However, the variation considered in this study is that which exists between birds irrespective of season, developmental or reproductive state. Variation in the adrenocortical response to a stressor exists between individual quail in a random-bred quail population such as the quail sourced for our previous studies. The selection of low and high adrenocortical response lines over many generations makes such lines an ideal model for testing hypotheses on the relationship between behavioural, physiological and genetic traits and the relationship between different aspects of 142 Chapter Five stress and the adrenocortical response Gones et al., 1 992a; Jones et al., 1994a; Satterlee and Jones, 1 997; Jones et al., 2000; Od eh et al., 2003a; Odeh et al., 2003b). Despite the well established differences between individual quail in adrenocortical responsiveness to stressors, few studies have been conducted to identify the physiological basis of these differences. Changes in plasma corticosterone concentrations are the main indicator used to measure activation of the HPA-axis to a stressor. In the bird, corticosterone secretion is stimulated by adrenocorticotrophic hormone (ACTH) which is released from the anterior pituitary, which in turn is stimulated by corticotrophin-releasing factor (CRF) released from the hypo thalamus (reviewed by Carsia and Harvey, 2000). The hypothalamic-hypophysiotrophic factors arginine vasotocin (AVT) and mesotocin (MT) also stimulate ACTH secretion or have a permissive role in the stimulation of ACTH by CRF (Castro et al., 1986; Westerhof et al., 1 992; Romero and Wingfield, 1998). In turn, the release of many of these factors is influenced by feedback from glucocorticoids. Differences between birds in their corticosterone responses could be mediated at several levels of the HPA-axis, adrenal sensitivity to ACTH; pituitary sensitivity to CRF, arginine vasotocin (A VT) or mesotocin (MT); or the perception of a stressor. The current experiments were conducted to determine the level or levels of the HPA-axis at which variation between birds in their corticosterone responses arise. Corticosterone responses to mechanical restraint were measured in 70 quail and the 20 birds with the lowest response (lowest 30%) and 20 birds with the highest response (top 30%) identified . The two different response groups were tested for differences in adrenal or pituitary responsiveness to doses of ACTH or CRF that stimulate a sub-maximal corticosterone response similar to tha t observed when birds undergo mechanical restraint. Corticosterone responses to ACTH and CRF treatment were measured in these birds to determine relationships between adrenal sensitivity to ACTH, pituitary sensitivity to CRF and the size of individual corticosterone responses to mechanical restraint. Ovine CRF was uses as avian CRF has not been sequenced and avian pituitary cells respond to ovine CRF (Carsia et al., 1986). A synthetic adrenocorticotrophin (Synacthen™, synthetic peptide Chapter Five 143 corresponding to amino acids 1 -24 from the N-terminal region of human ACTH) was used to test adrenal sensitivity; this form of ACTH stimulates corticosterone secretion in chickens (Littin and Cockrem, 2001) and ducks (Harvey et al., 1980; NoirauIt et al., 1999; Faure et al., 2003). 5.2 Methods and materials 5.2 .1 An imals and hous ing 5.2. 1 . 1 Source and type of bi rds Seventy 7 week-old male Japanese quail (Coturnix coturnix japonica) were purchased from a suppher (Canter Valley Farms, Christchurch) and held in quail rooms for 2 weeks before any manipulation. The birds had been raised under a long-day photoperiod (15L:9D) at air temperatures of 20 to 25°C in mixed sex groups. The 7 week-old birds were sexually mature at time of purchase with large c10acal glands and c10acal foam present. 5.2. 1 .2 Housing conditions Each bird was identified with a numbered leg band. They were housed in individual cages measuring 20 cm (W) x 35 cm (D) x 24 cm (H) in animal rooms in the Veterinary Science building. Quail were held on a long day photoperiod (16L:8D; lights on from 0900 - 0100 h). An extractor fan provided ventilation for each room. A temperature control unit enabled the air temperature to be maintained at 20°C. Light in each room was provided by two 75W incandescent light bulbs controlled by a 24 hour/7 day time switch (HPM Excel Light Switch and Timer, Cat XL770T). The light intensity in the quail rooms was 130 lux in cages at the top, and 1 5 lux in cages near the floor of the room (MinoIta Illuminance meter) . 5.2.1 .3 Food and water Quail were provided with fresh water and food (Harvey Farms quail layer mash) ab libitum. 144 Cha pter Five 5.2.2 Experimental design Timel ine of Chapter Five treatments: Week 0 2 3 4 5 6 7 8 9 Age (weeks) 7 8 9 1 0 1 1 1 2 1 3 1 4 1 5 1 6 � � � � � Study Study Study Study Study 5.2.2.1 5.2.2.2.1 5.2.2.2.2 5.2.2.3.1 5.2.2.3.2 Identification of ACTH ACTH CRF CRF low/high dose challenge of dose challenge of response quail response IOW/high response low/high response quail response quail 5.2.2. 1 Identification of birds with low or high corticosterone responses to mechanical restraint in individual quai l The adrenocortical responses of 70 nine week old male Japanese quail were determined by subjecting each bird to 15 minutes of immobilisation by mechanical restraint in a metal cage using the method described in Chapter 4. Blood samples were collected immediately after the bird was removed from the quail room (0 min), and at 15 and 30 min) . Corticosterone concentrations were measured in all blood samples by radioimmunoassay. Twenty birds with small corticosterone responses « 3 ng/ml increase between 0 and 15 min) and 20 birds with high responses (> 5 ng/ml increase between 0 and 15 min) were identified and held for 14 days before use in further experiments. Quail with corticosterone responses < 3 and > 5 ng/ml between 0 and 15 min were chosen as the 20 lowest versus 20 highest responders to mechanical restraint respectively. 5.2.2.2 Relationship between the adrenocortical response and adrenal sensitivity to ACTH 5 .2 .2 .2 . 1 Relationship between plasma corticosterone and dose of ACTH Forty male Japanese quail were randomly selected from the pool of 70 quail and were assigned into five treatment groups (n = 8 per group). The group allocation and known corticosterone response of each bird (as identified in Section 5.2.2.1 ) was checked to avoid placing a large proportion of low or high corticosterone response birds into one group. The corticosterone response the birds in each group s iden These groups received a single 100 III injection of either saline or 0.05, 0.10, 0.20 or 0.50 IU of ACTH (Synacthen® depot, Novartis Pharmaceuticals, Auckland) into the jugular vein. Stock Synacthen® depot was diluted in a isotonic saline vehicle. A blood sample was taken at 0900 h (0 min, immediately before the injection) and Chapter Five 145 then further samples were taken 15, 30, 60 and 120 min after the injection. Plasma concentrations of corticosterone were measured in all blood samples. The birds were then rested for at least 10 days before further studies. 5 .2 .2 .2 .2 Relationship between a sUb-maximal dose of ACTH and plasma corticosterone concentrations in low and h igh response quai l Synacthen® at a dose of 0.10 IU was selected as it caused a submaximal corticosterone response as shown in Section 5.2.2.2.1 . Quail in the low and h igh response groups were given a single 100 ,ul injection of saline (n= 6) or 0.10 IU of ACTH (Synacthen® immediate, Novartis Pharmaceuticals, Auckland) in 100 ,ul (n =14) of saline into the jugular vein. A blood sample was taken at 0900 h (0 min, immediately before the injection) and then further samples were taken at 15, 30, 60 and 120 min after the injection. Plasma concentrations of corticosterone were measured in all blood samples. The birds were then rested for at least 10 days before further studies. 5.2.2.3 Relationship between the adrenocortica l response and pitu itary sensitivity to CRF 5.2 .2 . 3 . 1 Relationship between plasma corticosterone and dose of CRF Forty male Japanese quail were randomly selected from the pool of 70 quail and were assigned into five treatment groups (n= 8 per group). The group allocation and known corticosterone response of each bird (as identified in Section 5.2.2.1 ) was checked to avoid placing a large proportion of Iow or high corticosterone response birds into one group. These groups received either a single 100 ,u1 injection of saline or 2, 4, 6 or 8 ,ug of CRF (synthetic ovine corticotrophin releasing factor C-3167, Sigma Chemical) dissolved in 100 ,u1 of saline into the jugular vein. A blood sample was taken at 0900 h (0 min, immediately before the injection) and then further samples were taken at 15, 30, 60 and 120 min after the injection. Plasma concentrations of corticosterone were measured in all blood samples. The birds were then rested for at least 10 days before further studies. 146 Chapter Five 5.2 .2 .3 .2 Relationship between a sub-maximal dose of CRF and plasma corticosterone concentrations in low and high adrenocortical response quai l CRF at a dose of 6 J.Lg was selected as it was the only dose that stimulated a consistent corticosterone response in all birds as shown in Section 5.2.2.3.1 . Quail in the low and high response groups were given a single 100 J.LI injection of saline (n=6) or 6 ;..tg of CRF (synthetic ovine corticotrophin releasing factor C-3167, Sigma Chemical) in 100 ;..tl (n =14) of saline into the jugular vein. A blood sample was taken at 0900 h (0 min, immediately before the injection) and then further samples were taken at 15, 30, 60 and 120 min after the injection. Plasma concentrations of corticosterone were measured in all blood samples. The experiments were conducted under a protocol approved by the Massey University Animal Ethics Committee. 5.2.3 Data col lection 5.2.3.1 Corticosterone admin istration ACTH (Synacthen® depot, Novartis Pharmaceuticals, Auckland) and CRF (synthetic ovine corticotrophin releasing factor C-3167, Sigma Chemical) solutions were prepared on the day of the experiment and then kept on a magnetic stirrer (Chiltern Scientific) at 4°C. Stock solutions were dissolved or mixed in 0.9% saline. ACTH and CRF solutions were administered with 1 ml insulin syringes and needles. 