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Neurosteroids and stress physiology in adult songbirds Newman, Amy Elida Margaret 2009

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NEUROSTEROIDS AND STRESS PHYSIOLOGY IN ADULT SONGBIRDS  by  AMY ELIDA MARGARET NEWMAN B.Sc., Queens University, 2004  A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY  in  THE FACULTY OF GRADUATE STUDIES (Neuroscience)  THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver)  April 2009  © Amy Elida Margaret Newman, 2009  ABSTRACT Stress increases adrenal glucocorticoid secretion, and chronic elevation of glucocorticoids can have detrimental effects on the brain. Dehydroepiandrosterone (DHEA) is an androgen precursor synthesized in the adrenal glands, gonads or the brain and has anti-glucocorticoid properties. However, little is known about the role of DHEA in the stress response, particularly in the brain. In Chapter 2, I validated a solid phase extraction technique for extracting steroids from lipid-rich brain tissue and plasma of songbirds. In Chapter 3, I demonstrated that acute stress had statistically significant effects on plasma corticosterone and DHEA in wild adult male song sparrows that were season and vein specific. For corticosterone, acute stress increased jugular levels more than brachial levels during the molt. For DHEA, acute stress did not affect brachial DHEA but decreased jugular DHEA during the breeding season and increased jugular DHEA during the molt. These results suggest that corticosterone and DHEA are locally synthesized in the brain during molt. In Chapter 4, I measured the effects of acute stress and season on corticosterone and DHEA in brain tissue and jugular plasma. Compared to jugular plasma, corticosterone levels were up to 10× lower in brain, whereas DHEA levels were up to 5× higher in brain and were highest in the hippocampus. Acute stress increased corticosterone levels in jugular plasma and brain, except during molt, when stress decreased corticosterone levels in the hippocampus. In Chapter 5, I tested the effects of corticosterone and DHEA treatments on the brain. Corticosterone and DHEA had additive effects on the volume, neuron number and recruitment of new cells into HVC. Elsewhere in the brain, DHEA increased BrdU+ cells only in the absence of corticosterone suggesting that corticosterone can interfere with the action of DHEA. Together, these studies demonstrate that acute stress and season have distinct effects on corticosterone and DHEA in plasma and brain. Furthermore, I demonstrate that corticosterone and DHEA can have additive effects on cell survival and recruitment in the adult brain and that, in some cases, corticosterone can inhibit the actions of DHEA in the brain. ii  TABLE OF CONTENTS ABSTRACT ................................................................................................................................. ii TABLE OF CONTENTS ........................................................................................................... iii LIST OF TABLES....................................................................................................................... v LIST OF FIGURES.................................................................................................................... vi ABBREVIATIONS.................................................................................................................... vii ACKNOWLEDGEMENTS ..................................................................................................... viii DEDICATION ............................................................................................................................ ix CO-AUTHORSHIP STATEMENT ........................................................................................... x 1  GENERAL INTRODCUTION ........................................................................................... 1 GLUCOCORTICOIDS, STRESS and the BRAIN ........................................................... 1 DHEA, STRESS and the BRAIN...................................................................................... 2 LOCAL STEROID SYNTHESIS IN THE BRAIN .......................................................... 3 SONG SPARROWS AS A MODEL SYSTEM................................................................ 3 OVERVIEW ...................................................................................................................... 5 FIGURES........................................................................................................................... 7 REFERENCES .................................................................................................................. 8  2  ANALYSIS OF STEROIDS IN SONGBIRD PLASMA AND BRAIN BY COUPLING SOLID PHASE EXTRACTION TO RADIOIMMUNOASSAY .................................... 11 INTRODUCTION ........................................................................................................... 11 MATERIALS and METHODS ....................................................................................... 13 RESULTS ........................................................................................................................ 17 DISCUSSION.................................................................................................................. 20 TABLES .......................................................................................................................... 24 FIGURES......................................................................................................................... 26 REFERENCES ................................................................................................................ 29  3  DHEA AND CORTICOSTERONE ARE REGULATED BY SEASON AND ACUTE STRESS IN A WILD SONGBIRD: JUGULAR VERSUS BRACHIAL PLASMA..... 32 INTRODUCTION ........................................................................................................... 32 MATERIALS and METHODS ....................................................................................... 34 RESULTS ........................................................................................................................ 38 DISCUSSION.................................................................................................................. 42 TABLES .......................................................................................................................... 48 FIGURES......................................................................................................................... 53 REFERENCES ................................................................................................................ 56  iii  4  CORTICOSTERONE AND DHEA IN SONGBIRD PLASMA AND BRAIN: EFFECTS OF SEASON AND ACUTE STRESS ............................................................. 61 INTRODUCTION ........................................................................................................... 61 MATERIALS and METHODS ....................................................................................... 62 RESULTS ........................................................................................................................ 67 DISCUSSION.................................................................................................................. 71 TABLES .......................................................................................................................... 76 FIGURES......................................................................................................................... 80 REFERENCES ................................................................................................................ 84  5  EFFECTS OF CORTICOSTERONE AND DHEA ON NEURAL PLASTICITY IN THE ADULT SONGBIRD BRAIN.................................................................................... 89 INTRODUCTION ........................................................................................................... 89 METHODS ...................................................................................................................... 91 RESULTS ........................................................................................................................ 98 DISCUSSION................................................................................................................ 101 TABLES ........................................................................................................................ 108 FIGURES....................................................................................................................... 112 REFERENCES .............................................................................................................. 118  6  GENERAL DISCUSSION .......................................................................................... 123 SUMMARY................................................................................................................... 123 LIMITATIONS ............................................................................................................. 124 FUTURE DIRECTIONS ............................................................................................... 126 CONCLUSIONS ........................................................................................................... 128 REFERENCES .............................................................................................................. 129  iv  LIST OF TABLES Table 2.1 Table 2.1 Recovery of 3H-DHEA and 3H-E2 from chicken serum using solid phase extraction……………………………………………………………………………21 Table 2.2 Recovery of 3H-DHEA and 3H-E2 from chicken serum extracts dried under N2 or under air after solid phase extraction………………………………………………..22 Table 3.1 No significant differences between 30 min restraint and 60 min restraint………….48 Table 3.2 Effect of Season, Vein and Stress on plasma CORT and DHEA is dependent on season and vein………………..…………………………………………………….49 Table 3.3 Effects of Vein and Stress on plasma CORT and DHEA within each season……...50 Table 3.4 CORT/DHEA ratio at baseline and after acute restraint tress……………………....51 Table 3.5 Seasonal changes in body condition, fat and cloacal protuberance length………….52 Table 4.1 Effects of season, stress and brain region on corticosterone and DHEA in the brain…..……………………………………………………………………………..76 Table 4.2 Effects of season and sample type on corticosterone and DHEA levels……………77 Table 4.3 Effects of season and stress on corticosterone levels in peripheral tissues……...….78 Table 4.4. Effects of season and stress on DHEA levels in peripheral tissues……………...….79 Table 5.1 Effects of steroid treatment on NeuN measurements and BrdU+ cell number.…...108 Table 5.2 Effects of steroid treatments on plasma corticosterone levels (ng/mL) (Experiment 1)……………………………………………………………………..109 Table 5.3 Effects of corticosterone treatment on plasma corticosterone levels (ng/ml) (Experiment 2)………………………………………………………………...…...110 Table 5.4 Effects of steroid treatments on plasma DHEA levels (ng/ml) (Experiment 1)…...111  v  LIST OF FIGURES Figure 1.1  Simplified diagram of steroid biosynthesis………………………………………7  Figure 2.1  Effects of ethanol on the DHEA radioimmunoassay ………………………..….26  Figure 2.2  Comparing Dichloromethane Extraction and Solid Phase Extraction….…….…27  Figure 2.3  Solid phase extraction protocol for songbird plasma and brain tissue…….……28  Figure 3.1  Serially diluted plasma in corticosterone and DHEA radioimmunassays.…...…53  Figure 3.2  Effects of season on baseline corticosterone and DHEA in brachial and jugular plasma…………….……………………………………………………………..54  Figure 3.3  Effects of season and acute restraint stress on corticosterone and DHEA in jugular and brachial plasma……………………….…………………………….55  Figure 4.1  Effects of season and acute stress on corticosterone in jugular plasma and brain………………………………………………………………………….….80  Figure 4.2  Effects of season and acute stress on DHEA levels in plasma and brain……….81  Figure 4.3  Comparison of corticosterone and DHEA in plasma and brain…………………82  Figure 4.4  Effects of season and acute stress on corticosterone / DHEA ratio…………..…83  Figure 5.1  Timeline for experiments 1 and 2……………………………………...