5.2.3.2 Tissue and blood samples Blood samples were collected by venipuncture of the brachial vein in the wing with a 25 g needle, with 200 ;..tl of blood collected into heparinised capillary tubes. All samples were collected within 1 .5 minutes from the time the bird was removed from the cage. The blood was stored on ice until centrifuged at 2 000 g for 15 minutes at 4°C (Heraeus Christ 5000S refrigerated centrifuge). The plasma was removed and stored at -20°C until assayed. Chapter Five 147 5.2.3.3 Corticosterone assay Corticosterone concentrations in quail plasma were measured by radioimmunoassay using an ICN ImmunoChem r .. Double Antibody Corticosterone 1251 RIA kit (ICN Biomedicals, Costa Mesa, CA). Samples were assayed in duplicate in five assays. Plasma was thawed and centifuged in a 1 .5 ml Eppendorf tube at 14 000 g for 5 min (IEC Micromax ventilated microcentrifuge OM3590) to separate clear plasma from lipid within the plasma sample. A 10 ILl aliquot of plasma was incubated with 100 ILl of antibody (anti-corticosterone Cat # 07-120113) and 1 00 ILl of iodinated corticosterone (approximately 5 000 cpm; 125I-corticosterone Cat # 07- 120121) at room temperature (22-25°C) for 2 hours. After incubation, 100 ILl of precipitant solution (Cat # 07-166624) was added to all tubes and mixed using a Chiltern Scientific SS70 orbital shaker for approximately 5 s. Tubes were then centrifuged at 4 500 g for 15 min at 4°C and the supernatant aspirated. The precipitates were counted (3 min per sample) in a LKB Wa11ac 1261 Multigamma Gamma Counter. The limit of sensitivity of the radioimmunoassay, defined as the hormone concentration at the mean minus two standard deviations from the zero hormone point on the standard curve and expressed as corticosterone concentration in plasma, was 0.80 ng/ml. Serial dilutions of quail plasma in diluent buffer were parallel to the corticosterone standard curve (n= 3) . Recovery of 125I-corticosterone added to quail plasma was 90.9 ± 3.8%, 99.2 ± 2.5%, and 90.4 ± 6.6% for three samples. Solutions of corticosterone in PBSG were prepared at concentrations that gave approximately 20, 50, and 80% binding on the standard curve. These solutions were used as high, medium and low quality control samples in every assay. Intra­ assay coefficients of variation have yet to be determined. Inter-assay coefficients of variation in a 11 6 assays were 1 1 .3, 10.5, and 9.9%. The cross-reactions of the antibody with steroids tested by ICN Biomedicals were corticosterone (100.00%), desoxycorticosterone (0.34%), testosterone (0.10%), cortisol (0.05%), aldosterone (0.03%), progesterone (0.02%), androstenedione (0.01 %), 5a-dihydrotestosterone (0.01 %), cholesterol, dehyrdroepiandrosterone, dehyrdroepiandrosterone-sul p ha te, 1 1-desoxycortisol, 20a-dihydroprogesterone, 148 Chapter Five estrone, oestradiol-170', oestradiol-17,6, oestriol, pregnenolone, 170'- hydroxypregnenolone, and 170'-hydroxyprogesterone (all < 0.01 %) . 5.2.4 Calculation of the area under the cort icosterone curve The area under the curve for plasma corticosterone concentrations versus time was calculated for each bird, using the method described in Chapter Two. The ' total integrated response' (Cockrem and Silverin, 2002a) is a measure of the amount of corticosterone secreted during the sampling period and is expressed as nglml·min. Differences in basal corticosterone concentrations between birds were corrected by calculating a second measure called the 'corrected integrated response' by multiplying the corticosterone concentration at Time 0 by the duration of experiment and subtracting this from the total integrated response area. The corrected integrated response was only calculated for the period of the main response between 0 and 30 min, and not for birds where 15 and 30 min corticosterone concentrations fell below initial values. Areas under the curve were calculated using the trapezoid rule using GraphPad Prism 4.0 (GraphPad Software Inc.) . 5.2.5 Statistical analyses All results are presented as the mean ± SE (standard error) unless it was more appropriate to present individual data. All the data were examined for normality using Shapiro-Wilks tests and for equality of variance using Levene's homogeneity of variance test. Hormone data were loglO transformed to avoid heteroscedasticity. If variances were homogenous then groups were compared with repeated measures two-way analyses of variance (ANOV A) with time and treatment as the grouping factors. Comparisons between times within each treatment and between treatments for each time were examined with post-hoc repeated measures contrasts. Data not complying with normality or variance criteria were compared using a Kruskal-Wallis non-parametric test followed by Mann-Whitney U independent group comparisons tests, or Friedman's test for repeat measures data. Chapter Five 149 All comparisons of means were conducting using SPSS TM 1 1 .0 (SPSS Inc., Illinois) or Systat ™ 8.0 (SPSS Inc., Illinois). 5.3 Resu lts 5.3 .1 Identifying low and h igh corticosterone responses to mechanical restraint The adrenocortical response to 15 min of mechanical restraint varied between birds (Figure 5.1a) . Initial corticosterone concentrations ranged from 1 .19 to 8.86 nglml. Thereafter concentrations remained relatively constant in some birds but increased (2.95 - 17.99 nglml) in others after 15 min of mechanical restraint. Corticosterone concentrations had declined by 30 min in most birds, although it remained elevated or increased above baseline values in a few birds. Birds that showed low or high responses at 15 min were identified for a low (n=20) and high response (n =20) groups (Figure 5.1b-c) . There was a significant effect of mechanical restraint on corticosterone concentrations (F2,209 = 37.374, P < 0.001; Figure 5.2a) . Corticosterone concentrations were initially low (5.20 ± 0.20 nglml), increased at 15 min (9.52 ± 0.43 nglml) then decreased 30 min after the initial sample (6.60 ± 0.34 nglml) and remained higher than initial values (post-hoc Bonferroni test, p = 0.007). A separate analysis of birds selected for low and high corticosterone responses showed a significant effect of time (F2,76 = 85.394, P < 0.001 ; Figure 5.2a) and group (F1,38 = 30.499,p < 0.001), and a significant interaction between time and group (F2,76 = 59.555, P < 0.001) . Corticosterone concentrations in low response birds did not change over 30 min. Initial corticosterone concentrations did not differ between low and high birds (F1,38 = 0.767, P = 0.387). Corticosterone concentrations increased in high response birds after mechanical restraint and were significantly higher than low response birds at 15 min (F1 ,38 = 1 19.385, P < 0.001 ), then declined after 15 min and were higher than but did not differ significantly from low response birds at 30 min (F1,38 = 3.958, P = 0.054). 150 Chapter Five Total and corrected integrated corticosterone responses of the high response group were greater than that of the low response group (F1,39 = 55.563, P < 0.001 and F1,27 = 48.656, P < 0.001 respectively; Figure 5.2b). 5.3 .2 Relationship between the adrenocortical response and adrenal sens itivity to ACTH 5.3.2.1 Relationship between plasma corticosterone and dose of ACTH Initial corticosterone concentrations were low in all groups and ranged from 1 .07 - 7.24 ng/ml (Figure 5.3) . Corticosterone concentrations remained low and relatively constant in all birds injected with saline (Figure 5.3a) . There were increases in corticosterone concentrations between 0 and 1 5 min in 50% (4/8) of birds injected with the lowest dose (0.05 IV) of ACTH (Figure 5.3b) . Corticosterone concentrations in these birds had largely declined to baseline by 60 min after the birds were first removed from their cages and remained relatively low at 120 min. Treatment with the next highest dose (0.10 IV) of ACTH stimulated a strong corticosterone response in 75% (6/8) of birds (Figure 5.3c) . Corticosterone concentrations a t 15 min varied between 5.64 and 14.00 ng/ml and then declined to baseline between 15 and 30 min in 88% of birds and remained low from 60 to 120 min. Treatment with 0.20 IV of ACTH stimulated a strong response in all birds, with corticosterone concentrations increasing to 7.49 - 19.37 ng/ml at 15 min, then declining at 30 min and remaining low thereafter (Figure 5.4d). The highest ACTH treatment (0.50 IV) stimulated the largest responses (Figure 5.3e), with corticosterone concentrations increasing in all birds to 1 0.72 - 25.50 ng/ml at 15 min. Corticosterone declined to initial concentrations in all but two birds by 30 min. Corticosterone continued to increase in one bird and was still high in another bird at 30 min, but declined at 60 min in both birds. Corticosterone responses varied between ACTH dose and between time, with a significant interaction between dose and time (see Table 5 .1 for all statistics for this analysis) . Corticosterone concentrations were initially low in all five groups (control- and ACTH-treated) of birds (Figure 5.4a). Mean corticosterone concentrations in birds injected with saline remained low for the duration of the experiment. All doses of ACTH caused mean plasma corticosterone concentrations Chapter Five 151 to increase between 0 and 15 min, with a dose response relationship (Figure 5.4b) . Corticosterone concentrations at 15 min in birds treated with 0.10, 0.20 or 0.50 ill per bird did not differ significantly from each other, but were significantly greater in the 0.05 IU group. Corticosterone declined and did not differ from initial concentrations between 15 and 30 min in all ACTH treated groups, and remained low and relatively constant at 60 and 120 min. Corticosterone at 15 min was no greater for birds treated with 0.50 IU than 0.20 IU, indicating that the maximum corticosterone response to an ACTH injection had been reached. The 0.10 IU dose resulted in a corticosterone response most similar to birds subjected to mechanical restraint. There were significant differences between the groups in the total integrated corticosterone response between the groups (Figure 5.5; see Table 5.2 for all statistics for this analysis) . The total integrated response increased in dose dependent manner, with the responses to 0.20 and 0.50 IUlbird of ACTH significantly larger than in birds injected with saline. The corrected corticosterone response varied between groups, and also increased in a dose dependent manner with a Significant difference between the saline treated and 0.50 IU ACfH groups. 152 Chapter Five 20 All birds (a ) 1 5 1 0 5 0 20 ,..-... Low response (b) E -0) 1 5 C --- Q) c e 1 0 Q) ..... Cl) 0 () 5 t 0 0 0 20 High response (c) 1 5 1 0 5 O�--------------�--------------� o 1 5 30 Time (min) Figure 5.1 Plasma corticosterone in individual male quail immobilised by mechanical restraint for 15 min for (a) all birds (n =70), (b) birds with a low response to mechanical restraint (n=20), and (c) birds with a high response to handling (n=20) . Results are plotted as raw data. Chapter Five 1 5 E 1 0 Dl c Q) c 0 l-Q) - (I) 8 t 5 0 () 0 0 400 E Dl c � Q) (I) c 0 Cl. (I) Q) l- Q) C 0 "- Q) -(I) (.) t 8 "0 Q) 16 "-0) Q) - 0 c � . _ . _ _ __ _ lr . _ . _ _ . _ • • _ - - - - - . � - - . - . - - - - � �� . - - - - - - . _ - - 1 5 30 Time (min) -All birds • «>. Low response - High response 45 1 6 Low High Low High All birds response response All birds response response Group _ Total integrated response c::::::::J Corrected integrated response 153 (a) (b) Figure 5.2 Mean plasma corticosterone concentrations (a) and total and corrected integrated corticosterone responses (b) in male quail immobilised by mechanical restraint for 1 5 min. Total integrated corticosterone responses were calculated from the areas under the corticosterone vs time curves for each bird, while corrected responses were calculated between 0 and 30 min. The number of birds per group are inset inside each column. Results are plotted as means ± standard error. 154 E --O'l C Q) c 0 L-Q) -en 0 () :;::; L-0 0 30 Saline 20 1 0 :;:.�:t:�)·l?'� :';:·;:�:.:,�:�:;;:; �:�: O�.-�--�--��==�====�- 30 20 1 0 0 30 20 1 0 o o 1 5 30 60 0 .05 1 U 0.20 1U 1 20 30 20 o 1 5 30 Time (min) Chapter Five 0 . 1 0 1U 0 .50 1U 60 1 20 Figure 5.3 Plasma corticosterone concentrations in individual male quail that received a single injection of saline or 0.05, 0.10, 0.20, or 0.50 ID ACTH/bird immediately after the 0 min blood sample. Results are plotted as raw data. Chapter Five E --0> C (1) C o .... Q.l - ()') 8 t o o E --0> c c E lO ..-- - co Q) c 0 .... (1) - ()') 8 :e 0 0 20 1 5 1 0 5 0 20 1 5 1 0 5 0 155 .... 0-... Saline (a ) -l:r- 0.05 1U · ·Q.· 0 . 1 0 IU .... - 0 .20 IU - 0.50 IU ::-:.:::: ::-: -. -'-.--.. -. 0 1 5 30 60 1 20 lime (min) (b) I -i I 1/ 1- / ,/i • I 0 0.05 0 .1 0.2 0.5 ACTH dose ( IU per bi rd) Figure 5.4 Mean plasma corticosterone concentrations in male quail that received a single injection of ACfH at a dose of 0.05, 0 .10, 0.20 or 0.50 IU/bird (a); and mean plasma corticosterone concentrations 15 min after an injection of saline or ACTH (b) . Results are plotted as means ± standard error. 156 --c 800 .� E 700 -0> C ID 600 Cl) c 2- 500 Cl) ID L- ID 400 c 0 L-ID ...... Cl) 300 0 u t 200 0 U -0 ID 1 00 ...... ro L-0> ID ...... 0 c Sal ine 0.05 Chapter Five 0. 10 0 .20 0 .50 Sal ine 0.05 0 . 1 0 0.20 0 .50 ACTH dose ( I U per bird) - Total integrated response c:::::::J Corrected integrated response Figure 5.5 Total and corrected corticosterone responses in quail treated with saline or ACTH (Synacthen TM ) at doses of 0.05, 0.10, 0.20 or 0.50 IU/bird. Total integrated corticosterone responses were calculated from the areas under the corticosterone vs time curves for each bird, while corrected responses were calculated between 0 and 30 min. The number of birds per group are inset inside each column. Results are plotted as means ± standard error. Chapter Five 157 Table 5.1 Two-way repeated measures ANOVA for plasma corticosterone concentrations in male quail treated with ACTH. Plasma corticosterone Effect F df p Dose 4,340 4,35 0.006 •• Time 41.144 4,140 < 0.00] •• Interaction of dose and time 2.1 13 ]6,140 0.012 • Comparisons within dose for each dose o vs 15 min Saline 1 .184 1,35 0.284 0.05 1U 2.400 1,35 0.130 O.10 IU 9.093 ] ,35 0.005 •• 0.20 IU ]7.032 ] ,35 < 0.001 • • 0.50 IU 25.736 1,35 < 0.001 • • 15 vs 30 min Saline 0.285 1 ,35 0.597 0.05 IU 20.553 ],35 < 0.001 • • 0.10 IU 8 .]21 ] ,35 0.007 • • 0.20 IU 27.690 ] ,35 < 0.001 • • 0.50 IU 24.708 1 ,35 < 0.001 • • 30 v s 60 min Saline 2.037 ] ,35 0.162 0.05 IU 0.787 1,35 0.381 0.10 IU 3.001 1 ,35 0.092 0.20 IU 1 .487 1,35 0.231 0.50 IU 1 3.691 1 ,35 0.001 • • 60 vs 120 min Saline 8.445 1 ,35 0.006 • • 0.05 IU 2.979 1,35 0.093 0.10 IU 6.882 1,35 0.01 3 • 0.20 IU 5.207 1 ,35 0.029 • 0.50 1U 0.086 1 ,35 0.772 o vs 30 min Saline 0.301 1 ,35 0.587 0.05 IU 1 .919 1,35 0.175 O.10 IU 0.539 1 ,35 0.468 0.20 1U 0.050 1 ,35 0.825 0.50 IU 1 .286 1 ,35 0.264 o vs 60 min Saline 2.122 1 ,35 0.1 54 0.05 IU 0.584 1 ,35 0.450 0.10 IU 0.171 1 ,35 0.682 O.20 IU 0.331 1 ,35 0.569 0.50 IU 1 .700 ] ,35 0.201 o vs 120 min Saline 20.291 1 ,35 < 0.001 •• 0.05 IU 6.582 1 ,35 0.015 • 0.10 1U 9.515 1 ,35 0.004 •• 0.20 1U 8.516 ],35 0.006 •• 0.50 1U 2.937 ] ,35 0.095 Note. The first three rows show the results of the two-way repeated measures ANOVA for the main effects and their interaction. The remaining rows show the results of post-hoc comparisons of times for each trea tment and comparisons of treatments a t each time. Significant differences (p < 0.05) are marked with a single asterisk, while highly significant d ifferences (p < 0.01) are marked with a double asterisk. 158 Chapter Five Table 5.1 cont Two-way repeated measures ANOVA for plasma corticosterone in male quail treated with ACTH. Comparisons between doses for each time F df P O min Saline vs 0.05 IU 0.114 1 ,35 0.738 Saline vs 0.10 IU 0.203 1 ,35 0.655 Saline vs 0.20 IU 0.349 1,35 0.559 Saline vs 0.50 IU 0.013 1,35 0.911 0.05 vs 0.10 IU 0.620 1 ,35 0.436 0.05 vs 0.20 IU 0.862 1,35 0.360 0.05 vs 0.50 IU 0.203 1,35 0.655 0.10 vs 0.20 IU 0.020 1 ,35 0.889 0.10 vs 0.50 IU 0.1 14 1,35 0.738 0.20 vs 0.50 IU 0.229 1 ,35 0.636 15 min Saline vs 0.05 IU 5.741 1,35 0.022 • Saline vs 0.10 IU 20.823 1 ,35 < 0.001 • • Saline vs 0.20 IU 33.820 1,35 < 0.001 • • Saline v s 0.50 IU 40.568 1,35 < 0.001 • • 0.05 v s 0.10 IU 4.697 1,35 0.037 • 0.05 vs 0.20 IU 11 .693 1,35 0.002 • • 0.05 vs 0.50 IU 15.787 1,35 < 0.001 • • 0.10 v s 0.20 IU 1 .598 1,35 0.219 0.10 vs 0.50 IU 3.262 1 ,35 0.080 0.20 vs 0.50 IU 0.307 1,35 0.583 30 min Saline vs 0.05 IU 1 .1 33 1 ,35 0.294 Saline vs 0.10 IU 2.521 1,35 0.121 Saline vs 0.20 IU 1 .363 1 ,35 0.251 Saline vs 0.50 IU 3.115 1 ,35 0.086 0.05 vs 0.10 IU 7.034 1,35 0.012 • 0.05 vs 0.20 IU 4.982 1 ,35 0.032 0.05 vs 0.50 IU 8.007 1 ,35 0.008 • • 0.10 vs 0.20 I U 0.177 1,35 0.677 0.10 vs 0.50 IU 0.031 1,35 0.860 0.20 vs 0.50 IU 0.357 1,35 0.554 60 min Saline vs 0.05 IU 0.181 1,35 0.673 Saline vs 0.10 IU 1 .414 1,35 0.242 Saline vs 0.20 IU 1 .265 1,35 0.268 Saline vs 0.50 IU 0.040 1,35 0.842 0.05 vs 0.10 IU 0.583 1,35 0.450 0.05 vs 0.20 IU 0.489 1 ,35 0.489 0.05 vs 0.50 IU 0.051 1 ,35 0.823 0.10 vs 0.20 IU 0.004 1,35 0.949 0.10 vs 0.50 IU 0.977 1,35 0.330 0.20 vs 0.50 IU 0.854 1 ,35 0.362 120 min Saline vs 0.05 IU 1.917 1,35 0.175 Saline vs 0.10 IU 1 .984 1,35 0.168 Saline vs 0.20 IU 2.636 1,35 0.113 Saline vs 0.50 IU 5.437 1,35 0.026 • 0.05 vs 0.10 IU 0.001 1,35 0.981 0.05 vs 0.20 IU 0.057 1,35 0.813 0.05 vs 0.50 IU 0.897 1,35 0.350 0.10 vs 0.20 IU 0.046 1 ,35 0.831 0.10 vs 0.50 IU 0.852 1,35 0.362 0.20 vs 0.50 IU 0.502 1,35 0.483 Chapter Five 159 Table 5.2 One-way ANOVA for the integrated corticosterone responses in quail treated with ACTH. Comparisons between doses F df p Integrated corticosterone response Total 4.087 4,35 0.008 • • Corrected 3.715 4,17 0.024 • Bonferroni post-hoc comparisons between groups Total response Corrected response df p df p Saline vs 0.05 IU 1 ,35 1 .000 1 ,17 1 .000 0.10 IU 1 ,35 0.289 1 ,17 0.918 0.20 IU 1 ,35 0.029 • 1 ,17 0.588 0.50 IU 1 ,35 0.034 • 1 ,17 0.021 • 0.05 vs 0.10 IU 1 ,35 1 .000 1 ,17 1 .000 0.20 IU 1 ,35 0 .222 1 ,17 1 .000 0.50 IV 1 ,35 0.259 1,17 0.177 0.10 vs 0.20 IV 1 ,35 1 .000 1,17 1 .000 0.50 IV 1 ,35 1 .000 1 ,17 0.685 0.20 vs 0.50 IV 1 ,35 1 .000 1,17 1 .000 5.3.2.2 Relationship between a sub-maximal dose (O. 1 0 IU/bird) of ACTH and plasma corticosterone concentrations in low and high response quai l Corticosterone concentrations were low in four of six birds in both the low and high corticosterone response groups injected with saline, and remained relatively constant throughout the period of sampling in these birds (Figure 5.6) . The initial corticosterone concentration was elevated (8.33 - 1 0.65 nglml) in two birds in each group. Corticosterone remained relatively unchanged or declined after the saline injection and did not differ between groups at 60 min. Initial corticosterone concentrations varied substantially in both the low and high groups before ACTH injection, ranging from 1 . 38 - 1 1 .62 nglml (Figure 5 .6) . The administration of ACTH resulted in similar increases in corticosterone in both the low and high response groups. Corticosterone concentrations increased between 0 and 15 min, declined between 15 and 30 min, then generally continued to decline and remained low and relatively constant between 30 and 60 min. Corticosterone varied between time, but not between groups, with a significant 160 Chapter Five interaction between group and time (see Table 5.3 for all statistics for this analysis). Corticosterone concentrations at 0 min did not differ between saline and ACTH treated birds, or low and high response groups (Figure 5.7a) . There was no significant change in plasma corticosterone concentrations in the low and high response birds injected with saline during the sampling period. Corticosterone concentrations in low and high response birds injected with 0. 10 IUlbird ACTH increased to peak values by 15 min (1 1 .83 ± 3.05 nglml and 1 1 .45 ± 3 .06 nglml respectively), declined between 15 and 60 min then remained low for the remainder of the experiment (Figure 5.7a) . Concentrations at 30 min were greater than initial concentrations, but generally did not differ from initial corticosterone concentrations at 60 and 120 min. Corticosterone did not differ at any time between the low and high response birds that had been treated with ACTH. There were Significant differences in the total integrated corticosterone response between groups (Table 5.4; Figure 5.7b) . The corrected integrated response (0-30 min) was significantly greater for ACTH than saline treated low or high response birds. Chapter Five Low response birds 20 Saline ACTH 15 i ': :::�:':;2�:::��;��.i'i 0) � O�--.--.------'------------r�r--.--.------.------------. Q) c e Q) ...... Cl) 8 20 t o 1 5 () o 1 5 30 60 1 20 0 1 5 30 60 120 High response birds Saline O�--.