………112  Figure 5.2  Representative immunocytochemical staining for BrdU and NeuN…………..113  Figure 5.3  Effects of corticosterone and DHEA treatment on HVC………………………114  Figure 5.4  Effects of corticosterone and DHEA treatment on the telencephalon…………115  Figure 5.5  Effects of corticosterone and DHEA treatment on the hippocampus………….116  Figure 5.6  Effects of corticosterone and DHEA treatment on BrdU+ cells in the ventricular zone………………………………………………………………..117  vi  ABBREVIATIONS 11β-HSD: 11β-hydroxysteroid dehydrogenase  HPA: hypothalamic-pituitary-adrenal HPG: hypothalamic-pituitary-gonadal  3β-HSD: 3β-hydroxysteroid dehydrogenase/isomerise  Hp: hippocampus  ACTH: adrenocorticotropin hormone  HSD: honestly significant difference  AE: androstenedione  HVC: HVC (proper name)  ANOVA: analysis of variance  MeOH: methanol  BDNF: brain derived neurotrophic factor  N2: nitrogen  BrdU: bromodeoxyuridine  NCM: caudal medial nidopallium  CBG: corticosteroid-binding-globulin  P450c11: cytochrome P450 11β−hydroxylase  cDien: caudal diencephalon rDien: rostral diencephalon CORT: corticosterone  P450c21: cytochrome P450 21αhydroxylase P450scc: cytochrome P450 side chain cleavage  CRH: corticotropin-releasing-hormone cmTel: central medial telencephalon  PBSg: phosphate buffered saline with gelatine  DAB: diaminobenzidine  Pgp: P-glycoprotein  DCM: dichloromethane  PREG: pregnenolone  DHEA: dehydroepiandrosterone  PROG: progesterone  DHEAS: dehydroepiandrosterone-sulphate  RIA: radioimmunoassay  dTel: dorsal telencephalon  SEM: standard error of the mean  E2: 17β-estradiol  SPE: solid phase extraction  EtOH: ethanol  SVZ: subventricular zone  GC: glucocorticoid  T: testosterone  GnRH: gonadotropin-releasing-hormone  VZ: ventricular zone  vii  ACKNOWLEDGEMENTS I am indebted to many. First, to Dr. Kiran Soma, as an advisor, teacher, critic, and importantly, mentor. Your honesty, insight, guidance and stalwart patience have been essential, both for the work summarized here and my development as a scientist. I am proud to have been a part of the lab from its inception and have learned much about who I am and who I can be. Thank you. To Peter Arcese, Liisa Galea and Joanne Weinberg for providing the attention to detail and kind criticism that kept me on my toes and improved the depth and clarity of this thesis. Your time and thoughtful input are deeply appreciated. To Scott MacDougall-Shackleton for kindly providing a part-time academic home, for tolerating my taking over the wet lab, for sacrificing countless hours to the cause of Chapter 5 and for a door always open. To the members of the Soma laboratory, both past and present, you have enriched my life in many ways, and I shall avoid mutual embarrassment by skimping on details. However, you have all certainly helped me immensely, particularly when I’ve been distant; you’ve kept me a part of the group and welcomed me with warmth during my periodic returns. I am hopeful our friendships shall continue long into the future: Princess, Shuffles, Lor-lets keep it clean-etta, RoNo-like-you’re-smarter-veena, Prickaleena (thank you for your friendship and keeping me well fed), and those fortunate enough to escape a moniker, Kelvin, Thierry (for our unforgettable adventures in poultry science, your comfy futon, and tolerating my attempts en francais), Eunice (always one to keep long nights in the lab interesting), Amit, Matt & Sarah (I hope to have further opportunities to employ sushi bribes). Last, Mikee, thank you for breaking the mold of first impressions, for your friendship, and for “laying me down a pallet on your floor”. To the brave souls, David Hope, Alex Leung, Lani Sheldon, Yong-Seok An, Brandon Yardy, Prick and Kim who braved pre-sunrise mornings in all seasons, tramped through the treatment plant and landfill, straddled sewage outflow pipes, tolerated voracious insects, shook bushes is the stealthiest of manners and gripped frigid aluminum in gathering snow, all to set a net and make a catch. Also, to the gentlemen of the islands, although your boating left something to be desired, your company surely did not. Your resilience was inspiring and many a good yarn was stitched forever into the tapestry. To my family, particularly my mum and brother who, while not knowing what it is I do or why I do it, continue to lend their unwavering support, love and sense of humour while reminding me of the other important things in life. To Too, I appreciate your encouragement and our long chats. Yenni, these past 2 years have been easier thanks to you. To Dolly, a true friend, excellent lap warmer and one who reminded me to get out for fresh air and perspective. And to my extended family, friends, places and things that maintained my sanity (almost) throughout this journey, including pph, bbt, gg, pt, lt, mrb, b&s, enlightened 4th floor dwellers, Jodi Pawluski, inhabitants of my ipod, HBO, Skeleton Lake and Islay. I am grateful for the financial support of the Natural Sciences and Engineering Research Council (NSERC) postgraduate scholarship and Canadian graduate scholarship and the Michael Smith Foundation for Health Research Junior Trainee Award. Finally, to Ryan, my husband (who read every word), best friend and greatest love, you swept me off my feet years ago and have kept me aloft ever since. You never cease to surprise me and continue to be a deep well of inspiration. I am fortunate we travel this road together. viii  EDICATION  For Brutus, and his fine feathered friends  ix  CO-AUTHORSHIP STATEMENT Chapter 2 was co-written with Eunice Chin. We equally shared in designing and performing the experiments, as well as analyzing the data. Kim Schmidt aided in sample collection and analysis. The initial extraction technique was learned in Kathy Wynne-Edward’s laboratory at Queen’s University with the assistance of Leah Bond. I prepared the manuscript. Devaleena Pradhan assisted with field work for Chapter 3. I collected and analyzed the data for Chapter 4 and wrote the manuscript. The experiments described in Chapter 5 were conducted in Dr. Scott MacDougal-Shackleton’s laboratory and Yong-Seok An and Pralle Kriengwatana assisted with field work, blood sample collection and animal care.  x  1  GENERAL INTRODCUTION  GLUCOCORTICOIDS, STRESS and the BRAIN It is well known that acute stress activates the hypothalamic-pituitary-adrenal (HPA) axis. In response to stress, the hypothalamus secretes corticotropin-releasing-hormone (CRH) which stimulates the anterior pituitary to release adrenocorticotropin hormone (ACTH) into the general circulation. ACTH stimulates adrenal synthesis and release of glucocorticoids which are potent modulators of behaviour and physiology (Sapolsky et al., 2000). The term “stress” is widely used and suffers from ambiguity. Throughout this dissertation, we adopt Bruce McEwen’s definition whereby a stressor is a threat, either real or perceived, to the physical and/or psychological integrity of the individual (McEwen, 2000). Further, the acute response to stress is characterized by elevated glucocorticoid levels. While the HPA response to stress is adaptive over the short-term, long-term exposure to stress or elevated glucocorticoids can disrupt behaviour, physiology and neuroanatomy (McEwen, 2007). Specific regions in the brain are especially susceptible to stress and glucocorticoids. For example, the mammalian hippocampus expresses high levels of glucocorticoid receptors (McEwen, 1999) and chronic elevations in corticosterone can lead to hippocampal atrophy and corresponding deficits in learning and memory (Sapolsky, 2000). In a primate, chronic stress increased cortisol levels and reduced cell proliferation in the hippocampus (Czeh et al., 2001). In the adult male rat dentate gyrus, acute stress and corticosterone treatment decrease cell proliferation (Tanapat et al., 2001; Cameron and Gould, 1994), while adrenalectomy can increase cell proliferation (Cameron and Gould, 1994). Interestingly, in adult female rats, exposure to an acute stressor (predator odour) does not affect cell proliferation in the dentate gyrus (Falconer and Galea, 2003). Also, in male songbirds, food restriction stress or exogenous corticosterone treatment during early development reduces the size of the song control nucleus HVC during adulthood (Buchanan et al., 2004). Last, in humans, stress can precipitate or worsen psychiatric illnesses, such as 1  depression (Gold and Chrousos, 2002), and a reduction in hippocampal volume has been associated with depression (Duman, 2004).  DHEA, STRESS and the BRAIN Dehydroepiandrosterone (DHEA), a precursor to testosterone and estradiol, and its sulphated ester DHEAS, are the most abundant steroids in the human circulation. Like glucocorticoids, DHEA can be regulated by ACTH in human plasma (Arvat et al., 2000), and by acute stress and ACTH in the rodent brain (Corpechot et al., 1981; Torres and Ortega, 2003). Although there is a large volume of literature on the regulation of glucocorticoids by stress, there is little known about the regulation of DHEA by stress. There is some behavioural evidence that DHEA treatment may alleviate symptoms of stress-related psychiatric illness, such as negative mood and cognitive deficits associated with depression (Wolkowitz et al., 1997). Also, the cortisol/DHEA ratio, rather than measurements of either steroid alone, more accurately predict the occurrence and duration of symptoms associated with psychiatric illness (Young et al., 2002; Ritsner et al., 2004), which further suggests a relationship between glucocorticoids and DHEA. In the brain, DHEA is thought to have neuroprotective and antiglucocorticoid effects. Male rats subcutaneously implanted with 100 mg DHEA pellets have reduced neuronal injury following ischemia (Li et al., 2001). Also, DHEA can protect the hippocampus from excitatory amino acid toxicity induced by neurotransmitters such as glutamate (Kimonides et al., 1998; Veiga et al., 2003). DHEA also counteracts the effects of corticosterone on cell proliferation and survival in the hippocampus, both in vivo and in vitro. In adult male rats, DHEA inhibits corticosterone-induced suppression of adult hippocampal neurogenesis (Karishma and Herbert, 2002). In vitro, DHEA reduces corticosterone-induced cell death in primary hippocampal tissue cultures (Kimonides et al., 1999). Together, these data suggest that DHEA has powerful neuroprotective effects. 2  LOCAL STEROID SYNTHESIS IN THE BRAIN Over 25 years ago, Baulieu and his colleagues first coined the term “neurosteroid” when they provided evidence that DHEA was locally synthesized in the rodent brain independent of the gonads and adrenals (Corpechot et al., 1981). Since then, several studies have confirmed that, in addition to synthesis by the adrenal glands, glucocorticoids and DHEA can be synthesized de novo from cholesterol in the brain (Fig 1.1) (Davies and MacKenzie, 2003; Gomez-Sanchez et al., 2005; Labrie et al., 2005). Local glucocorticoid synthesis in brain tissue could increase neural glucocorticoid concentrations independently of changes in peripheral glucocorticoid secretion. Similarly, locally synthesized DHEA may play a direct role in buffering the effects of increased concentrations of corticosterone in the brain. These studies contribute to an evolving shift from the traditional view of the brain as a passive recipient of systemic steroids to the more recent view that local steroid synthesis may play an integral role in neural development and function (Schmidt et al., 2008). However, the regulation of brain-derived steroids, particularly after stress and across seasons, is not well understood.  SONG SPARROWS AS A MODEL SYSTEM While DHEA and DHEAS are the most abundant circulating steroid hormones in humans, traditional small animal models such as rats and mice have very low levels of circulating DHEA(S) (Corpechot et al., 1981). Moreover, experimental doses of DHEA in rodent studies have been well above the physiological range (e.g. Karishma and Herbert, 2002), and thus are difficult to interpret. In contrast, several lines of evidence suggest that songbirds are a good model with which to investigate the actions and mechanisms of DHEA in the nervous system. Songbirds have high circulating levels of DHEA (Soma and Wingfield, 2001), which supports the face validity of this system as DHEA(S) are the most abundant circulating steroids in 3  humans. Second, a physiological dose of DHEA has significant effects on neuroanatomy and behaviour, such that treatment of nonbreeding song sparrows with a 7mm silastic DHEA implant significantly increased song behaviour and increased HVC size by 51% (Soma et al., 2002). Third, acute restraint stress and ACTH treatment increase peripheral glucocorticoid levels in song birds (Romero et al., 1998), similar to the effects of acute stress and ACTH in humans (de Jong et al., 2007; McEwen, 2000), thus supporting the construct validity of this system. Furthermore, wild songbirds, are exposed to a range of natural stimuli that change across seasons, and understanding the effects of season is important for a complete understanding of neurosteroids and stress physiology. In songbirds, the activity of both the hypothalamicpituitary-gonadal (HPG) and HPA axes change seasonally. In male house sparrows during the breeding season, the numbers of gonadotropin releasing hormone (GnRH) neurons in the songbird hypothalamus are increase, gonads recrudesce and levels of circulating testosterone (T) are elevated. During the nonbreeding season, GnRH neuron numbers decline, gonads regress and T levels are nondetectable (Hahn and Ball, 1995). Like T, baseline corticosterone levels are also elevated during the breeding season, which is in contrast to the HPA-HPG interaction in male rodents, where elevations in corticosterone suppress testosterone production and reproductive behaviours (Wingfield and Sapolsky, 2003). Interestingly, levels of DHEA are elevated during both the breeding and nonbreeding season (Soma and Wingfield, 2001), suggesting a seasonal shift from systemic to local steroid synthesis. In conclusion, the song sparrow (Melospiza melodia) provides an excellent opportunity for studying stress and neurosteroids because 1) adult neurogenesis is robust in songbirds (Nottebohm, 2002), 2) the adult song control circuit is plastic and changes both seasonally (Tramontin and Brenowitz, 2000) and in response to DHEA treatment (Soma et al., 2002) and 3)  4  levels of plasma corticosterone have been well-studied in response to acute stress and across seasons (Romero, 2002).  OVERVIEW In this thesis, I use a multidisciplinary approach to compare the effects of season and acute restraint stress on corticosterone and DHEA levels in the brain and periphery, and to examine the effects of corticosterone and DHEA treatment on the adult songbird brain. First, to understand the relationship between neurosteroids and acute restraint stress, an effective and reliable method with which to measure steroids in the brain is required. Chapter 2 describes an extensive set of validations for coupling solid phase extraction with radioimmunoassay to measure steroids in songbird brain tissue and plasma. In the subsequent chapters of this thesis, I test three 3 predictions to examine the hypothesis that, in the brain, DHEA and corticosterone are affected by acute stress, and that DHEA mitigates the effects of corticosterone in the brain. 1) Acute stress affects steroid levels differently in jugular versus systemic plasma (Chapter 3). In songbirds, jugular blood is known to be enriched with neurally synthesized steroids (Schlinger and Arnold, 1992; 1993), thus the effects of acute stress on neurosteroid levels may be reflected by changes in jugular steroid levels. Furthermore, season has profound effects of the stress response (Romero et al., 2006; 2008), thus the effects of acute stress on jugular corticosterone and DHEA may depend on season. In wild adult male song sparrows, we compared concentrations of corticosterone and DHEA in plasma collected from the jugular and brachial veins in different seasons. 