--.------'------------r�.--.--.------.------------. o 15 30 60 1 20 0 15 30 60 1 20 Time (min) 161 Figure 5.6 Plasma corticosterone in individual male quail with low or high responses to mechanical restraint. Birds were injected with saline (n=6) or 0.10 IU ACTH/bird (Synacthen H'; n= 14). Results are plotted as raw data. 1 62 1 5 E Cl 1 0 c: Q.) c: e Q.) ..... (f) 8 :;::; 5 ... o o .-.. c: .� E -... Cl c: Q) (J) c: 0 0. (J) Q) .... Q) c: a .... Q) .-(J) a (.) :;::; L-a (.) '"0 Q.) m .... Cl Q.) .-c: 0 1000 7 500 0 ( a ) 0 1 5 (b) Low High Saline 30 60 " , "i'. � Low response (Saline) -v- low response (ACTH) ..... High response (Saline) -.. - High response (ACT H) TIme (min) 4 Low High Low High low High ACTH Saline ACTH Group -Total integrated response c=:J Corrected integrated response Chapter Five 1 20 Figure 5.7 Mean plasma corticosterone concentrations (a); and the total and corrected integrated corticosterone responses (b) in male quail selected for low or high responses to mechanical restraint that were treated with saline (n=6) or 0.10 IU ACTH/bird (Synacthen H'; n=14). Total integrated corticosterone responses were calculated from the areas under the corticosterone vs time curves for each bird, while corrected responses were calculated between 0 and 30 min. The number of birds per group are inset inside each column Results are plotted as means ± standard error. Chapter Five 163 Table 5.3 Two-way repeated measures ANOVA for corticosterone concentrations in low and high response quail treated with 0.10 ID ACTHlbird. Plasma corticosterone Effect F df p Group 1 .502 3,36 0.230 Time 21 .234 4,144 < 0.001 •• Interaction of group and time 2.752 12,144 0.003 •• Comparisons within group for each group o vs 1 5 mm Low (saline) 0.405 1 ,36 0.528 Low (ACTH) 45.480 1 ,36 < 0.001 • • High (saline) 1 .1 25 1 ,36 0.296 High (ACTH) 36.596 1,36 < 0.001 • • 15 v s 30 nun Low (saline) 0.002 1 ,36 0.967 Low (ACTH) 28.040 1 ,36 < 0.001 • • High (saline) 0.971 1 ,36 0.331 High (ACTH) 33.398 1 ,36 < 0.001 • • 30 vs 60 mm Low (saline) 0.127 1 ,36 0.724 Low (ACTH) 16.046 1 ,36 < 0.001 •• High (saline) 2.304 1,36 0.138 High (ACTH) 12.204 1 ,36 0.001 • • 60 v s 120 min Low (saline) 3.079 1 ,36 0.088 Low (ACTH) 6.470 1 ,36 0.015 • High (saline) 1 .320 1 ,36 0.258 High (ACTH) 0.180 1,36 0.674 o vs 30 mm Low (saline) 0.373 1 ,36 0.545 Low (ACTH) 13.616 1 ,36 0.001 - - High (saline) 2.626 1 ,36 0.1 14 High (ACTH) 7.406 1 ,36 0.010 • o vs 60 min Low (saline) 0.486 1,36 0.490 Low (ACTH) 0.040 1 ,36 0.842 High (saline) 0.007 1 ,36 0.933 High (ACTH) 0.285 1 ,36 0.596 o vs 120 mm Low (saline) 0.639 1 ,36 0.429 Low (ACTH) 6.165 1 ,36 0.018 • High (saline) 0.864 1 ,36 0.359 High (ACTH) 0.925 1,36 0.343 Note. The first three rows show the re ults of the two-way repeated measures ANOVA for the mam effects and their interaction. The remaining rows show the results of post-hoc comparisons of times for each treatment and comparisons of treatments at each time. Significant differences (p < 0.05) are marked with a single asterisk, while highly significant d ifferences (p < 0.01 ) are marked with a double asterisk. 164 Chapter Five Table 5.3 cont Two-way repeated measures ANOVA for corticosterone concentrations in low and high response quail treated with 0.10 IU ACTHlbird. Comparisons between groups for each time F df P O min Low (saline) vs Low (ACTH) 0.038 1 ,36 0.847 Low (saline) vs High (saline) 0.121 1,36 0.730 Low (ACTH) vs High (ACTH) 0.041 1,36 0.841 High (saline) vs High (ACTH) 0.581 1,36 0.451 15 min Low (saline) vs Low (ACTH) 14.486 1,36 0.001 * * Low (saline) vs High (saline) 0.028 1,36 0.868 Low (ACTH) vs High (ACTH) 0.065 1 ,36 0.800 High (saline) vs High (ACTH) 14.486 1 ,36 0.001 * * 30 min Low (saline) vs Low (ACTH) 4.714 1,36 0.037 * Low (saline) vs High (saline) 0.107 1 ,36 0.745 Low (ACTH) vs High (ACTH) 0.274 1,36 0.604 High (saline) vs High (ACTH) 1 .899 1,36 0.177 60 min Low (saline) vs Low (ACTH) 0.314 1 ,36 0.579 Low (saline) vs High (saline) 0.687 1,36 0.413 Low (ACTH) vs H igh (ACTH) 0.003 1,36 0.959 High (saline) vs High (ACTH) 0.145 1 ,36 0.706 120 min Low (saline) vs Low (ACTH) 0.573 1,36 0.454 Low (saline) vs H igh (saline) 0.470 1 ,36 0.497 Low (ACTH) vs H igh (ACTH) 3.871 1 ,36 0.057 High (saline) vs High (ACTH) 2.492 1 ,36 0.123 Table 5.4 One-way ANOV A for the integrated corticosterone responses in low and high response quail treated with 0.10 IU ACTH/bird. Comparisons between doses Integrated corticosterone response Total Corrected Bonferroni post-hoc comparisons between groups Low (saline) vs Low (ACTH) High (saline) Low (ACTH) vs High (ACTH) High (saline) vs High (ACTH) F 1 .383 13.879 Total response df p 1 ,36 1 .000 1 ,36 1 .000 1 ,36 1 .000 1 ,36 0.442 df 3,36 3,29 p 0.264 < 0.001 * * Corrected response df p 1 ,29 < 0.001 * * 1 ,29 0.572 1 ,29 1 .000 1 ,29 0.013 * Chapter Five 5.3.3 Relationship between the adrenocortical response and p itu itary sensitivity to CRF 165 5.3.3 . 1 Relationship between plasma corticosterone and dose of CRF Initial corticosterone concentrations varied between birds and ranged from 0.76 - 8.44 nglml (Figure 5.8). Corticosterone concentrations remained relatively constant in all but one of the saline treated birds during the study. Corticosterone concentrations increased between 0 and 15 min in birds injected with 2 /-Lg CRFlbird, although the response varied and some birds did not respond to treatment (Figure 5.8b). Corticosterone concentrations generally declined between ]5 and 30 min and remained low and relatively constant for the rest of the treatment. There was no consistent pattern to corticosterone concentrations in birds injected with 4 /-Lg CRF/bird. Two birds showed large increases in corticosterone concentrations between 0 and 15 min while there was little change in the other six birds after treatment (Figure 5.8c). Treatment with 6 /-Lg CRFlbird resulted in large increases in corticosterone to 6.60 - 14.24 nglml in 75% of birds at 15 min. Corticosterone increased to a lesser extent in the remaining birds (Figure 5.8d). Corticosterone declined to initial concentrations in 75% of birds between 15 and 30 min and remained low, while it remained elevated in two birds at 60 and 120 min. Treatment with 8 /-Lg CRFlbird did not result in a clear increase in corticosterone concentration (Figure 5.8e), with increases in three birds but and little change in the other five birds. Corticosterone concentrations varied between time but not between doses, with a significant interaction between dose and time (see Table 5.5 for all statistics for this analysis). Initial corticosterone concentrations were low and did not differ between the five groups (Figure 5.9a). Mean corticosterone concentrations in birds injected with saline remained low for the duration of the experiment, and the 2, 4, and 8 fLg groups did not differ from the saline group at any of the sampling times. Treatment with 6 fLg CRF was the only treatment that stimulated a significant corticosterone response (Figure 5.9a). Treatment with 6 /-Lg CRFlbird induced a significant increase in mean plasma corticosterone concentrations between 0 and 15 min. Corticosterone concentrations at 15 min were 10.74 ± 3.80 nglml and were significantly higher than any of the other groups, then declined between 15 and 166 Cha pter Five 30 min and were no different from the other groups for the remainder of the study. The 6 J-Lg CRF dose resulted in a net increase in corticosterone concentrations of approximately 6 nglml; an increase similar to that caused by 0.10 IU of ACTH or mechanical restraint. There were no differences between groups in the total or corrected integrated corticosterone response between the groups (Figure 5.9b; see Table 5.6 for all statistics for this analysis). Chapter Five 1 5 (a ) Saline 1 0 5 O�.--.--.-----.-----------�- E 1 5 (b) -0) C ----Q) 1 0 C e Q) - en 8 t o () 5 O��-.--�----.-----------�- 1 5 6 �g 1 0 5 O�.-�--.-----.-----------�- o 1 5 30 60 120 1 5 (c) 4 �g o���--�----�----------�- 1 5 (e) 8 �g 10 T 5 t�����:.,��:��;,,;.�:����� O�.--.--.-----�----------�- o 1 5 30 60 1 20 Time (m in) 167 Figure 5.8 Plasma corticosterone concentrations in individual male quail that received a single injection of saline or 2, 4, 6, or 8 JLg CRF/bird (n=8) immediately after the 0 min blood sample. Results are plotted as raw data . 168 1 5 E en 10 c:: a> c:: o .... a> Vi o t) t o u 5 (a) - Saline · ·';· ' 2 119 -" - 4 119 - 6 119 ··· .. ··· 8 119 ...... :.,. �-.:.::.:. ::.::.,-,::_ •....... _ .. -•.....• --------- . _ - - - -. ... _ - ... ... _- ... ... ... .. _- Chapter Five o�._--_.----.---------_.--------------------� -c:: 800 E E en c:: -; 600 f/) c:: o 0. f/) a> .... a> c:: o .... a> (5 l3 t o t) "0 a> ro .... Cl a> C 400 200 o o 15 Saline 2 30 4 6 60 Time (min) 8 Saline 2 CRF dose ( fl9 per bird) _ Total integrated response 4 [==:J Corrected integrated response 1 20 6 8 Figure 5.9 Mean plasma corticosterone concentrations (a); and the total and corrected integrated corticosterone responses (b) in male quail treated with saline (n=8) or 2, 4, 6, or 8 J.Lg CRF/bird (n=8). Total integrated corticosterone responses were calculated from the areas under the corticosterone vs time curves for each bird, while corrected responses were calculated between 0 and 30 min. The number of birds per group are inset inside each column. Results are plotted as means ± standard error. Chapter Five 169 Table 5.5 Two-way repeated measures ANOVA for corticosterone concentrations in male quail treated with CRF. Plasma corticosterone Effect F df p Dose 1 .292 4,35 0.292 Time 34.915 4,140 < 0.001 * * Interaction of dose and time 3.266 16,140 < 0.001 * * Comparisons within dose for each dose o vs 1 5 min Saline 0.377 1 ,35 0.543 2p.g 15.803 1,35 < 0.001 * * 4 p.g 4.858 1,35 0.034 * 6p.g 63.129 1,35 < 0.001 * * 8 p.g 4.940 1 ,35 0.033 * 15 vs 30 min Saline 1 .443 1,35 0.238 2 p.g 12.945 1,35 0.001 * * 4p.g 2.268 1 ,35 0.141 6 p.g 43.489 1 ,35 < 0.001 * * 8 p.g 1 .528 1,35 0.225 30 vs 60 min Saline 1 .102 1 ,35 0.301 2 p.g 6.070 1,35 0.019 * 4p.g 2.037 1 ,35 0.162 6 p.g 0.174 1 ,35 0.679 8 p.g 3.921 1 ,35 0.056 60 vs 1 20 mm Saline 0.412 1 ,35 0.525 2p.g 2.330 1 ,35 0.136 4 p.g 2.599 1 ,35 0.1 16 6 p.g 1 .853 1 ,35 0.182 8 p.g 0.258 1,35 0.615 o vs 30 mm Saline 0.347 1 ,35 0.560 2 p.g 0.486 1 ,35 0.490 4 p.g 0.835 1 ,35 0.367 6 p.g 4.419 1 ,35 0.049 * 8 p.g 1 .504 1 ,35 0.228 o vs 60 min Saline 1.735 1,35 0.196 2 p.g 1 .068 1,35 0.309 4 p.g 0.009 1 ,35 0.926 6 p.g 5.328 1 ,35 0.027 * 8 p.g 0.029 1 ,35 0.865 o vs 1 20 mm Saline 3.910 1 ,35 0.056 2 p.g 6.011 1 ,35 0.019 * 4 p.g 2.309 1 ,35 0.138 6 p.g 1 .630 1,35 0.210 8 p.g 0.398 1,35 0.532 Note. The first three rows show the results of the two-way repeated measures ANOV A for the main effects and their interaction. The remaining rows show the results of post-hoc comparisons of times for each treatment and comparisons of treatments at each time. Significant differences (p < 0.05) are marked with a single asterisk, while highJy significant differences (p < 0.01) are marked with a double asterisk. 170 Chapter Five Table 5.5 cont Two-way repeated measures ANOVA for corticos terone concentrations in male quail treated with CRF. Comrarisons between doses for each time F dt l!. O min Saline v s 2 J.Lg 0.1 1 8 1 ,35 0.733 Saline vs 4 J.Lg 0.092 1 ,35 0.763 Saline vs 6 J.Lg 0.751 1,35 0.392 Saline vs 8 J.Lg 0.035 1 ,35 0.853 2 vs 4 J.Lg 0.002 1 ,35 0.969 2 vs 6 J.Lg 0.274 1 ,35 0.604 2 vs 8 J.Lg 0.025 1 ,35 0.876 4 vs 6 J.Lg 0.317 1,35 0.577 4 vs 8 J.Lg 0.014 1 ,35 0.907 6 vs 8 J.Lg 0.463 1 ,35 0.501 15 min Saline vs 2 J.Lg 3.841 1 ,35 0.058 Saline vs 4 J.Lg 0.678 1 ,35 0.416 Saline vs 6 J.Lg 17.540 1,35 < 0.001 •• Saline vs 8 J.Lg 0.850 1 ,35 0.363 2 vs 4 J.Lg 1 .291 1 ,35 0.264 2 vs 6 J.Lg 4.965 1,35 0.032 • 2 vs 8 J.lg 1.077 1 ,35 0.307 4 vs 6 J.lg 11 .320 1,35 0.002 • • 4 v s 8 J.Lg 0.010 1 ,35 0.922 6 vs 8 J.Lg 10.666 1 ,35 0.002 ** 30 min Saline vs 2 J.lg 0.288 1,35 0.595 Saline vs 4 J.lg 0.500 1 ,35 0.484 Saline vs 6 J.Lg 0.925 1 ,35 0.343 Saline vs 8 J.lg 1 .003 1 ,35 0.323 2 vs 4 J.Lg 0.029 1,35 0.866 2 vs 6 J.Lg 0.180 1,35 0.674 2 vs 8 J.lg 0.216 1,35 0.645 4 vs 6 J.Lg 0.065 1 ,35 0.801 4 vs 8 J.Lg 0.087 1 ,35 0.770 6 vs 8 J.Lg 0.002 1,35 0.968 60 min Saline vs 2 J.Lg 0.013 1,35 0.909 Saline vs 4 J.Lg 0.401 1,35 0.531 Saline vs 6 J.lg 3.645 1 ,35 0.064 Saline vs 8 J.Lg 0.473 1,35 0.496 2 vs 4 J.lg 0.560 1,35 0.459 2 vs 6 J.lg 4.098 1,35 0.051 2 vs 8 J.lg 0.645 1,35 0.427 4 vs 6 J.lg 1 .628 1 ,35 0.210 4 vs 8 J.Lg 0.003 1,35 0.957 6 vs 8 J.lg 1 .491 1,35 0.230 120 min Saline vs 2 J.lg 0.462 1,35 0.501 Saline vs 4 J.lg 0.001 1,35 0.975 Saline vs 6 J.lg 2.237 1,35 0.144 Saline vs 8 J.lg 0.622 1 ,35 0.436 2 vs 4 J.lg 0.506 1,35 0.482 2 vs 6 J.Lg 4.731 1,35 0.036 .. 2 vs 8 J.Lg 2.155 1,35 0.151 4 vs 6 J.Lg 2.143 1,35 0.152 4 vs 8 J.lg 0.573 1 ,35 0.454 6 vs 8 J.lg 0.500 1 ,35 0.484 Cha pter Five 171 Table 5.6 One-way ANOVA for the integrated corticosterone responses in quail trea ted with CRF. Comparisons between doses F df p Integrated corticosterone response Total 1 .480 4,35 0.229 Corrected 3.757 4,1 9 0.020 * Bonferroni post-hoc comparisons between grou ps Total response Corrected response df p df P Saline vs 2J.Lg 1 ,35 1 .000 1 ,19 1 .000 4 J.Lg 1,35 1 .000 1,1 9 1 .000 6 J.Lg 1 ,35 0.349 1 , 19 0.569 B J.Lg 1,35 1 .000 1 ,19 1 .000 2 vs 4 J.Lg 1,35 1 .000 1,19 0 .447 6 J.Lg 1 ,35 0.585 1,19 1 .000 8 J.Lg 1 ,35 1 .000 1 ,19 0.352 4 vs 6 J.Lg 1,35 1 .000 1 , 19 0.079 8 J.Lg 1 ,35 1 .000 1,19 1 .000 6 vs B J.Lg 1,35 1 .000 1 , 19 0.053 5.3.3 .2 Relationship between a sub-maximal dose of CRF and plasma corticosterone concentrations in low and high response quail Initial corticosterone concentrations across al l four groups (saline- vs. CRF-treated low and high corticosterone responders to 15 min of mechanical restraint) ranged between 1 .04 and 10.93 nglml (Figure 5.10). Corticosterone concentrations fluctuated and changed up to 3-4 fold in low and high response groups injected with saline, and did not differ between any of the, groups at 0 min (Figure �.l1a) . Administration of CRF did not affect corticosterone concentrations in low or high response groups and there was no consistent temporal pattern to secretion in either group. Corticosterone varied between time but not between treatment, with a significant interaction between treatment and time (Figure 5.11a; see Table 5.7 for statistical analysis). There was no corticosterone response to the administration of CRF in either low or high response groups. Corticosterone in saline and CRF treated low response groups did not change significantly during the sampling period. Corticosterone in saline and CRF-treated high response groups remained relatively constant between 0 and 30 min, then gradually declined between 30 and 120 min to be lower than initial concentrations at 120 min. There were no differences in either total or corrected integrated corticosterone response between the groups (Table 5.8; Figure 5.11b). 172 Low response b irds 12 12 Saline 1 0 10 8 8 ......... 6 Q ••••• .D •••••• • ········.. .. • •••••.•.•.••••••••• , ........ '" ,' . . ...... : .:.r; . . a': .0, •• >::.� .. .a-.-.-....... - - 6 ,-.... E 4 - 0> 2 c: -- ... �. .. D _ •• • - .·····0 •. _ •.• _ •.. :: •••• ::.:: .. �. -'0 4 >.\.:::"'�- ... ::·i':····---····: ::::::::;:::::a_·:::::�::··_.·.·_ ........ �.·.·:.� ..... ·:�:::.:��:::::::=::::8 ..:\�. '0- ... < .... __ .............. _ .. . .0 ••• _........... 2 ·· h ···· ···········0 Chapter Five Q) O-'-r--,------,---,------------, c: O-'-r--,----.---�------� e Q) ...... Cl) 8 1 2 t 1 0 o o D High response b irds 12 Saline 1 0 8 8 · �::::�,��:; ;;;,�:;��::�::;�i;;��Z� : CRF o O-'-r--.--,,---�------� o 1 5 30 60 1 20 0 15 30 Time (m in) 60 1 20 Figure 5.10 Plasma corticosterone in individual male quail selected for low or high responses to mechanical restraint. Birds were injected with saline (n =6) or 6 J.Lg of CRF (n =14). Results are plotted as raw data. Chapter Five E --Cl C Q) C o L-Q) ..... (f) 8 � L-o U ,-.. c E E --Cl c Q) (f) c 0 c.. (f) Q) L- Q) c 0 L-Q) ..... (f) 8 � L-0 u "0 Q) ..... ro L-Cl Q) ..... c 8 6 4 2 0 600 400 200 50 25 0 (a) - -T- - Low response (Saline) -'f'- Low response (CRF) - -.... . . High response (Saline) I -... - High response (CRF) ,,----"' -! ! I ' I It-------� 1:: "" "it:: ::-:: lit?��1;- " " " " " " " " " " " " "" " " " " " " " t ! ''f'-- I .. . . . . .. I�I - . - . - • • • " . . .. ...... . . I ' " I " " 1 ... ..... ... - .. ... .. .. .. ... .. ' " · · · · · · - · · · ·i I 0 1 5 30 60 1 20 Time (min) (b) 3 1 0 9 Low High Low High Low High Low High Saline CRF Saline CRF Group _ Total i ntegrated response c=J Corrected i ntegrated response 173 Figure 5.11 Mean plasma corticosterone concentrations (a); and the total and corrected integrated corticosterone responses (b) in male quail selected for low or high responses to mechanical restraint that were treated with saline (n=6) or 6 /-Lg CRF/bird (n= 14). Total integrated corticosterone responses were calculated from the areas under the corticosterone vs time curves for each bird, while corrected responses were calculated between 0 and 30 min. The number of birds per group are inset inside each column. Results are plotted as means ± standard error. 1 74 Chapter Five Table 5.7 Two-way repeated measures ANOVA for corticosterone concentrations in low and high response quail treated with 6 J.Lg CRF/bird. Plasma corticosterone Effect F df p Treatment 1 .3 17 3,36 0 .284 Time 12.855 4,144 < 0.001 • • Interaction o f treatment and time 1 .853 12,144 0.045 • Comparisons within treatment for each treatment O vs 15 min Low (saline) 0.956 1 ,36 0.335 Low (CRF) 1 .670 1 ,36 0.204 High (saline) 0.090 1 ,36 0.765 High (CRF) 0.018 1 ,36 0.893 15 vs 30 min Low (saline) 1 .429 1,36 0.240 Low (CRF) 0.612 1 ,36 0.439 High (saline) 2.916 1 ,36 0.096 High (CRF) 0.005 1 ,36 0.945 30 vs 60 min Low (saline) 0.071 1 ,36 0.79] Low (CRF) 2.703 1 ,36 0.109 High (saline) 2.420 1 ,36 0.129 High (CRF) 8.552 1 ,36 0.006 • • 60 vs 120 min Low (saline) 0.677 1 ,36 0.416 Low (CRF) 1 .438 1 ,36 0.238 High (saline) 3.807 1 ,36 0.059 High (CRF) 9.895 1 ,36 0.003 >t . o vs 30 min Low (saline) 0.004 1 ,36 0.950 Low (CRF) 0.258 1 ,36 0.615 High (saline) 1 .026 1 ,36 0.318 High (CRF) 0.004 1 ,36 0.951 o vs 60 min Low (saline) 0.044 1 ,36 0.835 Low (CRF) 0.303 1 ,36 0.585 High (saline) 3 .100 ] ,36 0.087 High (CRF) 2.814 ] ,36 0.102 o vs 120 min Low (saline) 0.357 1 ,36 0.554 Low (CRF) 2.953 1 ,36 0.094 High (saline) 1 3 .336 1 ,36 0.001 •• High (CRF) 22.456 1 ,36 < 0.001 • • Note. The first three rows show the results o f the two-way repeated measures ANOVA for the main effects and their interaction. The remaining rows show the results of post-hoc comparisons of times for each treatment and comparisons of treatments at each time. Significan t differences (p < 0.05) are marked with a single asterisk, while highly significant d ifferences (p < 0.01 ) are marked with a double asterisk. Chapter Five 175 Table 5.7 cont Two-way repeated measures ANOVA for corticosterone concentrations in low and high response quail treated with 6 f..Lg CRF/bird. Comparisons between doses for each time F df p O min Low (saline) vs Low (CRF) 0.656 1,36 0.423 Low (saline) vs High (saline) 0.263 1,36 0.61 1 Low (saline) vs High (CRF) 0.754 1,36 0.391 Low (CRF) vs High (saline) 0.041 1 ,36 0.840 Low (CRF) vs High (CRF) 0.006 1,36 0.940 High (saline) vs High (CRF) 0.069 1,36 0.795 15 min Low (saline) vs Low (CRF) 6.106 1,36 0.018 • Low (saline) vs High (saline) 2.248 1 ,36 0.143 Low (saline) vs High (CRF) 3.507 ] ,36 0.069 Low (CRF) vs High (saline) 0.486 1,36 0.490 Low (CRF) vs High (CRF) 0.597 1,36 0.445 High (saline) vs High (CRF) 0.010 1 ,36 0.922 30 min Low (saline) vs Low (CRF) 1 .288 1 ,36 0.264 Low (saline) vs High (saline) 0.145 1,36 0.705 Low (saline) vs High (CRF) 0.810 1,36 0.374 Low (CRF) vs High (saline) 2.516 1 ,36 0.121 Low (CRF) vs High (CRF) 0.092 1,36 0.764 High (saline) vs High (CRF) 1 .826 1 ,36 0.185 60 min Low (saline) vs Low (CRF) 0.018 1 ,36 0.894 Low (saline) vs High (saline) 1 .548 1,36 0.221 Low (saline) vs High (CRF) 0.320 1,36 0.575 Low (CRF) vs High (saline) 2.580 1,36 0.117 Low (CRF) vs High (CRF) 0.816 1 ,36 0.372 High (saline) vs High (CRF) 0.821 1,36 0.371 120 min Low (saline) vs Low (CRF) 0.020 1,36 0.889 Low (saline) vs High (saline) 3.245 1,36 0.080 Low (saline) vs High (CRF) 2.191 1,36 0.148 Low (CRF) vs High (saline) 5.162 1,36 0.029 • Low (CRF) vs High (CRF) 4.378 1,36 0.044 • High (saline) vs High (CRF) 0.424 1,36 0.519 176 Chapter Five Table 5.8 One-way ANOVA for the integrated corticosterone responses in low and high response quail treated with 6 f-Lg CRF. Comparisons between doses F df p Integrated corticosterone response Total 1 .678 3,36 0.189 Corrected 0.809 3,21 0.503 Bonferroni post-hoc comparisons between groups Total response Corrected response df p df P Low (saline) vs Low (CRF) 1 ,36 1 .000 1,21 1 .000 High (saline) 1 ,36 1 .000 1 ,21 1 .000 High (CRF) 1 ,36 1 .000 1 ,21 1 .000 Low (CRF) vs High (saline) 1 ,36 0.312 1 ,21 1 .000 High (CRF) 1 ,36 0.614 1 ,21 1 .000 High (saline) vs High (CRF) 1 ,36 1 .000 1 ,21 1 .000 5.4 D iscussion Mechanical restraint causing immobilisation was a suitable stressor for identifying differences in the adrenocortical response of individual quail, allowing the identification of birds for low and high response groups. There was a positive dose­ dependent relationship between ACTH and plasma corticosterone concentrations. The corticosterone response to a sub-maximal dose of ACTH did not differ between low and high response quail, showing there was no difference in adrenal sensitivity between the two response groups. Corticosterone responses to CRF were inconsistent, with only one of the four doses initiating clear responses. However, when this dose was given to low and high response birds it did not elicit a response in either group. This finding is similar to results reported in snow buntings (Plectrophenax nivaIis) (Romero et al., 1998b) and Gambel's white-crowned sparrow (Zonotrichia leucophrys gambelii) (Romero and Wingfield, 1 999) where high doses of ovine CRF failed to affect corticosterone concentrations. 