2) The effects of acute stress on neurosteroid concentrations are dependent on season and brain region (Chapter 4). We measured the effects of season and acute stress on corticosterone and DHEA concentrations in jugular plasma and directly in specific brain regions 5  of wild, adult male song sparrows. Specific areas of the brain are known to be sensitive to stress and elevated glucocorticoids (e.g., hippocampus: McEwen, 1999). We predicted that stress may have a greater effect on neurosteroid concentrations in these sensitive brain areas. 3) DHEA counteracts the effects of corticosterone on neuroanatomy of the songbird brain (Chapter 5). In the dentate gyrus of the adult rat, corticosterone decreased cell proliferation and survival, whereas DHEA increased cell proliferation and survival, and when coadministered, there was no effect (Karishma and Herbert, 2002). However this study used pharmacological levels of DHEA. Using captive nonbreeding adult male song sparrows, we used physiological doses of corticosterone and DHEA, alone or in combination, to examine the effects of these steroids on HVC and hippocampus volume, on neuron number in HVC, and on recruitment of new cells throughout the telencephalon. This is the first study to test the effects of corticosterone treatment on new cell recruitment into HVC.  6  FIGURES  Figure 1.1 Simplified diagram of steroid biosynthesis.  Cholesterol P450scc PREG  P450c17  3β-HSD PROG  P450c17 DHEA  17OH PREG  P450c17  P450c21 11Deoxycorticosterone (DOC) P450c11 Corticosterone 11β-HSD 2 1 Dehydrocorticosterone  3β-HSD 17OH PROG  3β-HSD P450c17  AE  P450c21 11Deoxycortisol  P450aro  E1  17β-HSD T  P450aro  17β-HSD E2  P450c11 Cortisol 11β-HSD 2 1 Cortisone  Figure 1.1 Steroids are shown in bold and enzymes in italics. Steroids: PREG, pregnenolone; 17OH PREG, 17α-hydroxy-pregnenolone; DHEA, dehydroepiandrosterone; PROG, progesterone, 17OH PROG, 17α-hydroxy-progesterone; AE, androstenedione; T, testosterone; E2, 17β-estradiol; E1, estrone. Enzymes: P450scc, cytochrome P450 side chain cleavage; P450c17, cytochrome P450 17α−hydroxylase/17,20 lysase; 3β-HSD, 3β-hydroxysteroid dehydrogenase/isomerase; 17β-HSD, 17β-hydroxysteroid dehydrogenase; P450aro, cytochrome P450 aromatase; P450c21, cytochrome P450 21α-hydroxylase; P450c11, cytochrome P450 11β−hydroxylase; 11β−HSD 1/2, 11β-hydroxysteroid dehydrogenase (type 1 which catalyzes the conversion of dehydrocorticosterone to corticosterone, and type 2 which catalyzes the conversion of corticosterone to dehydrocorticosterone). Adapted from Schmidt et al., 2008.  7  REFERENCES Arvat, E., DiVito, L., Lanfranco, F., Maccario, M., Baffoni, C., Rossetto, R., Aimaretti, G., Camanni, F., Ghigo, E., 2000. Stimulatory effect of adrenocorticotropin on cortisol, aldosterone, and dehydroepiandrosterone secretion in normal humans: dose-response study. J Clin Endocrinol Metab 85, 3141-3146. Buchanan, K.L., Leitner, S., Spencer, K.A., Goldsmith, A.R., Catchpole, C.K., 2004. 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Romero, L.M., Meister, C.J., Cyr, N.E., Kenagy, G.J., Wingfield, J.C., 2008. Seasonal glucocorticoid responses to capture in wild free-living mammals. Am J Physiol Regul Integr Comp Physiol 294, R614-622. Romero, M.L., Cyr, N.E., Romero, R.C., 2006. Corticosterone responses change seasonally in free-living house sparrows (Passer domesticus). Gen Comp Endocrinol 149, 58-65. Romero, M.L., Soma, K.K., Wingfield, J.C., 1998. Hypothalamic-pituitary-adrenal axis changes allow seasonal modulation of corticosterone in a bird. Am J Physiol Regul Integr Comp Physiol 274, R1338-R1344. Sapolsky, R.M., 2000. Glucocorticoids and Hippocampal Atrophy in Neuropsychiatric Disorders. Arch Gen Psychiatry 57, 925-935.  9  Sapolsky, R.M., Romero, L.M., Munck, A.U., 2000. How Do Glucocorticoids Influence Stress Responses? Integrating Permissive, Suppressive, Stimulatory, and Preparative Actions. Endocr Rev 21, 55-89. Schlinger, B.A., Arnold, A.P., 1992. Circulating estrogens in a male songbird originate in the brain. Proc Natl Acad Sci USA 89, 7650-7653. Schlinger, B.A., Arnold, A.P., 1993. Estrogen synthesis in vivo in the adult zebra finch: additional evidence that circulating estrogens can originate in brain. Endocrinology 133, 26102616. Schmidt, K.L., Pradhan, D.S., Shah, A.H., Charlier, T.D., Chin, E.H., Soma, K.K., 2008. Neurosteroids, immunosteroids, and the Balkanization of endocrinology. Gen Comp Endocrinol 157, 266-274. Soma, K.K., Wingfield, J.C., 2001. Dehydroepiandrosterone in songbird plasma: Seasonal regulation and relationship to territorial aggression. Gen Comp Endocrinol 123, 144-155. Soma, K.K., Wissman, A.M., Brenowitz, E.A., Wingfield, J.C., 2002. Dehydroepiandrosterone (DHEA) increases territorial song and the size of an associated brain region in a male songbird. Horm Behav 41, 203-212. Tanapat, P., Hastings, N.B., Rydel, T.A., Galea, L.A.M., Gould, E., 2001. Exposure to fox odor inhibits cell proliferation in the hippocampus of adult rats via an adrenal hormone-dependent mechanism. J Comp Neurol 437, 496-504. Torres, J.M., Ortega, E., 2003. DHEA, PREG and their sulphate derivatives on plasma and brain after CRH and ACTH administration. Neurochem Res 28, 1187-1191. Tramontin, A.D., Brenowitz, E.A., 2000. Seasonal plasticity in the adult brain. Trends Neurosci 23, 251-258. Veiga, S., Garcia-Segura, L.M., Azcoitia, I., 2003. Neuroprotection by the steroids pregnenolone and dehydroepiandrosterone is mediated by the enzyme aromatase. J Neurobiol 56, 398-406. Wolkowitz, O.M., Reus, V.I., Roberts, E., Manfredi, F., Chan, T., Raum, W.J., Ormiston, S., Johnson, R., Canick, J., Brizendine, L., Weingartner, H., 1997. Dehydroepiandrosterone (DHEA) treatment of depression. Biol Psychiatry 41, 311-318. Wingfield, J.C., Sapolsky, R.M., 2003. Reproduction and resistance to stress: When and how. J Neuroendocrinol 15, 711-724. Young, A.H., Gallagher, P., Porter, R.J., 2002. Elevation of the cortisol-dehydroepaindrosterone ratio in drug-free depressed patients. Am J Psychiatry 159, 1237-1239.  10  2  ANALYSIS OF STEROIDS IN SONGBIRD PLASMA AND BRAIN BY COUPLING SOLID PHASE EXTRACTION TO RADIOIMMUNOASSAY 1  INTRODUCTION Analyzing steroid levels via immunoassay is a very common method to determine endocrine status. For example, plasma levels of corticosterone and sex steroids are often measured to evaluate stress physiology (Newman and Soma, 2006; Romero et al., 2006) or reproductive state in songbirds and other species (Williams et al., 2005). In addition to circulating steroids in the plasma, there is accumulating evidence of local steroid synthesis in the brain and other organs (Baulieu, 1997; Soma et al., 2004; Soma et al., 2005; Soma 2006; Demas et al., 2007). Thus it is critical to measure steroid levels within target tissues. When measuring steroids, especially in tissues, sample preparation prior to immunoassays is a crucial step in the analytical process and has a direct impact on the accuracy and precision of the assay (Chard, 1995). A major obstacle in analyzing steroids in tissues is that lipids, such as free fatty acids and triglycerides, can interfere with immunoassays (Rash et al., 1980; Jawad et al., 1981). In particular, brain tissue is rich in lipids, and measuring steroids in egg yolk is also complicated by the high lipid content (Schwabl, 1993). Extremely lipemic plasma samples can also be a problem, especially in animals with a lipid-rich diet, such as seabirds (Speake et al., 1990), carnivores, and nestling birds. Female birds that are depositing yolk into eggs can also have high levels of plasma lipids. Thus, separating steroids in plasma and tissue samples from lipids and other sources of assay interference can be important for accurate and reliable measurements. Traditionally, organic solvents (e.g., dichloromethane (DCM), diethyl ether, hexane) have been used to extract steroids from samples (Samuels, 1947). However, many variables  1  A version of this chapter has been published. Newman, A.E.M., Chin, E.H., Schmidt, K.L., Bond, L., WynneEdwards, K.E. and Soma, K.K. (2008) Analysis of steroids in songbird plasma and brain by coupling solid phase extraction to radioimmunoassay. Gen. Comp. Endocrinol. 155:503-510  11  must be considered when choosing an appropriate extraction technique; these include the steroid being measured, study species, sex, season, reproductive state, and sample type (tissue, plasma, serum). Organic solvent extraction may be appropriate for plasma samples with low lipid content or for immunoassays that require very small amounts of plasma. However, for plasma samples or tissue samples with high lipid content, organic solvent extraction can yield low recoveries (O’Grady, 1968), variable recoveries (Fuqua et al., 1995), or incomplete removal of interfering substances. Importantly, the effectiveness of organic solvent extraction should be determined empirically on a case-by-case basis. Another consideration is that organic solvents used for steroid extraction are often highly reactive or extremely flammable. For example, DCM is an organochlorine with carcinogenic properties, and diethyl ether has high volatility and a low ignition point. Solid phase extraction (SPE) is an alternative to traditional organic solvent extractions (Thurman and Mills, 1998; Telepchak et al., 2004). Previous studies using SPE report high and reliable analyte recovery and more effective removal of interfering substances (Lee and Goeger, 1998). SPE is relatively straightforward to set up in a laboratory and the materials are commercially available. In SPE, a solid sorbent material, typically an alkyl bonded silica, is packed into a column. Extracting relatively non-polar compounds (such as steroids) from a polar matrix (such as water) requires a solid sorbent containing non-polar functional groups of octadecyl (C18) bonded silicas. Steroids bond with C18 groups on the sorbent and are extracted from the sample matrix. Steroids are then eluted from the column. Here we describe validations of SPE to isolate steroids from avian plasma and brain. We address two primary issues: 1) optimization of steroid recovery using radio-labeled and radio-inert steroids and 2) removal of substances that interfere with radioimmunoassay (RIA). Our experiments focus on songbirds, which are widely studied for a variety of research questions about hormonal and environmental influences on physiology and behaviour (Goodson 12  et al., 2005). To our knowledge, measurement of steroid levels in songbird brain has not been previously described.  MATERIALS and METHODS Subjects Plasma pools were created from 1) wild nestling European starlings (Sturnis vulgaris), 2) captive nestling finches (Taeniopygia guttata), and 3) wild adult song sparrows (Melospiza melodia) in non-breeding condition. Plasma from nestling European starlings appeared lipemic upon visual inspection. Brain tissue was collected from wild nestling European starlings, captive adult zebra finches, and wild adult male song sparrows in non-breeding condition. Commercially available chicken serum (Sigma C-540) was used for preliminary validations. Animal use protocols were approved by the Animal Care Committee at the University of British Columbia and conform to the guidelines promulgated by the Canadian Council on Animal Care.  Reagents Radioinert dehydroepiandrosterone (DHEA), corticosterone (CORT), and 17β-estradiol (E2) were purchased from Steraloids (Newport, Rhode Island). Radiolabeled steroids were purchased from Perkin-Elmer (3H-DHEA, NET814; 3H- E2, NET517). HPLC-grade methanol (MeOH) and HPLC-grade dichloromethane (DCM) were purchased from Fisher Scientific. Absolute ethanol (EtOH) was obtained from the UBC Chemistry Department.  SPE Protocol The SPE protocol involves 7 major steps: 1) Solvation, columns are primed with ethanol, 2) Equilibration, water is passed through columns to prepare sorbent for sample loading, 3) Sample Loading, samples in aqueous matrix are passed through the columns, 4) Interference Elution, 13  water is passed through columns to wash out interfering polar substances, 5) Sample Elution, steroids are eluted from columns with a small amount of eluant into 7mL glass vials, 6) Drying, eluates are dried, and 7) Resuspension, dried eluates are resuspended in assay buffer. For SPE, we used a 24-place vacuum manifold (United Chemical Technologies, UCTVMF024GL, approx $930 cdn), non-endcapped C18 columns (500mg C18 material, 6mL column volume, UCTCUC18156, approx. $2 cdn/column), and polytetrafluoroethylene tips. We systematically manipulated several variables after sample loading to evaluate effects on 1) steroid recovery and 2) removal of assay interference. To measure steroid recovery, plasma and brain samples were spiked with known quantities of tritiated steroids. A known amount of tritiated steroid was added to samples prior to loading samples onto columns. To evaluate loss of tritiated steroids during the process, aliquots were counted in a liquid scintillation counter (Beckman Coulter LS-6500) for 2 min. After initial studies using chicken serum spiked with 3H-DHEA or 3H-E2, the protocol was applied to song sparrow brain tissue spiked with either 3H-DHEA or 3H-E2. In addition, steroid recovery was evaluated by spiking samples with radioinert steroids prior to extraction and measuring extracted samples using RIA. By comparing unspiked and spiked samples, we were able to calculate recovery of exogenous radioinert steroids.  Eluting Solvent Choosing the appropriate solvent for elution is important in maximizing extraction efficiency. We compared solvents (5mL, all HPLC-grade): methanol, 90% MeOH (9:1 MeOH:dH2O), and DCM. These solvents are common in SPE (MeOH) or traditional organic solvent extraction methods (DCM).  14  Drying Method Following sample elution, the eluates are dried prior to resuspension in assay buffer. We compared recovery efficiency when eluates were dried under air or nitrogen (N2). To dry eluates under air, the vacuum manifold was used to pull a steady stream of air through the columns and over the eluates at room temperature. To dry eluates under N2, samples were placed in a water bath at 38˚C under a steady stream of medical-grade N2.  