5.4 . 1 ACTH dose response After treatment with ACTH plasma corticosterone concentrations increased in a dose dependent manner after 15 minutes and returned to basal after 30 minutes. Chapter Five 177 ACTH treatment has been shown to stimulate an increase in corticosterone concentrations in chickens (Beuving and Vonder, 1 978; Beuving and Vonder, 1 986; Littin and Cockrem, 2001), ducks (Harvey et al., 1 980; Noirault et al., 1999; Faure et al., 2003), and a range of passerine species (Romero et al., 1998a; Romero et al., 1998b; Romero and Wingfield, 1998; Wilson and Holberton, 2001). Carsia et al. (1988) showed that corticosterone secretion increased in Japanese quail adrenocortical cells incubated with ACTH. Beuving and Vonder (1986) reported that a dose of 0.1 IU/kg porcine ACTH initiated a sub-maximal corticosterone response in chickens. In many of these studies large doses of ACTH resulted in a maximal corticosterone response or a prolonged elevation of corticosterone. Studies conducted on passerines (Romero et al., 1998a; Romero et al., 1998b; Romero and Wingfield, 1998) focused mainly on the adrenal sensitivity to ACTH by stimulating a maximal corticosterone response at different stages of the lifecyc1e . The smallest dose of ACTH (0.05 ID; 0.2 ID/kg body weight) resulted in a small but significant increase in corticosterone concentrations 15 minutes after administration. Peak corticosterone concentrations at 15 min increased with doses of O. lD, 0.20 and 0.50 IV (0.4, 0.8 and 2.0 IU/kg respectively). A dose of 0.8 IU/kg generally stimulated a maximal corticosterone response by 15 minutes. Beuving and Vonder (1986) found that a dose of 0.5 IU/kg stimulated a maximal response in chickens after 25 minutes, while Noirault et al. (1999) reported that corticosterone concentrations in ducks peaked 10 minutes after an injection of 0.125 IU/kg. Single reports of adrenal responsiveness to ACTH in a species are not sufficien t to determine whether there are differences between species in these responses, as there are differences in the type of ACTH, age of the birds, species, sexes, and environmental conditions, all making direct comparisons difficult. Unlike previous studies which have focussed on the adrenocortical potential to secrete corticosterone in response to ACTH, the present study was designed to identify a dose that stimulated a sub-maximal corticosterone response. In the current study a dose of 0.05 IU/bird (0.2 IU/kg) stimulated a small increase (approXimately 2 nglml) in corticosterone, which was smaller than the response of birds to mechanical restraint. A maximal response was generally elicited from birds 178 Chapter Five given doses of 0.20 and 0.50 IU ACTH (0.8 and 2.0 IU/kg respectively), with little variation between individuals in the corticosterone response between these two doses. The 0.10 IU (0.4 IU/kg) dose resulted in an increase in corticosterone concentrations of approximately 5 ng/ml after 15 minutes. This was similar to the response of birds to mechanical restraint and, more importantly, there was a wide range of corticosterone responses as seen in responses to mechanical restraint. 5.4.2 Adrenal sensitivity to ACTH in b irds selected for low or high responses There was no difference in adrenal sensitivity to ACTH between Japanese quail selected for low or high responses to mechanical restraint. These results differ from the in vitro results of Carsia et al. (1988) who worked with adrenocortical cells from random-bred quail and from quail selected for a high corticosterone response to immobilization. Carsia et al. (1988) showed that maximal ACTH-induced corticosterone production by cells from the high response quail was greater than that of the random-bred quail. The injection of ACTH stimulated an increase in corticosterone concentrations in both low and high response groups, with net increases in corticosterone at 15 minutes of 5.5 and 5.3 ng/ml respectively. This compares with an increase of 4.3 ng/ml in quail that were mechanically restrained. Plasma corticosterone declined after the peak at 15 minutes and returned to basal at 60 minutes. As shown in the dose response study, the peak in corticosterone concentrations at 15 minutes varied between birds, with similar variation in the low and high response groups. However, the mean corticosterone response to 0.10 IU ACTH was the same in low and high response birds, indicating that differences between birds in their corticosterone responses to mechanical restraint were not due to differences in adrenal sensitivity to ACTH. Chapter Five 179 5.4.3 CRF dose response Intrajugular injection of 2 to 8 J..Lg CRF (8, 1 6, 24 or 32 J..Lglkg body weight) stimulated an increase in plasma corticosterone in some birds after 15 minutes. However, the increase in corticosterone was not consistent and there was no dose-dependent response. The 6 J..Lg (24 J.Lglkg) dose was the only one to stimulate a Significant increase in corticosterone. It is well established that CRF stimulates ACTH secretion from the pituitary in birds, as shown by the incubation of ovine CRF with duck (Castro et al., 1986) or chicken (Carsia et al., 1986) pituitary cells in vitro. CRF stimulates corticosterone secretion from the adrenal gland via ACTH, so changes in corticosterone do not directly reflect the effect of CRF on ACTH secretion. However, the absence of differences between birds with low and high corticosterone responses to mechanical restraint in terms of their adrenal sensitivity to ACTH indicates that corticosterone responses to CRF are likely to reflect differences in pituitary sensitivity to CRF or CRF production. Westerhof et al. (1992) showed that ovine CRF treatment caused corticosterone concentrations to increase in a dose dependent manner in racing pigeons (Calumba livia damestica), with doses of 10 and 100 J..Lglkg stimulating a significant increase in corticosterone concentrations 15, 30 and 60 minutes after the intravenous injection. In the present study corticosterone declined and returned to basal between 15 and 30 minutes in quail treated with 6 J..Lg of CRF (24 J.Lglkg), whereas Westerhof et al. (1992) reported that corticosterone continued to increase between 15 and 60 minutes after the CRF treatment before declining to basal at 120 minutes. In the current study, birds treated with 2 or 6 J.Lg CRF (8 and 24 J.Lglkg) showed the most consistent corticosterone responses to CRF, although only the 6 J..Lg/bird dose produced a significant response. The absence of a consistent and significant corticosterone response in birds treated with 8 J..Lg CRF may indicate possible down­ regulation of CRF-responsiveness of the anterior pituitary gland. The inconsistent corticosterone responses might have resulted from the use of these birds in three previous mechanical restraint or ACTH treatment studies. The range of responses 1 80 Chapter Five to CRF administration between the doses, and the odd responses by single individuals or groups could reflect an effect of repeated manipulations. Jones et al., (2000) found experience-dependent sensitisation of the adrenocortical response in high stress line Japanese quail. It is possible that the corticosterone responses of some individuals be sensitive to repeated handling associated with blood sampling and the injection rather than CRF. In Japanese quail, Launay et al. (1993) reported a slight but significant increase in corticosterone 30 minutes after a single injection of 15 J.Lglkg of human CRF. Ovine CRF was used in the present study, and has been used successfully to stimulate an increase in corticosterone release in a variety of avian species (Castro et al., 1986; Carsia et al., 1986; Westerhof et al., 1992; Romero et al., 1998b). The 6 J.Lglbird CRP dose stimulated a 7 ng/ml increase in corticosterone between 0 and 15 minutes, compared with a 5 ng/ml increase in response to 0.10 IU ACTH. However, although there was more variation between birds in their responses to the ACTH dose than the CRF dose, the 6 ILg dose of CRF was used to treat low and high response groups since it was the only CRF dose to stimulate a significant corticosterone response. 5.4.4 P ituitary sensitivity to CRF i n b irds selected for low and h igh responses Male quail with a low or high corticosterone responses to mechanical restraint were given an intrajugular injection of saline or 6 J.Lg CRF/bird (24 J..Lglkg body weight) . Treatment with CRF did not stimulate a corticosterone response in either the low or high response groups. The absence of a corticosterone response from the CRF treated birds contrasted with the CRF dose response study which showed a clear response to 6 ILg CRF/bird. The corticosterone response to CRF treatment was similar in birds that had been previously treated with saline or a high dose of CRF in the previous CRF dose response study. This suggests that the absence of any significant effect of CRF on corticosterone secretion in low and high response to mechanical restraint quail was Chapter Five 181 not due to previous treatment with CRF. It is more likely that the CRF was ineffective, or that the response to CRF was affected by the three previous sampling sessions experienced by the birds. Corticosterone concentrations at 0 minutes were higher and more variable than had been previous measured and this confounded the results. These quail were 15 weeks old and had been used in up to six previous experiments. It is possible that the birds were responding before being picked up when people were entering the quail rooms . This could be due to their previous experiences of being handled and blood sampled several times over the previous 8 weeks, as seen in the response in domestic fowl which habituate to repeated injections of corticotrophin (Freeman et al., 1 979) . The results of the present study can also be compared with those previously reported in passerines where responses to with an ovine CRF dose selected to saturate the pituitary have been inconsistent (Romero et al., 1998a; Romero et al., 1998d; Romero and Wingfield, 1999) . 5.4.5 Summary The aim of this s tudy was to determine if individual variation of the corticosterone response is caused by differences in adrenal or pituitary sensitivity to ACTH or CRF respectively, in Japanese quail selected for low or high corticosterone responses to mechanical restraint. Adrenal sensitivity to ACTH did not differ between quail with low or high corticosterone responses, while CRF treatment did not stimulate corticosterone responses consistently. The stress response initiates physiological and behavioural responses that allow a bird to adapt to the source of the stressor, with some individuals better able to respond to stressors than others. There is marked variation of the corticosterone response between birds (Wingfield et al., 1992; Wingfield, 1 994; Astheimer et al., 1995) and this variation exists due in part, to differences in the HPA-axis (Carsia et al., 1 988) and higher centres in the brain. In mammals, exposure to stressful stimuli has been shown to alter an individual's subsequent responses (Beylin and Shors, 2003), and it is likely that a similar mechanism exists in birds. There is also a genetic basis to corticosterone responses, as shown by the selection of quail for low 182 Chapter Five or high responses (Satterlee and Johnson, 1988) . In this study, adrenal sensitivity to ACTH did not differ between low and high response quail indicating that the difference in sensitivity to a pharmacological dose of ACTH found by Carsia et al. (1988) may have a negligible if any influence on variation of the corticosterone response to a stressor. The present data indicate that individual variation is caused at a site higher than the adrenal gland. The use of ovine CRF to assess pituitary sensitivity in low or high corticosterone response quail produced inconsistent results. Ovine and human CRF have been used in a variety of bird species to stimulate ACTH and corticosterone secretion with mixed success (Westerhof et al., 1992; Romero and Wingfield, 1998) . Despite the sensitivity of some birds to ovine and human CRF indicating the existence of an avian equivalent, an avian CRF has yet to be characterised. There is some evidence that AVT and MT are necessary for the action of CRF (Westerhof et al., 1992; Romero and Wingfield, 1998), although the stimulation of ACTH secretion has been inconsistent with both AVT and MT in isolation and when used in combination with CRF. The inconsistent responses to CRF meant that the role of pituitary gland sensitivity to CRF in contributing to variation between birds in their corticosterone responses could not be assessed. The inconsistent responses might have resulted from reusing birds from the earlier mechanical restraint, ACTH, and CRF dose response studies. This problem could be overcome in a future study by choosing birds with low or high responses to mechanical restraint from a new group of quail, thereby avoiding the reuse of birds in the ACTH study. Alternatively, establishing a colony of quail and breeding for low or high corticosterone responses to mechanical restraint as done in the Satterlee laboratory would produce completely naive birds for CRF treatment. It was not possible to conduct such studies for the present thesis research. However, the current study did show that adrenal sensitivity to a physiological dose of ACTH does not differ between quail selected for low or high corticosterone responses to mechanical restraint, indicating that individual variation is mediated at the pituitary gland or at high brain centres responsible for activa ting the HPA-axis. Chapter Six Chapter 6 General discussion 183 The aims of the research described in this dissertation were to determine the effects of different stressors on corticosterone and sex steroid secretion in Japanese quail, and to investigate the neuroendocrine basis of variation between birds in their responses to stressors. The size of corticosterone and testosterone responses to stressors were dependent on the type and duration of the stimuli. Manual and mechanical restraint elicited larger corticosterone responses than other stressors, whilst the size of the corticosterone response increased with increasing duration of exposure to stressors. Manual and mechanical restraint also had the greatest inhibitory effect on testosterone secretion. This inhibitory effect on testosterone occurred independently of any change in LH secretion. This is one of the first reports of testosterone secretion in birds being inhibited by a single, short period of handling, and has implications for future studies in domestic and free-living birds where sex steroid concentrations are measured in birds that may already have been handled. The effect that a stressor has on the reproductive system is influenced by an individual's perception of the stressor and the magnitude of the response to the stressor. The magnitude of the response of a bird to a stressor has a genetic basis, and is also influenced by the bird's previous experience of stressors. These factors differ between individuals and lead to variation in the stress response between individuals. The corticosterone response differed markedly between individual random-bred Japanese quail in this study, and between individuals of the strains of 184 Chapter Six quail selected for low or high stress responses by Satterlee and Johnson (1988). Individual differences in the ability to respond to stressors may contribute to variation in breeding success and survival (reviewed by Sapolsky and Wingfield, 2003). There has been considerable interest in corticosterone responses in free­ living birds in relation to environmental conditions and latitude (Wingfield, 1994; Romero et al., 1998a) . However, little research has been conducted on how individual variation of the corticosterone response arises. In the present study, it was found that variation between birds in their corticosterone responses was caused by differences in adrenal sensitivity to ACTH. This indicates that individual variation is due to individual differences in pituitary sensitivity to CRF, AVT, MT or by differences at a higher level in the brain. The importance of these findings will be discussed in greater detail in the remainder of this chapter, and suggestions will be made for future potential areas of study. Japanese quail were selected as a model species due to their availability and the large amount of previous research conducted overseas . There are no previous reports of experimental work with Japanese quail in New Zealand, so a series of pilot studies, not included in this dissertation, were conducted to identify the conditions that either stimulated gonadal regression or prevented gonadal growth as a prerequisite to studies of the effects of corticosterone on the control of the reproductive system. In Chapter 2, a single subcutaneous 1 .2 mg corticosterone injection was used to increase plasma corticosterone to pharmacological concentrations to determine if corticosterone treatment inhibited sex steroid secretion. LH concentrations in males were unaffected by corticosterone treatment showing that a short « 30 min) elevation in corticosterone did not influence LH secretion from the pituitary. Handling associated with the blood collection and injection caused a decline in plasma testosterone that masked any effect of corticosterone treatment. Eskeland and Blom (1979) and Johnson (1981) had previously reported a decline in testosterone after repeated handling at 5 minute intervals. Less frequent blood sampling and similar doses of corticosterone have been used in Japanese quail and caused a decline in testosterone secretion (Boyd, 2000). Corticosterone is also Chapter Six 185 known to inhibit several enzymes involved in testicular steroidogenesis, causing a decrease in testosterone secretion in isolated rat leydig cells (Bambino and Hsueh, 1981, Mann and Orr, 1990; Orr and Mann 1992, Monder et al., 1994) . It is proposed that corticosterone acts in a similar fashion in the Japanese quail testes, and the increase in corticosterone associated with repeated handling accounts for the decline in testosterone secretion. While a variety of stimuli were shown to stimulate a corticosterone response, manual and mechanical restraint were the most effective in stimulating an increase in plasma corticosterone and a decrease in testosterone secretion. A single period of manual or mechanical restraint caused a decrease in testosterone for more than 2 hours. In contrast, there is a single report of an increase in testosterone secretion following 10 minutes of mechanical restraint (Heiblum et al., 2000). Mechanical restraint was the preferable method for characterising individual variation in the adrenocortical response in our quail and was comparable to the method used by Satterlee and Johnson (1988) . Mechanical restraint was lOgistically easy to apply making it preferable over the intermittent handling method used previously for quail (Boyd, 2000) and chickens (Littin and Cockrem, 2001). Individual variation of the corticosterone response to immobilisation by manual or mechanical restraint has been reported in a range of avian species (Boyd, 2000; Littin and Cockrem, 2001; Cockrem and Silverin, 2002a) . Free-living birds also differ in their stress response to an identical stressor. This variation, influenced by differences in body condition and social status may be related to variation in survival and reproductive performance (reviewed by Wingfield and Sapolsky, 2003). In previous studies, quail have been selected for divergent corticosterone responses and these strains have been used to study the relationship between the adrenocortical response and behavioural, physiological and genetic traits (Satterlee and Johnson, 1988; Jones et al., 1992a; Jones et al., 1994a; Satterlee and Jones, 1997; Jones et al., 2000; Od eh et al., 2003a; Odeh et al., 2003b). However, few studies have specifically addressed the exact causes of individual variation of the corticosterone response. 