Resuspension Volume Resuspending dried eluates prior to immunoassay is an important step that greatly affects recovery. Steroids have low solubility in aqueous buffers and may adhere to glass vials after drying. To determine whether the volume of resuspension buffer affects recovery, two volumes of phosphate buffered saline with gelatin (PBSG) were added to dried eluates: 220μl or 440μl. These volumes are sufficient for one or two immunoassays depending on expected steroid concentrations and assay sensitivity.  Ethanol in Resuspension We also determined whether EtOH aids in resuspending steroids. Using high-grade absolute ethanol, we compared the effects of 0, 5, and 10% EtOH in the resuspension buffer on recovery of radiolabeled steroids from dried eluates. The absolute EtOH was added directly to the bottom of the glass vials containing dried eluates, the vials were vortexed briefly, the assay buffer was added, and the vials were vortexed (2 min). An important concern is whether EtOH in the resuspension buffer affects the immunoassay (e.g., by affecting the primary or secondary antibodies). To address this issue, we determined whether 5 or 10% EtOH had an effect on the measured concentrations of known amounts of steroids. 15  Resuspension Mixing Method We determined whether a manual vortexer (VWR, 58816-121) or a rack shaker (IKA Vibrax VWR basic) affected recovery of radiolabeled steroids. Resuspension buffer was added to dried eluates, vials were either vortexed (4 min) or shaken (60 min), stored at 4˚C overnight, and either vortexed (4 min) or shaken (60 min) again.  Radioimmunassays We examined recovery of exogenous radioinert DHEA, CORT and E2 via RIA. Known quantities of radio-inert DHEA, CORT or E2 were added to samples prior to SPE. We calculated recovery by comparing the quantity of steroids in unspiked and spiked samples. To determine whether SPE effectively removed substances that interfere with the RIAs, we serially diluted plasma and brain samples after SPE. If interfering substances are removed, then serially diluted samples should be parallel to the standard curve, when comparing the percentage of tracer bound (Chard, 1995). To measure DHEA, we used a specific double antibody RIA (DSL 8900, Diagnostic Systems Laboratories, Webster, TX). The DHEA RIA was modified according to Granger et al (1999) to increase sensitivity and has been previously used for songbird plasma (Goodson et al., 2005; Soma, 2006). To measure E2, we used a specific double antibody RIA (DSL 39100, Diagnostic Systems Laboratories, Webster, TX) which was modified according to Shirtcliff et al. (2000). To measure CORT, we used a specific double antibody RIA (ImmuChem 07-120103, MP Biomedicals, Orangeburg, NY) which was modified according to Washburn et al. (2002).  16  Statistical Analysis To determine parallelism between serial dilutions of plasma and brain tissue with the standard curve, we tested equality of slopes using an analysis of covariance (ANCOVA). A lack of significant interaction between serial dilution and standard curve indicates that the line slopes are similar. If the interaction term is significant, then the slopes of the serially diluted sample and the standard curve are not parallel. We used JMP IN 5.1 (SAS, Cary, NC) for linear models and Sigma Stat 3.0 (Jandel Scientific, San Rafael, CA) for between group comparisons. We considered test results significant for p< 0.05. Means ± standard errors represent central tendency and variability, respectively.  RESULTS Recovery of radiolabeled steroids The highest recoveries of radiolabeled DHEA and E2 from chicken serum were obtained when samples were eluted with 90% MeOH and then resuspended in buffer with 10% EtOH, although the difference between 5% and 10% EtOH in resuspension buffer was slight (Table 1). Resuspending dried extracts in either 220μl or 440μl resuspension matrix did not alter the recovery of 3H-DHEA (Table 1). Drying eluates under N2 at 38˚C, particularly with 5 or 10% EtOH in the resuspension buffer, significantly increased recovery of radiolabeled DHEA and E2 (Table 2). Resuspending dried eluates with either a manual vortexer or automatic shaker did not affect recovery of radiolabeled DHEA when EtOH was added to the resuspension buffer (5% EtOH: t-test, t=-0.49, p=0.64; 10% EtOH: t-test, t=-0.13, p=0.90; data not shown). However, if no EtOH was added to the resuspension buffer, recovery increased significantly from 46 ± 1.4% to 56 ± 3.9% with the rack shaker (t-test, t= -2.51, p=0.04).  17  Lastly, highest recovery of radiolabeled DHEA from songbird brain tissue was 68.9 ± 0.8% and was achieved with 5% EtOH in resuspension buffer (ANOVA, F(2, 11)=4.27, p=0.05). Highest recovery of radiolabeled E2 from songbird brain tissue was 66.6 ± 2.4% and was unaffected by percentage of EtOH in the resuspension buffer (ANOVA, F(2,11)=0.007, p=0.9).  Recovery of Radioinert Steroids We also determined recovery of known amounts of radioinert steroids that were added to samples prior to SPE and then measured via RIA (n=4 replicates for each trial). Except where stated otherwise, 5% EtOH was used in the resuspension of dried eluates. For DHEA, recovery of 50pg DHEA added to chicken serum was 113% with 5% EtOH in resuspension and 120% with 10% EtOH in resuspension. Recovery of 50pg DHEA added to nestling starling plasma was 104%. Recovery of 25pg DHEA added to 0.5mg song sparrow brain tissue was 80% and recovery of 50pg DHEA added to 1mg song sparrow brain tissue was 72%. Recovery of 0.5pg 17ß-E2 added to nestling starling plasma was 107%. Recovery of 1pg 17ß-E2 added to nestling starling brain tissue was 62%. For CORT, recovery of 25pg added to nestling starling plasma and brain tissue were 86% and 71%, respectively. We also determined the effect of absolute EtOH in the resuspension buffer on RIAs. For DHEA RIA, 5% and 10% EtOH in the resuspension buffer did not affect the concentration of DHEA in known controls (Fig 1). Similarly, for CORT or E2 RIA, when compared with 0%, 5% EtOH in the resuspension buffer did not affect the concentration of CORT in a 125pg control (0% EtOH: 122.5 ± 4.07pg; 5% EtOH: 123.2 ± 3.58pg) or E2 in known controls (data not shown).  18  Removal of interfering substances To determine whether SPE effectively removed interfering substances, such as lipids, from plasma and brain samples, we compared serially diluted samples and standard curves. For DHEA, we examined plasma and brain tissue that were not extracted by either organic solvents or SPE. Without extraction, serially diluted song sparrow plasma and brain tissue yielded displacement curves that were not parallel to the standard curve (data not shown; Plasma: ANCOVA, R2=0.96, significant interaction, F2,15 = 102.31, p = 0.007; Brain Tissue: ANCOVA, R2 = 0.98, significant interaction, F2, 13 = 244.98, p < 0.0001). Indeed, the serially diluted brain tissue yielded a flat displacement curve. Second, we examined the effects of DCM extraction. With DCM extraction, serial dilutions of nestling zebra finch plasma, nestling European starling plasma, and song sparrow brain tissue yielded displacement curves that were not parallel to the DHEA standard curve (Fig 2.2A, Brain Tissue: R2=0.96, significant interaction, F2,13=28.87, p=0.0002; Plasma: F2,13= ANCOVA, R2=0.97, significant interaction, F2,18=22.18, p<0.0001). Third, we evaluated the efficacy of other organic solvent extractions. Song sparrow plasma extracted with diethyl ether or hexane/ethyl acetate (9:1) also yielded displacement curves that were not parallel to the DHEA standard curve (data not shown; ANCOVA, R2=0.96, significant interaction, F3, 20 = 13.87, p = 0.0004). In contrast, with SPE, serially diluted nestling European starling plasma and song sparrow brain tissue yielded displacement curves that were parallel to the DHEA standard curve (Fig 2.2B, Brain Tissue: R2= 0.98, no significant interaction, F2, 13 = 0.58, p = 0.46; Plasma: ANCOVA, R2=0.98, no significant interaction; F2,10 = 0.04, p = 0. 84). Similarly, for E2 RIA, serially diluted adult zebra finch brain extracted with SPE was parallel to the E2 standard curve (Fig 2.2C, ANCOVA, R2=0.56, no significant interaction, F1,8=4.3, p=0.1,). Lastly, for CORT  19  RIA, serially diluted adult song sparrow brain extracted with SPE was parallel to the CORT standard curve (Fig 2.2D, ANCOVA, R2=0.99, no significant interaction, F1,10 = 3.84, p = 0.10). DISCUSSION Extraction of steroids from lipid-rich samples has been an obstacle for analyzing steroids in lipemic plasma and tissue. These results indicate that solid phase extraction (SPE) is a suitable method for extracting steroids from lipid-rich avian plasma and brain samples. SPE is a straightforward procedure that can easily be established in a laboratory (Fig 3). SPE yields relatively high and consistent recovery of radiolabeled and radioinert steroids. After SPE, serially diluted plasma and brain samples yield displacement curves that are parallel to the standard curve. Interestingly, steroid levels in songbird brain have not been previously described. This method provides a reliable way to quantify steroids in small tissues or tissues that are rich in interfering substances. Extraction techniques can be examined using radiolabeled steroids, serially diluted samples, and radioinert steroids. Adding known quantities of radiolabeled steroids to samples prior to extraction provides information on recovery, but does not indicate if the extraction removes substances that interfere with immunoassay. Examining parallelism between serially diluted samples and a standard curve reveals the ability of the extraction to remove interfering substances, but does not indicate recovery efficiency. Serial dilutions are also useful in determining an appropriate sample volume (Chard, 1995). Lastly, calculating recovery of known amounts of radioinert steroids added prior to extraction and measured with immunoassay provides information on both recovery and removal of interfering substances. Eluting the steroids from C18 columns with 90% HPLC-grade MeOH in deionized water yields the highest recovery of radiolabeled steroids. Methanol forms hydrogen bonds with the silica surface and breaks the van der Waals interactions between the steroids and the C18 material. Thus, it is an effective solvent for eluting non-polar analytes, such as steroids. 20  Adjusting the solvent can affect which analytes are eluted. For example, eluting with a lower percentage of MeOH in water can specifically elute sulfated steroids, but not unconjugated steroids (Liere et al., 2004). Other non-polar solvents, such as acetonitrile and ethyl acetate, may also work well for non-polar, lipophilic analytes and should be examined in future studies. Drying the eluates is necessary prior to resuspension in assay buffer. Drying the eluates under a steady stream of medical-grade N2 yields significantly higher recovery of radiolabeled steroids than drying under a stream of air. Importantly, the N2 is free of possible contaminating substances that may be present in air, and inert N2 is less likely to cause steroid oxidation. Furthermore, drying under N2 at 38˚C is much faster than drying at room temperature. Additionally, eluates can be dried using a vacuum concentrator, and future studies will examine this method. After drying, dried eluates can be stored at -20ºC prior to resuspension in assay buffer. An addition of absolute EtOH in the resuspension, either 5 or 10%, significantly improves recovery of radiolabeled steroids. The EtOH is added directly to the dried eluates at the bottom of the vials and then brought up to final volume with assay buffer. In our laboratories, we routinely use 5% EtOH in resuspension buffer, which does not affect the DHEA, E2 or CORT RIAs in our laboratories. However, the effect of EtOH should be determined for each RIA or EIA used. These data also demonstrate that SPE effectively reduces assay interference in songbird samples for measurement of DHEA, E2, and CORT. For other analytes, it is possible that a second interference elution with dilute MeOH or another eluant will be important to further reduce assay interference (e.g., Vallee et al 2000). Further work on the interference elution(s) is a goal for future experiments. While SPE can separate unconjugated steroids from sulfated steroids (Liere et al., 2004), SPE has typically not been used to separate various unconjugated steroids from each other. In 21  this regard, SPE C18 columns generally have been used differently than Celite chromatography columns, which can separate several unconjugated steroids (Wingfield and Farner, 1975; Soma et al., 1999). Note that C18 columns can be used with HPLC to separate unconjugated steroids, but HPLC C18 columns are much longer than SPE C18 columns and thus have greater separating ability. Whether SPE C18 columns can be used to separate unconjugated steroids from each other is an important question for future studies. These data demonstrate that SPE is useful prior to RIA, but SPE is also useful prior to EIA (Acosta et al., 2000; Love et al., 2005; Williams et al., 2005), high-performance liquid chromatography (Hojo et al., 2004; Ziegler and Wittwer, 2005), and gas chromatograph-mass spectrometry (Mensah-Nyagan et al., 1998; Vallee et al., 2000). SPE is an attractive alternative to traditional organic solvent extractions, because recoveries are high and consistent, and removal of assay interference is more complete. The equipment and columns are commercially available, and set-up is straightforward. Also, it may be possible to reduce the cost of SPE by washing and reusing the C18 columns. These data highlight the importance of empirically determining how to prepare samples prior to steroid measurement. This should be done on a case-by-case basis. Many variables can affect the choice of sample preparation technique: the steroid measured, study species, sex, season, reproductive state, and sample type (e.g., tissue, plasma, serum). First, in some cases, no extraction may be needed. For example, Washburn et al (2002) demonstrate that CORT can be measured in avian plasma without extraction, when using a particular CORT RIA. Second, in other cases, an organic solvent extraction may be sufficient. For example, Wingfield and Farner (1975) demonstrate that T can be measured in avian plasma after DCM extraction, especially when DCM extraction is followed by Celite column chromatography. In addition, Tsutsui and Yamazaki (1995) measured pregnenolone and DHEA in Japanese quail brain after ethyl acetate extraction. Third, in other cases, SPE may be required. The present results 22  indicate that SPE is more effective than organic solvent extractions for removal of interfering substances from songbird brain and lipemic plasma. Migues et al (2002) measured DHEA in chicken brain after SPE, but to our knowledge, the present report is the first examination of steroid levels in the brain of songbirds, an important model system in neuroendocrinology. Also, to compare steroid levels in tissues and plasma, both types of samples should be processed similarly. That is, if SPE is required for tissue but not plasma samples, both types of samples should be prepared using SPE to avoid a confound. SPE should prove useful for a variety of researchers interested in measuring steroids in plasma, serum, brain, yolk, feces, urine and whole body homogenates from a wide range of organisms.  