186 Chapter Six The corticosterone response varies between individuals in response to a range of different stressors including immobilisation (Satterlee and Johnson, 1988), handling (Littin and Cockrem, 2001; Cockrem and Silverin, 2002a), low temperature (Brown and Nestor, 1973), or social stress (Gross and Siegel, 1985) . Variation could arise at the CNS or at some level of the HPA-axis, or a combination of both. The CNS plays an important part in the magnitude and duration of the HPA response activated in an individual. Following activation of the HPA-axis, differences in pituitary and adrenal sensitivity to products of the HPA-axis are also likely to influence the corticosterone response. The general pattern of the corticosterone response for an individual was found to be repeatable when quail were handled on three occasions 1 0 days apart. A single corticosterone response from a bird therefore can be used as a measure of that bird's responsiveness to the stressor. In previous studies, quail have been selected for low or high corticosterone responses based on the response to a stressor on one occasion. This study confirms that single corticosterone responses of quail can be used as a basis for selection of birds with low or high corticosterone responses. ACTH injections have been used previously to assess adrenal sensitivity to ACTH (Romero et al., 1998a; Romero and Wingfield, 1998; Romero et al., 1 998b). ACTH treatment has also been used to determine adrenocortical cel l sensitivity in vitro in random-bred quail and quail selected for a high corticosterone response to immobilisation (Carsia et al., 1988) . In the current study, an i.v. injection of ACTH at a dose of 0. 10 IUlbird stimulated a range of corticosterone responses that was similar to the range of responses to mechanical restraint. The 0.10 IU injection of ACTH was equally effective in stimulating a corticosterone response in birds with low or high corticosterone responses to mechanical restraint. This showed that individual variation in the corticosterone response to the restraint stressor is not regulated by adrenal sensitivity to ACTH. In contrast, Carsia et al . (1988) showed that the adrenocortical cells of quail selected for a high stress response to mechanical restraint were more sensitive to ACTH than those of random-bred quail . However, isolated adrenocortical cells of high stress quail only differed in their response to pharmacological doses of ACTH and basal corticosterone secretion Chapter Six 187 in cells from high response birds did not differ from that of cells from random-bred birds. Carsia et al. (1988) proposed that adrenocortical cells in random-bred and high response quail are equally sensitive to ACTH, but that the cells of the high stress line are more able to couple ACTH with transmembrane signals and steroidogenic enzymes associated with corticosterone synthesis. In the current study, birds were treated with a sub-maximal dose of ACTH at physiological concentrations, resulting in similar corticosterone responses in low and high response quail. This suggests that the in vitro difference in response to ACTH found by Carsia et al. (1988) does not contribute to the variation in vivo. The avian pituitary gland has been shown to be responsive to ovine CRF both in vitro and in vivo in a variety of species (Carsia et al., 1986; Castro et al., 1986; Westerhof et al., 1992; Launay et al., 1993; Romero et al., 1998b). Treating quail with a single i.v. injection of oCRF stimulated an increase in corticosterone secretion, indicating that ACTH secretion from the anterior pituitary gland was responsive to CRF. However, CRF-treatment of quail with low or high corticosterone responses to mechanical restraint was ineffective, in contrast to the CRF dose response study where some quail were responsive to a dose of 6 ILg CRFlbird. The absence of any response to CRF meant it was not possible to determine whether the variation in responses to mechanical restraint was due to variation in pituitary responsiveness to CRF. Previous studies (Romero et al., 1998b; Romero and Wingfield, 1999) have shown CRF treatment to be inconsistent, particularly where changes in corticosterone concentrations are used to assess ACTH secretion. It is possible that other factors are required in conjunction with CRF treatment to cause a repeatable and robust increase in corticosterone secretion. Treatment with CRF may need to be preceded, accompanied, or followed by treatment with A VT or MT. CRF treatment in vivo in combination with AVT or MT have been shown to be effective in causing an increase in plasma corticosterone via ACTH in a pigeons, quail and passerines (Westerhof et al., 1992; Romero et al., 1998b) . Further studies are necessary to examine the pituitary gland as a site where individual variation of the corticosterone response may be regulated. CRF treatment of low or high response quail needs to be repeated to assess the 188 Chapter Six responsiveness of the pituitary gland to CRF in quail which show a low or high response to mechanical restraint. Such a study would include AVT and MT treatment in isolation or in combination with CRF as well as the measurement of plasma ACTH concentrations. If the pituitary glands of low and high response quail did not differ in their responsiveness to physiological doses of CRF, A VT or MT in vivo, the responses of pituitary cells isolated from low and high response quail to pharmacological doses of CRF, A VT, or MT could differ in vitro. As discussed earlier, Carsia et al. (1988) showed that maximal ACTH-induced corticosterone production differed between quail with divergent corticosterone responses to mechanical restraint, and a similar difference may exist in pituitary cells of low or high response quail. Any such differences may have little effect in vivo at physiological concentrations of the hypothalamo-hypophysiotrophic factors. The CNS is an area that has received little a ttention in birds when assessing the causes of individual variation in stress responses. Stressors have been shown to activate different regions of the brain in quail (Takeuchi et al., 1996) and the degree of activation and activation of different regions are likely to vary between individuals exposed to the same stressor. The activation of the endocrine stress response is via the CNS, and differences in the measured corticosterone response between individuals exposed to the same s tressor may reflect differences in the activation of neurons in the CNS. Individual variation in the corticosterone response to mechanical restraint is likely to be mediated by differential activation of the regions of the brain involved with the initial stress response. Immunoreactivity of the immediate early gene c-fos can be used to identify the activation of neurons in the CNS in response to photoperiodic (Meddle and Follett, 1995), reproductive (Meddle et al., 1999) and stressor s timuli in Japanese quail (Takeuchi et al . , 1996). The distribution and intensity of c-fos immunoreactivity has been used in mammals and birds to determine the neural pathways activated by different stimuli. Immobilisation stimulates an increase in fos-like immunoreactivity in the lateral preoptic area, Chapter Six 189 median preoptic, paraventricular, arcuate, dorsomedial hypo thalamic nuclei and lateral hypo thalamus (Briski and Gillen, 2001). Measuring c-fos expression in quail selected for low or high corticosterone responses to mechanical restraint would show if the activated neural pathways differed between the two strains. Locations of c-fos protein expression could be identified in low or high corticosterone response quail using either an antibody raised against c-fos or probes for c-fos mRNA for in situ hybridisation. Once the precise location of stressor-induced c-fos expression in the quail hypo thalamus was determined, c-fos mRNA expression could be quantified using a c-fos mRNA quantitative competitive polymerase chain reaction (PCR) established and validated for quail hypothalamic tissue during a recent visit to a collaborator's laboratory. In order to understand the physiological basis of individual variation in the corticosterone response it is necessary to understand the neural and endocrine mechanisms that generate a corticosterone response following exposure to a stressor. The results of the current study (summarised in Figure 6) show that the adrenocortical responses of Japanese quail differ in relation to the type and duration of the stressor and show that the response to a standardised stressor is generally repeatable for an individual quail. Differences in adrenocortical responses between individuals are likely to have a genetic or neurological basis and to be modified by experience. The precise locations in the HPA-axis that mediate variation of the response are not known, although the results of this study indicate that the locations are the pituitary gland and higher centres in the brain. Therefore, future work with quail that have divergent corticosterone responses should include an examination of pituitary gland sensitivity to hypothalamo-hypophysial factors and the quantification of differences in neuronal activation in brains regions associated with the stress response. 190 Stressors - type - duration Brain Activate the HPA axis with varying intensity Chapter Six No Amygdala No HPA 4--+-Stressor is threat? Individual variation response GnRH 1 1 �o/ Individual variation [ Pitu itary g land H Individual variation ] Fy �H Gonad Sex steroids Adrenal g land No individual variation �-------.. Corticosterone Inhibits sex steroid secretion Figure 6 Schematic representation of the stress response in Japanese quail. An acute stressor activates the hypothalamic-pituitary-adrenal axis causing an increase in corticosterone secretion. 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