23  TABLES Table 2.1 Recovery of 3H-DHEA and 3H-E2 from chicken serum using solid phase extraction. % Ethanol in Resuspension Resuspenion Volume (μl) 220  0%  5%  10%  45.3 ± 2.2  72.8 ± 1.9  80.0 ± 2.2  440  65.8 ± 1.6  72.7 ± 6.2  81.2 ± 3.0  DHEA: 100% Methanol  220  70.9 ± 2.6  70.7 ± 2.2  78.1 ± 0.1  440  65.5 ± 1.0  67.3 ± 5.0  70.6 ± 7.5  DHEA: Dichloromethane  220  40.3 ± 2.9  67.3 ± 0.8  65.1 ± 1.7  440  37.8 ± 3.6  62.1± 3.4  68.5 ± 1.4  220  65.9 ± 1.6  73.9 ± 2.1  80.6 ± 0.7  220  46.6 ± 1.3  52.1 ± 0.9  56.2 ± 1.1  Steroid: Eluant DHEA: 90% Methanol  Estradiol: 90% Methanol Estradiol: Dichloromethane  Note: Data are means ± SEM and n = 4 for each group.  24  Table 2.2 Recovery of 3H-DHEA and 3H-E2 from chicken serum extracts dried under N2 or under air after solid phase extraction. N2 at 38˚C % Ethanol in resuspension 0% 5% 10%  Air at 22˚C  DHEA  E2  DHEA  E2  45.2 ± 2.2 72.8 ± 1.9 80.3 ± 2.2  65.9 ± 1.6 73.9 ± 2.1 80.6 ± 0.7  50.0 ± 3.3 56.2 ± 2.9 60.2 ± 1.0  50.0 ± 1.7 53.2 ± 1.5 58.2 ± 1.0  Note: Data are means ± SEM and n = 4 for each group.  25  FIGURES  Figure 2.1 Effects of ethanol on the DHEA radioimmunoassay.  1000 Standard A Standard B  DHEA (pg/mL)  800  600  400  200  0 0%  5%  10 %  % Ethanol  Figure 2.1 Adding absolute EtOH to the resuspension buffer (either 5 or 10% of total resuspension volume) did not change the concentration of DHEA in known standards measured with RIA. Data are means ± standard error.  26  Figure 2.2 Comparing Dichloromethane Extraction and Solid Phase Extraction.  Dichloromethane Extraction  A  100  80  80  60  60  % Bound  % Bound  Solid Phase Extraction (DHEA)  B  100  40 DHEA Standard Curve European starling plasma Zebra finch plasma Song sparrow brain tissue  20  40 DHEA Standard Curve European starling Plasma Song sparrow brain tissue  20  0  0 0.1  1  10  100  1000  0.1  DHEA (pg) or Sample (µl or mg)  % Bound  120  Solid Phase Extraction (E2)  100  1000  100  100  80  80  60  60  40  40  20 Estradiol Standard Curve Zebra finch brain tissue  20 0.01  10  Solid Phase Extraction (CORT)  D  % Bound  C  1  DHEA (pg) or Sample (µl or mg)  CORT Standard Curve Song sparrow brain tissue  0 0.1  1  10  Estradiol (pg) or Sample (µl or mg)  100  0.1  1  10  100  1000  CORT (ng) or Sample (µl or mg)  Figure 2.2 Solid phase extraction (SPE), but not dichloromethane (DCM) extraction, effectively removed substances that interfere with DHEA RIA. A) Serially diluted samples of nestling European starling plasma, nestling zebra finch plasma, and song sparrow brain tissue extracted with DCM are not parallel to the DHEA standard curve. For DCM extraction, steroids were extracted with 3mL HPLC-grade DCM (twice), and extracts were dried under N2 and resuspended in assay buffer. B) Serially diluted samples of nestling European starling plasma and song sparrow brain tissue extracted with SPE are parallel to the DHEA standard curve. C) Serially diluted zebra finch brain tissue extracted with SPE is parallel to the E2 standard curve. D) Serially diluted song sparrow brain tissue extracted with SPE is parallel to the CORT standard curve.  27  Figure 2.3 Solid phase extraction protocol for songbird plasma and brain tissue. Tissue Preparation  Plasma Preparation  SPE Protocol  Day One Weigh tissue, transfer to 13 x 100 mm test tube  Thaw plasma on ice  e.g. weight = 25 mg Add ice-cold dH2O (3x tissue mass) e.g. 25 mg x 3 = 75 µL  Measure plasma into 16 mm test tube  Add 10 mL dH2O to test tube  Homogenize tissue on ice  Open valves to allow EtOH to flow through columns until 1 mm remains, then close valves  Sonicate 15 min at room temp SPE  Resuspension: One day prior to assay, add absolute EtOH (5% final conc.) directly onto bottom of vial, vortex 1 sec, add buffer, vortex 3 sec Shake at ~ 2000 rpm for 1 hr  Equilibration: Add 5 mL dH2O to columns (twice) Open valves to allow dH2O to flow through columns until 1 mm remains, then close valves  Add HPLC grade MeOH (4x homogenate vol) e.g. 100 µL x 4 = 400 µL  Shake at 1000 rpm for 1 hr at room temp  Solvation: Add 3 mL HPLC EtOH to columns & soak for 2 min  Sample Loading: Add ~ 5 mL of sample (see left) to columns (twice)  Store at 4 °C overnight  Shake ~2000 rpm for 1 hr  Assay  Open valves to allow sample to flow through columns until 1 mm remains, then close valves  Store overnight at 4°C Interference Elution: Add 5 mL dH2O to columns (twice)  Day Two Shake at 1000 rpm for 1 hr at room temp Centrifuge at 3000 g for 10 min at 2 °C Transfer supernatant, (up to 1 mL) to a 16 mm glass test tube Add 10 mL dH2O to test tube  SPE  After second interference elution, open all valves for 5 min to remove excess dH2O, then close valves. Insert clean 7 mL glass vials into vacuum manifold Sample Elution: Add 5 mL 90% MeOH (HPLC-grade) to columns, soak for 2 min Use vacuum to prompt MeOH to flow, then turn off vacuum and wait for all of MeOH to drip slowly through columns, then turn on vacuum for 2 min to collect remaining MeOH Sample Drying: Dry eluates under N2 at ~40°C then store at –20°C, or proceed to resuspension  Figure 2.3 Summary of solid phase extraction protocol for tissue and plasma samples. C18 columns should be closed prior to addition of liquids for all steps. After liquid has been gently added to the column, columns are opened and liquid is allowed to drip into collecting vessel. If necessary, vacuum pressure may be used to begin dripping, but dripping should be continued without vacuum pressure to ensure a slow and constant drip rate. During SPE, it is crucial that a small amount of liquid (approximately 1mm above packing material) remains in the column between steps unless otherwise stated (for details see Thurman and Mills, 1998). 28  REFERENCES Acosta, T.J., Ozawa, T., Kobayashi, S., Hayashi, K., Ohtani, M., Kraetzl, W.D., Sato, K., Schams, D., Miyamoto., A. 2000. Periovulatory changes in the local release of vasoactive peptides, prostaglandin F2α, and steroid hormones from bovine mature follicles in vivo. Biol Reprod 63, 1253-1261. Baulieu, E.E., 1997. Neurosteroids: of the nervous system, by the nervous system, for the nervous system. Recent Prog Horm Res 53, 1-32. Chard, T., 1995. An Introduction to Radioimmunoassay and Related Techniques. Elsevier, New York. Demas, G.E., Cooper, M.A., Albers, H.E., Soma, K.K., 2007. Novel mechanisms underlying neuroendocrine regulation of aggression: a synthesis of rodent, avian and primate studies. In: Blaustein, J.D. (ed), Behavioral Neurochemistry and Neuroendocrinology. 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Clin Chem 26, 84-88. Romero, L.M., Cyr, N.E., Romero, R.C., 2006. Corticosterone responses change seasonally in free-living house sparrows (Passer domesticus). Gen Comp Endocrinol 149, 58-65. Samuels, L.T., 1949. Metabolism of steroids by tissues. I. Determination of testosterone and related steroids in tissue extracts. J Biol Chem 168, 471-475. Schwabl, H. 1995. Yolk is a source of maternal testosterone for developing birds. Proc Natl Acad Sci USA 90, 11446-11450. Shirtcliff, E.A., Granger, D.A., Schwartz, E.B., Curran, M.J., Booth, A., Overman, W.H., 2000. Assessing estradiol in biobehavioral studies using saliva and blood spots: simple radioimmunoassay protocols, reliability, and comparative validity. Horm Behav 38, 137-147. Soma, K.K., 2006. Testosterone and aggression: Berthold, birds and beyond. J Neuroendocrinol 18, 543-551. Soma, K.K., Alday, N.A., Hau, M., Schlinger, B.A., 2004. Dehydroepiandrosterone metabolism by 3β-hydroxysteroid dehydrogenase/Δ5-Δ4 isomerase in adult zebra finch brain: sex difference and rapid effect of stress. Endocrinology 145, 1668-1677. Soma, K.K., Sinchak, K., Lakhter, A., Schlinger, B.A., Micevych, P.E., 2005. Neurosteroids and female reproduction: estrogen increases 3β-HSD mRNA and activity in rat hypothalamus. Endocrinology 146, 4386-4390. Soma, K.K., Sullivan, K., Wingfield, J.C., 1999. Combined aromatase inhibitor and antiandrogen treatment decreases territorial aggression in a wild songbird during the nonbreeding season. Gen Comp Endocrinol 115, 442-453.  30  Soma, K.K., Wingfield, J.C., 2001. Dehydroepiandrosterone in songbird plasma: seasonal regulation and relationship to territorial aggression. Gen Comp Endocrinol 123, 144-155. Speake, B.K., Decrock, F., Surai P.F., Groscolas, R., 1999. Fatty acid composition of the adipose tissue and yolk lipids of a bird with a marine-based diet, the emperor penquin (Aptenodytes fosteri). Lipids 34, 283-290. Telepchak., M.J., August, T.F., Chaney, G., 2004. Forensic and Clinical Applications of Solid Phase Extraction. Humana Press, New Jersey. Thurman, E.M., Mills, M.S., 1998. Solid-phase Extraction: Principles and Practice. John Wiley & Sons Inc., New York. Tsutsui, K., Yamazaki, T., 1995. Avian neurosteroids. I. Pregnenolone biosynthesis in the quail brain. Brain Res 678, 1-9. Vallee, M., Rivera, J.D., Koob, G.F., Purdy, R.H., Fitzgerald, R.L., 2000. Quantification of neurosteroids in rat plasma and brain following swim stress and allopregnanolone administration using negative chemical ionization gas chromatography/mass spectrometry. Anal Biochem 287, 153-166. Washburn, B.E., Morris, D.L., Millspaugh, J.J., Faaborg, J., Schulz, J.H., 2002. Using a commercially available radioimmunoassay to quantify corticosterone in avian plasma. The Condor 104, 558-563. Williams, T.D., Ames, C.E., Kiparissis, Y., Wynne-Edwards, K.E., 2005. Laying-sequencespecific variation in yolk oestrogen levels, and relationship to plasma oestrogen in female zebra finches (Taeniopygia guttata). Proc Roy Sci B 272, 173-177. Wingfield, J.C., Farner, D.S., 1975. The determination of five steroids in avian plasma by radioimmunoassay and competitive protein-binding. Steroids 26, 311-321. Ziegler, T.E., Wittwer, D.J., 2005. Fecal steroid research in the field and laboratory: improved methods for storage, transport, processing, and analysis. Am J Primatol 67, 159-174.  31  3  DHEA AND CORTICOSTERONE ARE REGULATED BY SEASON AND ACUTE STRESS IN A WILD SONGBIRD: JUGULAR VERSUS BRACHIAL PLASMA 2  INTRODUCTION It is well-known that acute stress stimulates the hypothalamic-pituitary-adrenal (HPA) axis and results in glucocorticoid release from the adrenal glands in vertebrates. Interestingly, the HPA axis is plastic in adulthood and varies dramatically across the seasons (Pyter et al., 2007; Romero, 2002; Romero et al., 1998). Seasonal variation in HPA axis function may enable animals to adjust behavior and physiology to match seasonal changes in food, weather, predators, or social organization (Sapolsky et al., 2000). Like glucocorticoids, dehydroepiandrosterone (DHEA) is regulated by stress. DHEA is a sex steroid precursor with no known classical intracellular steroid receptor (Labrie et al., 2005; Widstrom and Dillon, 2004). In humans, DHEA and its sulfated ester, DHEA-S, are the most abundant steroids secreted by the adrenal cortex. Acute stress and adrenocorticotrophic hormone (ACTH) increase circulating DHEA levels in humans (Kroboth et al., 1999; Oberbeck et al., 1998; Salek et al., 2002) and DHEA is more sensitive than cortisol to low doses of ACTH (Arvat et al., 2000). Nonetheless, the function of DHEA in the stress response is not well understood. Importantly, DHEA is not only produced by the adrenals, but can also be synthesized by the gonads and the brain itself (Baulieu, 1998;Robel and Baulieu, 1995). In rats and mice, plasma DHEA levels are low, and the adrenals produce little or no DHEA (Labrie et al., 2005). Concentrations of DHEA are higher in the rat brain than plasma, even after adrenalectomy (Corpechot et al., 1981), and the brain expresses the necessary steroidogenic enzymes (Mensah-  2  A version of this chapter has been published. Newman, A.E.M., Pradhan, D.S., and Soma, K.K. (2008) DHEA and corticosterone are regulated by acute stress in a wild songbird: jugular versus brachial plasma. Endocrinology 149: 2537-2545.  32  Nyagan et al., 1999). Acute stress and ACTH treatment increase DHEA levels in the rat brain (Corpechot et al., 1981; Torres et al., 2001; Torres and Ortega, 2003). These data suggest that DHEA, as a neurosteroid, is regulated by stress. Recent evidence suggests a role for DHEA as an anti-glucocorticoid in the nervous system (Kalimi et al., 1994). For example, in humans, the ratio of cortisol to DHEA is increased in some studies of patients with stress-related psychiatric diseases, such as major depression and schizophrenia (Ritsner et al., 2004; Young et al., 2002). Further, DHEA treatment to depressed patients improves mood (Schmidt et al., 2005; Wolkowitz et al., 1997). In rats, DHEA is a potent anti-glucocorticoid in neuronal cultures (Kimonides et al., 1999) and protects against glucocorticoid-induced neuronal death in vivo (Karishma and Herbert, 2002). The mechanisms of action remain unclear because DHEA has low affinity for mineralocorticoid receptors and glucocorticoid receptors (Chen et al., 2005), but neural metabolism of DHEA to active sex steroids might be important (Hajszan et al., 2004). Here, we examined the effects of acute restraint on plasma DHEA and corticosterone (CORT) levels in a songbird, the song sparrow (Melospiza melodia). We measured DHEA and CORT in different seasons, because there are large seasonal changes in both steroids (Romero, 2002; Soma and Wingfield, 2001). Song sparrows, unlike rats and mice, have relatively high levels of DHEA in the plasma (Soma and Wingfield, 2001). In the nonbreeding season, DHEA levels are elevated in song sparrow adrenal glands (Soma, 2006), and recent evidence indicates high DHEA levels in the brain (Newman and Soma, 2007). A physiological dose of DHEA has significant effects on behavior and neuroanatomy in song sparrows (Soma et al., 2002). Last, DHEA metabolism to sex steroids in the songbird brain is very high (Pradhan et al., 2008; Schlinger et al., 2008; Soma et al., 2004). Neural metabolism of DHEA by 3β-hydroxysteroid dehydrogenase/isomerase (3β-HSD) is rapidly increased by restraint in breeding male song sparrows (Soma et al., 2002). 33  We measured DHEA and CORT in plasma from the brachial vein and jugular vein. Brachial plasma serves as a measure of systemic steroid levels, and jugular plasma serves as an indirect measure of the neural steroidal milieu. Importantly, in songbirds, jugular plasma is enriched with neurally-synthesized steroids, such as estradiol (Schlinger and Arnold, 1992). When radiolabeled androgen is administered peripherally, radiolabeled estrogens are higher in jugular plasma than carotid plasma (Saldanha and Schlinger, 1997; Schlinger and Arnold, 1992; Schlinger and Arnold, 1993). Recent studies suggest that the songbird brain can synthesize DHEA (London et al., 2006). If the brain is the main source of circulating DHEA, then jugular DHEA will be higher than brachial DHEA. During the nonbreeding season (when the gonads are regressed), the brain might be the main source of circulating DHEA (Soma and Wingfield, 2001). Thus, a difference between jugular and brachial DHEA levels could be season-specific. Last, if DHEA or its metabolites have anti-glucocorticoid properties in the brain, acute restraint stress might specifically alter the balance between neural DHEA synthesis and metabolism, thereby affecting jugular DHEA levels.  MATERIALS and METHODS Subjects Subjects were wild adult male song sparrows (n = 107 total). Song sparrows are a useful model system, because their behavior and physiology in the wild are well-characterized (Nice, 1937; Wingfield, 1994). Subjects were captured near Vancouver, British Columbia (49˚ 12’N, 123˚ 01’W). This population is sedentary, and males maintain territories year-round (Arcese et al., 2002; Wingfield, 1994). In 2005, individuals were captured during 4 seasons: 1) Breeding (May 1-25, n = 34); 2) Molt (Aug 15-Sept 8, n = 19); 3) Early nonbreeding (Oct 10-27, n = 38); and 4) Mid-nonbreeding (Dec 19-23, n = 16). During breeding, plasma testosterone levels are elevated, and males aggressively defend territories. During the molt, song sparrows replace 34  their feathers, have non-detectable plasma testosterone levels, and show little territorial aggression. During the nonbreeding season, the testes are regressed, plasma testosterone levels are non-detectable, but territorial aggression is expressed at high levels (Wingfield and Hahn, 1994). Protocols were approved by the University of British Columbia Committee on Animal Care and complied with the guidelines of the Canadian Council of Animal Care.  Field Protocol Subjects were caught using mist-nets and less than 5 min of conspecific song playback (mean ± SEM: 2.78 ± 0.02 min). A baseline blood sample (~150 μl) was collected within 3 min of capture (mean ± SEM: 2.2 ± 0.05 min) from the brachial or jugular vein (see below). Then, subjects were restrained in an opaque cloth bag for 30 or 60 min. After restraint, another blood sample (~150 μl) was collected (from the same vein as at baseline). Importantly, separate individuals were bled after 30 min restraint and 60 min in 2 of 4 seasons (breeding and early nonbreeding seasons). From the brachial vein, blood was collected with heparinized micro-hematocrit tubes after venipuncture with a sterile 26-gauge needle. From the jugular vein, blood was collected with a sterile heparinized 1 mL syringe with a fixed 28-gauge needle, as described previously (Hoysak and Weatherhead, 1991; Sheldon et al., 2008; Silverin et al., 2004). We used cotton and gentle pressure to stop the blood flow. Blood was kept on ice until returned to the laboratory (2-8 hr) and centrifuged. Plasma was stored at –20˚C. We also collected the following body measurements: length (to nearest 0.1 mm) of the tarsus, wing, and cloacal protuberance (androgen-dependent secondary sex characteristic); abdominal and furcular fat scores (5-level visual fat index (Helms and Drury, 1960)); and body mass (to the nearest 0.1 g). Last, subjects were given a unique combination of 3 plastic colour bands and a numbered aluminum band. Subjects were then released back onto their territory. 35  Corticosterone Radioimmunoassay To measure CORT, we used a double antibody 125I-radioimmunoassay (RIA) (ImmuChem 07120103, MP Biomedicals, Orangeburg, NY) which was modified for songbird plasma (Washburn et al., 2002). Each sample was measured in duplicate. Briefly, the manufacturer’s directions were followed, except that the volumes of all reagents were halved and plasma was diluted 1:50 (5 μl of plasma + 245 μl of assay buffer). The CORT antibody has a low crossreactivity with desoxycorticosterone (0.35%), testosterone (0.10%), cortisol (0.05%), androstenedione (0.03%), and DHEA (<0.01%). The intra-assay variation was 3.8% (n = 12 replicates in 1 assay), and the inter-assay variation was 2.4% for the low standard and 2.1% for the high standard (n = 4 assays). The lowest point on the standard curve was 3.1 pg CORT/tube, and the detection limit for our assays with 1 µL plasma/tube was 3.1 ng/mL. We validated this assay for song sparrow plasma. Using a plasma pool, we compared the CORT concentrations in samples assayed without a steroid extraction step and in samples assayed after a steroid extraction step. For steroid extraction, we used HPLC-grade dichloromethane (DCM) (3 mL x2). Extracts were dried under N2 at 40°C, resuspended in 250 μl assay buffer, and stored overnight at 4°C prior to RIA. We also examined parallelism between the standard curve and serially-diluted unextracted plasma. Parallelism between the two curves indicates that substances in the plasma do not interfere with the assay (Chard, 1995).  Dehydroepiandrosterone Radioimmunoassay To measure DHEA, we used a double antibody 125I-RIA (DSL 8900, Diagnostic Systems Laboratories, Webster, TX). The DHEA RIA was modified to increase sensitivity (Granger et al., 1999) and has been previously used for songbird plasma (Goodson et al., 2005). Each sample was measured in duplicate. The DHEA antibody has a low cross-reactivity with DHEA-  36  S (0.02%), 16β-OH DHEA (0.041%), androstenedione (0.46%), testosterone (0.028%), and CORT (<0.01%). The intra-assay variation was 1.6% (n = 6 replicates in 1 assay), and the interassay variation was 2.0% for the low standard (25 pg/tube) and 7.0% for the high standard (100 pg/tube) (n = 4 assays). The lowest point on the standard curve was 2 pg DHEA/tube, and the detection limit for our assays with 15 µL plasma/tube was 0.13 ng/mL. Previous data indicate that DHEA must be extracted from song sparrow plasma prior to RIA (Newman et al., 2008), and here we used DCM to extract DHEA (3 mL x2), as in (Soma and Wingfield, 2001). To confirm that DCM extraction was effective, we examined recovery of 50 pg of exogenous DHEA that was added to a pool of song sparrow plasma prior to extraction. DHEA was extracted from 33 μl of plasma with DCM, and extracts were dried under nitrogen at 40˚C. Dried extracts were resuspended in 220 μl assay buffer and stored at 4˚C overnight. Furthermore, plasma was extracted with DCM, and the extracts were serially diluted and measured via RIA. Parallelism between the standard curve and serially-diluted extracted plasma indicates that DCM extraction removes substances in the plasma that interfere with the assay (Chard, 1995). The serial dilution was also used to determine optimal plasma volume for the RIA.  Statistical Analysis To determine parallelism between an RIA standard curve and a serial dilution of plasma, we tested equality of slopes using an analysis of covariance (ANCOVA). A lack of significant interaction between the standard curve and serial dilution of plasma indicates that the lines are parallel. To calculate an index of body condition, we used the residuals from a linear regression of body mass on tarsus length (Schulte-Hostedde et al., 2005).  37  To test for the effects of Season and Vein on baseline steroid levels, we used a 2-factor ANOVA. To examine the effects of Season, Vein and Stress on hormone levels, we used a 3factor mixed-design ANOVA where Season and Vein were between-subject factors and Stress was a within-subject factor (Zar, 1999). Where applicable, significant interactions with Season were broken down using 2-factor ANOVA within each season to assess the effects of Vein and Stress. For post hoc tests, we used Tukey’s honestly significant difference (HSD) tests. We used JMP IN 5.1 (SAS, Cary, NC). We considered test results significant for p < 0.05. Data are presented as mean ± standard error of the mean.  RESULTS Validation of Radioimmunoassays CORT RIA The CORT concentration measured in plasma extracted with DCM (31.34 ± 0.94 ng/mL; n=6) was not different from the CORT concentration measured in unextracted plasma (31.56 ± 1.16 ng/mL; n=6) (t-test; t = 0.15, df =10, p = 0.88). Furthermore, a serial dilution of unextracted plasma showed parallelism with the CORT standard curve (ANCOVA, R2 = 0.99, no significant interaction, F1,12 = 1.11, p = 0.32) (Fig 3.1A). Taken together, these data indicate that CORT can be measured directly in song sparrow plasma, without an extraction step, as in (Washburn et al., 2002).  DHEA RIA Recovery of 50 pg DHEA added to song sparrow plasma prior to extraction was 104.12 ± 8.6% (n = 4 pairs). Recoveries of 25 pg and 100 pg DHEA standards were 89.6 ± 4.5% and 91.3 ± 8.3%, respectively (n = 4 assays each). Furthermore, a serial dilution of DCM-extracted plasma was parallel to the DHEA standard curve (ANCOVA, R2 = 0.99, no significant interaction, F1,10 38  = 2.44, p = 0.17) (Fig 3.1B). Lastly, all water blanks were non-detectable (<2 pg DHEA) (n = 2 per assay, 8 total). Taken together, these data indicate that extraction of DHEA from song sparrow plasma with DCM was effective.  Baseline Steroid Levels Baseline CORT At baseline, there was a significant effect of Season on plasma CORT (F3,105 = 55.20, p < 0.0001), however there was no main effect of Vein (F1,105 = 0.12, p = 0.73) and no interaction between Season and Vein (F3,105 = 1.31, p = 0.27). Baseline plasma CORT levels were highest during the breeding season and similarly low during the other 3 seasons (Tukey’s HSD, p < 0.05) (Fig 3.2A).  Baseline DHEA Similar to CORT, there was a significant main effect of Season on baseline plasma DHEA (F3,106 = 29.84, p < 0.0001), and no main effect of Vein (F1,106 = 0.08, p = 0.78) and no interaction between Season and Vein (F3,106 = 0.33, p = 0.80). Baseline plasma DHEA levels were highest during the breeding season and mid-nonbreeding season, compared to the molt and early nonbreeding season (Tukey’s HSD, p < 0.05) (Fig 3.2B).  Stressed Steroid Levels Within a vein and season, there were no significant differences in CORT or DHEA concentrations in plasma samples collected after 30 or 60 min of restraint (Table 3.1). Previous studies on sparrows have also shown that CORT levels do not differ between 30 and 60 min of restraint (Breuner and Orchinik, 2001; Romero and Romero, 2002). Therefore, 30 and 60 min data were pooled within a vein and within a season. Moreover, separate analyses using 1) 39  stressed steroid levels at 30 min or 2) pooled stressed steroid levels (30 and 60 min) were not different; thus pooling the data did not alter the results.  Stressed CORT To examine the effects of Season, Vein and Stress on plasma CORT, we used a 3-factor mixed design ANOVA. The main effects of Season, Vein and Stress were significant (Table 3.2). Also, the interaction between Season and Stress was significant, and thus this interaction was broken down using a 2-factor ANOVA within each season to examine the main effects of Vein and Stress on plasma CORT. Within all seasons, there was a significant main effect of Stress (Fig 3.3A, Table 3.3). Interestingly, only during the molt was there a significant main effect of Vein and a significant interaction between Vein and Stress (Fig 3.3A, Table 3.3). During molt, jugular CORT levels were significantly higher than brachial CORT levels after stress (Tukey’s HSD, p < 0.05) (Fig 3.3A).  Stressed DHEA To examine the effects of Season, Vein and Stress on plasma DHEA, we used a 3-factor mixed design ANOVA. There was a significant main effect of Season; however the main effects of Vein and Stress were not significant (Table 3.2). The interaction between Season and Vein was significant, as was the 3-way interaction of Season x Vein x Stress (Table 3.2). Because the 3way interaction was significant, we used a 2-factor ANOVA within each season to examine the effects of Vein and Stress on plasma DHEA. During the breeding season, the main effect of Vein was significant, the main effect of Stress approached significance and the interaction between Vein and Stress was significant (Table 3.3). Stressed jugular DHEA was significantly lower than baseline jugular DHEA 40  (Tukey’s HSD, p < 0.05) but stressed brachial DHEA was not different from baseline brachial DHEA (Tukey’s HSD, p > 0.05). These results indicate that stress decreased jugular DHEA but did not affect brachial DHEA during the breeding season (Fig 3.3B, Table 3.3). During the molt, the main effects of Vein and Stress were not significant, but the interaction between Vein and Stress was significant (Table 3.3). Stressed jugular DHEA was significantly higher than baseline jugular DHEA (Tukey’s HSD, p < 0.05). Stressed jugular DHEA was also significantly higher than baseline and stressed brachial DHEA (Tukey’s HSD, p < 0.05). Stressed brachial DHEA was not different from baseline brachial DHEA (Tukey’s HSD, p > 0.05). These results indicate that stress increased jugular DHEA but did not affect brachial DHEA during the molt (Fig 3.3B, Table 3.3). During the early and mid-nonbreeding seasons, neither the main effects of Vein and Stress, nor the interaction between Vein and Stress were significant (Fig 3.3B, Table 3.3).  Relationship between CORT and DHEA Baseline levels of CORT and DHEA were positively correlated across seasons (r(100) = 0.47, p < 0.0001). However, within each season, there was not a significant correlation between baseline levels of CORT and DHEA. Stressed levels of CORT and DHEA were also positively correlated across seasons (r(101) = 0.36, p = 0.0003). However, within each season, there was not a significant correlation between stressed levels of CORT and DHEA. The CORT/DHEA ratios at baseline and after acute restraint were significantly higher during the breeding season in brachial and jugular plasma (Table 3.4).  41  Body Condition Body condition was calculated using the residuals from a regression of body mass on tarsus length. Body condition changed significantly with season, and subjects in the mid-nonbreeding season had the highest body condition (Table 3.5). Baseline CORT levels were not correlated with body condition in any season; however, stressed CORT levels were negatively correlated with body condition in the breeding season (r(32) = -0.39, p = 0.05). In contrast, baseline DHEA levels were positively correlated with body condition in the breeding season (r(32) = 0.45, p = 0.02). Fat score also changed significantly with season, with fat stores being much greater during the mid-nonbreeding season (Table 3.5). The androgen-dependent cloacal protuberance length changed significantly with season (Table 3.5). The cloacal protuberance was largest during the breeding season, and decreased progressively in the molt, early nonbreeding season, and mid-nonbreeding season (Table 3.5). A small cloacal protuberance indicates low circulating testosterone levels (Soma et al., 2004; Soma et al., 2003).  DISCUSSION Plasma CORT and DHEA changed dramatically over the seasons. Both steroids were regulated by acute restraint stress; however the effect of stress was dependent on season and vein. Stress decreased jugular DHEA during breeding, whereas stress increased jugular DHEA during molt. Stress did not affect brachial DHEA in any season. Across seasons, stress increased CORT in both veins. However, during molt, stress increased jugular CORT significantly more than brachial CORT. Previous songbird studies have shown differences between jugular and systemic plasma levels of neurally-synthesized radiolabeled estrogens (Saldanha and Schlinger, 1997; Schlinger and Arnold, 1992; Schlinger and Arnold, 1993), but this study is the first to 42  suggest differences between jugular and systemic levels of endogenous steroids in adults. Moreover, this is the first study to examine acute regulation of DHEA in multiple seasons.  Baseline Steroid Levels At baseline, brachial CORT did not differ from jugular CORT in any season. Baseline CORT levels were significantly higher during the breeding season than during the rest of the year, and this robust seasonal pattern is consistent with previous studies in songbirds (Romero, 2002). Seasonal modulation of plasma CORT levels can occur due to changes at the level of the hypothalamus, the pituitary or the adrenals (Romero et al., 1998). Elevated baseline CORT in breeding males may facilitate energy mobilization to meet the demands of feeding and raising young, which is energetically demanding in this biparental species (Hambly et al., 2007; Romero, 2002). Baseline levels of brachial DHEA were significantly higher during the breeding and mid-nonbreeding seasons, consistent with previous data in song sparrows (Soma and Wingfield, 2001). During the breeding season, the testes are recrudesced and secrete high levels of androgens. During the non-breeding season, the testes are fully regressed and circulating testosterone is non-detectable (Wingfield and Hahn, 1994). Importantly, at baseline, jugular DHEA did not differ from brachial DHEA at any time of year. Thus, these data do not support the hypothesis that the brain is the main source of circulating DHEA in the non-breeding season. Nonetheless, it remains possible that DHEA is synthesized in the brain (Tsutsui et al., 2000) and acts locally, without being secreted into the jugular. Future studies will measure DHEA directly in specific brain regions. Baseline CORT and DHEA in both veins were positively correlated across all seasons. The striking similarity in the pattern of seasonal change suggests that both CORT and DHEA may be of adrenal origin.  43  Chronic elevations in CORT have detrimental effects on brain and immune function (Vanitallie, 2002). DHEA is a potent antiglucocorticoid, and thus an elevation in DHEA during the breeding season could mitigate the detrimental effects of CORT. For example, DHEA is a potent antiglucocorticoid in the brain and diminishes the neurodegenerative effects of CORT in the rodent brain (Kalimi et al., 1994; Karishma and Herbert, 2002). In mice, DHEA also ameliorates the immunosuppressive effects of CORT (Blauer et al., 1991). During the breeding season (when CORT is high), elevated DHEA may play a neuroprotective or immune-enhancing role. Interestingly, baseline DHEA was positively correlated with body condition during the breeding season, further suggesting a beneficial effect of high DHEA. The CORT/DHEA ratio was highest during the breeding season. An elevated CORT/DHEA ratio supports the idea that the breeding season is a particularly demanding period. The CORT/DHEA ratio is also increased in some studies of patients with major depression and schizophrenia (Ritsner et al., 2004; Young et al., 2002). Further studies are needed to clarify the significance of the CORT/DHEA ratio in songbirds.  Effects of Acute Restraint on Plasma CORT Restraint stress increased brachial and jugular CORT levels in every season, but the magnitude of the increase showed large seasonal variation, consistent with previous studies (Romero, 2002). This is the first study to examine seasonal changes in jugular CORT levels. In both veins, the CORT response to stress was significantly higher during the breeding season and similar in the other seasons. Increased CORT responsiveness during the breeding season may prepare an individual for a subsequent stressor (Sapolsky et al., 2000). Thus, stressed CORT should be maximal when the probability of encountering a stressor (e.g. predator, parasite, competitor for mates) is greatest, and for many species this period is the breeding season (Romero, 2002). 44  Specifically during the molt, the increase in jugular CORT was significantly greater than the increase in brachial CORT, even though baseline levels were similar in the two veins. The molt is the annual replacement of feathers after the breeding season (Romero et al., 2005). Previous studies have shown that baseline CORT and stressed CORT levels are dramatically reduced during molt (Romero, 2006). Further, CORT treatment slows feather growth in molting birds (Romero et al., 2005). During molt, the decrease in systemic CORT levels may facilitate protein deposition during feather growth and avoid the catabolic actions of CORT (Romero et al., 2005). The present data raise the intriguing hypothesis that during molt, when adrenal synthesis of CORT is downregulated, neural synthesis of CORT is upregulated. There is some evidence that the brain has the capacity to synthesize CORT in mammals (Davies and MacKenzie, 2003; Gomez-Sanchez et al., 2005; Mellon and Deschepper, 1993; Stoeffel-Wagner, 2003), and future studies will measure CORT directly in brain tissue. Increased local synthesis of CORT in the brain during molt might allow for behavioral responses to stressors (e.g. predators), while avoiding the costs of high circulating CORT on feather growth. Similar mechanisms may operate during the nonbreeding season to avoid the costs of high circulating testosterone (Wingfield et al., 2001). An alternative hypothesis is that circulating CORT is sequestered in the brain during molt and released in response to stress, but at present there is little evidence for this hypothesis.  Effects of Acute Restraint on Plasma DHEA Acute restraint stress for 30 or 60 min did not affect brachial DHEA levels in any season. These data are consistent with a previous study that used a 30 min restraint (Soma and Wingfield, 2001). Thus, even though CORT and DHEA levels are positively correlated across the seasons, suggesting similar long-term regulation, short-term regulation by stress is quite different for 45  these two steroids. In humans, both cortisol and DHEA are synthesized in the adrenal cortex and are acutely regulated by stress and ACTH (Engelmann et al., 2004; Kroboth et al., 1999; Oberbeck et al., 1998; Salek et al., 2002). In song sparrows, during the nonbreeding season, DHEA concentrations in the adrenals are nearly 10 times higher than in plasma (Newman and Soma, 2007; Soma and Wingfield, 2001). Nonetheless, acute stress has no effect on systemic DHEA levels. Similarly, the bovine adrenal synthesizes DHEA (Ogo et al., 1991), but shortterm ACTH treatment has no effect on systemic DHEA levels (Marinelli et al., 2007). In contrast, long-term ACTH treatment increases circulating DHEA levels in cows (Marinelli et al., 2007). If ACTH has chronic, but not acute, effects on adrenal DHEA synthesis in song sparrows, that might explain why seasonal changes in systemic CORT and DHEA are positively correlated but the effects of acute stress differ. Unlike brachial DHEA, jugular DHEA was significantly affected by acute stress in a season-specific manner. During the breeding season, stress significantly decreased jugular DHEA levels. A decrease in jugular DHEA suggests a decrease in neural DHEA synthesis or an increase in neural DHEA metabolism. In captive breeding male song sparrows, acute restraint increases the activity of brain 3β-HSD, the enzyme that metabolizes DHEA to androstenedione (Soma et al., 2002). These data are consistent with stress increasing neural DHEA metabolism in the breeding season. During the breeding season, there was greater individual variation in both brachial and jugular DHEA. This individual variability could be the result of differences in breeding substage. Within the breeding season, hormone levels fluctuate according to sub-stage (e.g. nest building, egg laying, incubation, feeding chicks, re-nesting) and also decline from the first brood to the second brood (Wingfield and Hahn, 1994). This variability may have affected our ability to assess the effect of Vein between individuals. However, the effect of Stress was assessed within individuals, which controlled for individual differences. 46  In contrast to the breeding season, stress increases jugular DHEA levels during the molt. An increase in jugular DHEA suggests either an increase in neural DHEA synthesis or a decrease in neural DHEA metabolism. Molt is the only season when jugular DHEA increases in response to stress, and this coincides with the data on jugular CORT during molt. During molt, when systemic levels of CORT and DHEA are low, it is possible that the downregulation of systemic steroid signals is accompanied by an upregulation of local steroid production. The balance between systemic and local steroid signaling mechanisms remains enigmatic (Soma et al., 2005) but has important implications for patients with adrenal insufficiency and Addison’s disease (Arlt and Allolio, 2003) and also for the stress hyporesposive period during development (Sapolsky and Meaney, 1986).  Conclusions The present results indicate pronounced season-dependent effects of acute restraint stress on CORT and DHEA levels. Further, the effects depend on whether the steroids are measured in brachial or jugular plasma. The positive correlation between baseline CORT and DHEA levels across seasons suggests that CORT and DHEA are regulated similarly in the long-term. However, systemic CORT but not systemic DHEA is regulated by stress in the short-term. Importantly, the effects of acute stress on jugular DHEA suggest a role for DHEA in the brain. Future studies shall focus on CORT and DHEA levels in specific brain regions and the effects of natural chronic stress (Clinchy et al., 2004). Lastly, these data highlight the importance of the blood sampling site. In endocrine studies, blood is collected from a variety of sites, including the brachial vein, jugular vein, tail vein, saphenous vein and retro-orbital sinus. It is possible that blood samples from these different sites have different steroid profiles.  47  TABLES Table 3.1 No significant differences between 30 min restraint and 60 min restraint.  Jugular  Brachial  Breeding Season  Early nonbreeding Season  30 min  60 min  t  p  30 min  60 min  t  p  Plasma CORT (ng/mL)  199.71 ± 21.96 (n = 8)  146.6 ± 23.05 (n = 8)  1.74  0.11  39.50 ± 6.80 (n = 9)  52.14 ± 6.64 (n = 9)  -1.30  0.21  Plasma DHEA (pg/mL) Plasma CORT (ng/mL) Plasma DHEA (pg/mL)  961.82 ± 177.17  1138.83 ± 204.58  -0.65  0.53  420.57 ± 48.72  385.89 ± 40.69  0.55  0.59  161.96 ± 22.71 (n = 8)  160.62 ± 22.71 (n = 10)  0.04  0.97  38.52 ± 3.83 (n = 10)  35.87 ± 3.83 (n = 10)  0.49  0.63  762.26 ± 212.48  619.9 ± 190.05  0.50  0.62  394.31 ± 41.15  395.91 ± 36.69  -0.03  0.98  Note: Sample sizes for brachial and jugular plasma samples are indicated in parentheses. The 60 min restraint was conducted only during the breeding season and early nonbreeding season.  48  Table 3.2 Effects of Season, Vein and Stress on plasma CORT and DHEA are dependent on season and vein. 3-way mixed design ANOVA Variable  CORT  DHEA  df  F-ratio  p  df  F-ratio  p  Season  3, 105  131.72  < 0.0001  3, 106  41.32  < 0.0001  Vein  1, 105  4.58  0.03  1, 106  0.07  0.67  Season x Vein  3, 105  0.78  0.51  3, 106  4.93  0.003  Stress  1, 105  420.94  < 0.0001  1, 106  1.22  0.27  Season x Stress  3, 105  5.06  0.0002  3, 106  1.58  0.20  Vein x Stress  1, 105  0.01  0.91  1, 106  1.59  0.21  Season x Vein x Stress  3, 105  1.55  0.20  3, 106  14.01  < 0.0001  49  Table 3.3 Effects of Vein and Stress on plasma CORT and DHEA within each season. 2-way mixed design ANOVA Season  Breeding  Molt  Early nonbreeding  Midnonbreeding  Variable  CORT  DHEA  df  F-ratio  p  df  F-ratio  p  Vein  1, 33  0.58  0.45  1, 34  4.46  0.04  Stress  1, 33  100.42  < 0.0001  1, 34  3.59  0.07  Vein x Stress  1, 33  0.39  0.54  1, 34  22.10  < 0.0001  Vein  1, 18  7.95  0.01  1, 18  0.70  0.41  Stress  1, 18  207.81  < 0.0001  1, 18  0.33  0.57  Vein x Stress  1, 18  4.81  0.04  1, 18  21.38  0.0002  Vein  1, 37  0.14  0.71  1, 37  0.99  0.33  Stress  1, 37  391.04  < 0.0001  1, 37  2.65  0.11  Vein x Stress  1, 37  1.09  0.40  1, 37  0.03  0.87  Vein  1, 15  1.21  0.29  1, 15  0.28  0.60  Stress  1, 15  83.72  < 0.0001  1, 15  0.03  0.87  Vein x Stress  1, 15  0.11  0.74  1, 15  0.02  0.88  50  Jugular  Brachial  Table 3.4 CORT/DHEA ratio at baseline and after acute restraint stress. Breeding (n = 16, 18)  Molt (n = 10, 8)  Early nonbreeding (n = 18, 18)  Mid-nonbreeding (n = 8, 8)  F-ratio  p  Baseline CORT/DHEA  1.58 ± 0.04 a  1.45 ± 0.04 b  1.40 ± 0.02 b  1.46 ± 0.03 b  13.52  < 0.0001  Stress CORT/DHEA  1.87 ± 0.03 a  1.73 ± 0.02 b  1.78 ± 0.02 b  1.73 ± 0.02 b  10.87  < 0.0002  Baseline CORT/DHEA  1.62 ± 0.02 a  1.45 ± 0.3 b  1.45 ± 0.02 b  1.49 ± 0.04 b  14.43  < 0.0001  Stress CORT/DHEA  1.95 ± 0.03 a  1.77 ± 0.04 b  1.76 ± 0.01 b  1.75 ± 0.03 b  12.63  < 0.0001  Note: Ratio calculated using log transformed corticosterone and DHEA concentrations. Numbers in parentheses indicate sample sizes for brachial and jugular plasma, respectively. Within a row, values that share the same letter are not significantly different.  51  Table 3.5 Seasonal changes in body condition, fat and cloacal protuberance length. Breeding (n = 34)  Molt (n = 19)  Early nonbreeding (n = 38)  Mid-nonbreeding (n = 16)  F-ratio  p  Body Condition  -0.33 ± 0.23 ab  0.44 ± 0.23 a  -0.73 ± 0.18 b  2.26 ± 0.36 c  24.70  < 0.0001  Fat Score  0.42 ± 0.06 a  0.69 ± 0.07 b  0.55 ± 0.06 ab  1.97 ± 0.46 c  38.48  < 0.0001  Cloacal Protuberance (mm)  9.50 ± 0.15 a  5.23 ± 0.18 b  4.46 ± 0.08 c  3.32 ± 0.16 d  353.55  < 0.0001  Note: Within a row, values that share the same letter are not significantly different.  52  FIGURES Figure 3.1 Serially diluted plasma in corticosterone and DHEA radioimmunassays A 100  CORT standard curve Unextracted plasma  Percent Bound  80 60 40 20 0  0.1  1  10  100  1000  CORT (pg) or plasma (µL)  B 100 DHEA standard curve DCM extracted plasma  Percent Bound  80 60 40 20 0  1  10  100  1000  DHEA (pg) or plasma (µL)  Figure 3.1 Validation of the (A) corticosterone and (B) DHEA radioimmunoassays. (A) Serially diluted, unextracted plasma was parallel to the CORT standard curve. (B) Serially diluted, DCM extracted plasma was parallel to the DHEA standard curve.  53  Figure 3.2 Effects of season on baseline corticosterone and DHEA in brachial and jugular plasma.  Baseline Plasma CORT (ng/mL)  A 60  A  Brachial Jugular  50 40 a  30  B  20 B  10  B b  b  b  0  a  1200 1050 900  a  750 A A  600 b  b  450 B  300 0  ed M id in -n g on br ee di ng  Ea r  ly  no nb re  B re e  M ol t  B  di ng  Baseline Plasma DHEA (pg/mL)  B  Figure 3.2 Baseline levels of plasma CORT and DHEA. Upper case letters refer to brachial plasma and lower case letters refer to jugular plasma. Data points that share the same letter are not significantly different. (A) Baseline CORT was highest during the breeding season and lower during the molt, early and mid-nonbreeding seasons. Baseline CORT did not differ between veins in any season. (B) Baseline DHEA was highest during the breeding and midnonbreeding seasons, and lower during the molt and early nonbreeding season. Baseline DHEA did not differ between veins in any season. Data are means ± SEM.  54  Figure 3.3 Effects of season and acute restraint stress on corticosterone and DHEA in jugular and brachial plasma.  Plasma CORT (ng/mL)  B  250  Plasma DHEA (pg/mL)  A  1200  Breeding  Molt  Early nonbreeding  Mid-nonbreeding  (16)  200  Brachial Jugular  (18)  150  * (9)  100 50  (8) (8)  (19) (18)  (10)  0  1050 (16)  900 750  (8) (8)  (18)  * (9)  600 450  *  300 0 Baseline  Stressed  (18) (20)  (10) Baseline  Stressed  Baseline  Stressed  Baseline  Stressed  Figure 3.3 Effects of acute restraint stress on plasma CORT and DHEA levels. (A) In all four seasons, stress increased plasma CORT in the brachial and jugular veins. Importantly, during the molt, stress increased CORT significantly more in the jugular vein. (B) Stress decreased jugular DHEA during the breeding season and increased jugular DHEA during the molt. Stress did not affect brachial DHEA in any season. 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Glucocorticoids are synthesized de novo from cholesterol by the adrenal glands and also by the brain itself (Davies and MacKenzie, 2003; Gomez-Sanchez et al., 2005; Ye et al., 2008). Local steroid synthesis could increase neural glucocorticoid concentrations independently of systemic glucocorticoid concentrations. Local glucocorticoid levels are regulated, in part, by 11β-hydroxysteroid dehydrogenase (11β-HSD) isozymes, which catalyze the conversion between active glucocorticoids and their inactive metabolites (Pelletier et al., 2007). Although numerous studies have measured systemic glucocorticoid levels in plasma or serum, very few studies have measured local glucocorticoid levels in the brain (Thoeringer et al., 2007; Droste et al., 2008) Dehydroepiandrosterone (DHEA), a sex-steroid precursor, can also be synthesized by the adrenals and brain (Baulieu, 1998; Labrie et al., 2005). DHEA has antiglucocorticoid properties (Kalimi et al., 1994) and reduces the neurodegenerative effects of elevated glucocorticoids on the rat hippocampus in vitro (Kimonides et al., 1999) and in vivo (Karishma and Herbert, 2002). Moreover, DHEA levels in the rat brain are increased by acute stress (Corpechot et al., 1981) and ACTH treatment (Torres and Ortega, 2003). DHEA may regulate local corticosteroid levels via effects on 11β-HSD isozymes (Apostolova et al., 2005; Balazs et al. 2008). Together, these data suggest that neural DHEA is regulated by stress and modulates the actions of glucocorticoids.  3  A version of this chapter has been accepted for publication. Newman, A.E.M., and Soma, K.K. Corticosterone and DHEA in songbird plasma and brain: effects of season and acute stress. European Journal of Neuroscience, in press.  61  In a wild songbird, the song sparrow (Melospiza melodia), acute restraint affects corticosterone and DHEA levels in plasma from the brachial and jugular veins (Newman et al., 2008a). Brachial plasma is an index of systemic steroid levels, and jugular plasma is enriched with neurally-synthesized steroids and is an indirect index of neural steroid levels (Schlinger and Arnold, 1992, 1993). During the breeding season, stress does not affect brachial DHEA levels but decreases jugular DHEA levels. During the molt, stress does not affect brachial DHEA levels but increases jugular DHEA levels. Also during the molt, stress increases corticosterone levels to a greater extent in jugular plasma than brachial plasma. A specific change in jugular steroid levels suggests a change in neurosteroid synthesis or metabolism. However, jugular plasma is an indirect index of neural steroid levels. Here, we directly examined the effects of acute restraint stress on corticosterone and DHEA concentrations in jugular plasma and brain of wild song sparrows under natural conditions. Hypothalamic-pituitary-adrenal axis activity varies across seasons (Romero et al., 2002; Pyter et al., 2007), and thus we examined animals from three distinct seasons of the year: breeding season, molt, and nonbreeding season. This design allowed us to examine the effects of acute stress and season, as well as compare plasma and brain steroid concentrations within individuals. We were particularly interested in the hippocampus, which is a major glucocorticoid target in the brain.  MATERIALS and METHODS Subjects Subjects were wild adult male song sparrows (n = 46 total). Song sparrows are an excellent model for studying seasonal plasticity in hormones and the adult brain (Tramontin and Brenowitz, 2000; Soma et al., 2008). This species shows large seasonal changes in circulating corticosterone and DHEA (Soma and Wingfield, 2001; Newman et al 2008a). Song sparrows, 62  unlike laboratory rats or mice, have relatively high levels of plasma DHEA. Furthermore, a physiological dose of DHEA has pronounced effects on neuroanatomy and behavior (Soma et al., 2002). DHEA is metabolized to androstenedione in the songbird brain, and this metabolism is affected by acute stress (Soma et al., 2004; Pradhan et al., 2008). The songbird brain might also synthesize DHEA de novo (London et al., 2006). Song sparrows were captured during 3 seasons: 1) Breeding (May 9-17, 2006; baseline n = 8; stressed n = 9); 2) Molt (August 12-18, 2006; baseline n = 5; stressed n = 6); and 3) Nonbreeding (Jan 3-11, 2006; baseline n = 9; stressed n = 9). Subjects were captured near Vancouver, British Columbia (49° 12’N, 123° 01’W). This population is sedentary, and males maintain territories year-round (Wingfield and Hahn, 1994; Soma, 2006). During the breeding season, plasma testosterone levels are elevated, and males aggressively defend territories. During the molt when song sparrows are replacing their feathers, plasma testosterone levels are nondetectable, and territorial aggression is reduced. During the nonbreeding season when testes are regressed, plasma testosterone is non-detectable, but territorial aggression is high. Protocols were approved by the UBC Committee on Animal Care and complied with the guidelines of the Canadian Council of Animal Care.  Blood Collection Subjects were captured using conspecific playback (3.50 ± 0.53 min) and mist nets. A baseline blood sample (~150 µL) was collected within 3 min of capture (2.05 ± 0.07 min) from the jugular vein with a heparinized syringe (Newman et al., 2008a). Subjects were sacrificed by rapid decapitation within 3 min of capture (2.60 ± 0.16 min) or restrained for 30 min in a cloth bag. After restraint, another blood sample was collected from the jugular vein, and subjects were sacrificed (32.71 ± 0.23 min after capture). Blood was kept on wet ice until centrifuged. Plasma was stored at -20°C. 63  Tissue collection After cooling on wet ice for 2-3 min, the brain was dissected into 6 regions: 1) rostral diencephalon (rDien); 2) caudal diencephalon (cDien); 3) hippocampus (Hp); 4) caudal medial nidopallium (NCM); 5) dorsal telencephalon (dTel) containing the song nucleus HVC; and 6) central medial telencephalon (cmTel) containing the septum, bed nucleus of the stria terminalis, and nucleus accumbens (Goodson et al., 2004; Montagnese et al., 2004). The dissection protocol closely followed Soma et al. (1999). Briefly, the hippocampi were isolated by making two parallel cuts approximately 1.5mm lateral to the midline and 1 mm deep. The songbird Hp is located on the dorsomedial surface of the telencephalon and lies directly above the lateral ventricle (Saldanha et al., 1998). Next, we removed a triangular piece of the dTel from the area lateral to the hippocampus, designed to match the location of HVC (Reiner et al., 2004). Next, NCM was removed from directly under the excised hippocampus by removing a rectangular piece of tissue 1.5mm lateral to the midline and approximately 1.5mm deep. The brain was then removed from the cranium and placed on its dorsal surface on a petri dish resting on wet ice. Then the optic lobes and hindbrain (to the level of the mammillary bodies) were removed. The diencephalon was removed to the depth of the anterior commissure and split into rDien and cDien. The remaining telencephalon was divided into three regions (rostral, central and caudal). The central telencephalon was further split into medial and lateral sections. All brain regions were divided into left and right halves; one half was used to measure steroid levels (present study) and the other half will be used to measure steroidogenic enzymes (to be presented separately). Left or right halves of brain tissues were chosen at random for steroid measurement. The body was kept on wet ice until dissected. We collected a piece of pectoral muscle, a piece of liver, the testes, and the adrenals. The entire dissection took less than 30 min. Tissues were stored separately in microcentrifuge tubes and frozen immediately on dry ice. In the laboratory, tissue was stored at -80°C. 64  Steroid Measurement Steroids were extracted from tissue and plasma samples (n = 571 total) using solid phase extraction with C18 columns, as previously described (Newman et al., 2008b). This extraction procedure results in high and consistent steroid recoveries and effectively removes interfering substances from plasma and lipid-rich brain tissue (Newman et al., 2008b). We have thoroughly validated this procedure with both plasma and brain tissue for corticosterone and DHEA assays (e.g., serial dilutions, recovery of radiolabeled and radioinert steroids) (Newman et al., 2008a,b; Schmidt and Soma, 2008). In addition, here we examined recovery of 30pg of exogenous corticosterone and 35pg of exogenous DHEA that were added to a song sparrow plasma pool (n = 6 replicates) and a brain tissue pool (n = 6 replicates) prior to extraction. Corticosterone and DHEA concentrations in these plasma and tissue samples were compared to concentrations in plasma and tissue samples from the pools that did not have exogenous steroid added (n = 6 each). Recovery of exogenous corticosterone was 94.3% from plasma and 73.9% from brain. Recovery of exogenous DHEA was 73.8% from plasma and 70.8% from brain. We used sensitive and specific radioimmunoassays (RIAs) to measure corticosterone (ImmuChem 07-120103, MP Biomedicials, Orangeburg, NY) and DHEA (DSL 8900, Diagnostic Systems Laboratories, Webster, TX) (Newman et al., 2008a,b; Schmidt and Soma, 2008). The corticosterone antibody has very low cross-reactivities to cortisol (0.05%), 11deoxycorticosterone (0.34%), dehydrocorticosterone (0.50%), progesterone (0.02%) and other steroids (Schmidt and Soma, 2008). The DHEA antibody has very low cross-reactivities to androstenedione (0.73%), testosterone (0.28%), progesterone (0.05%), DHEA-sulfate (0.02%) and other steroids (Boonstra et al., 2008; Newman et al., 2008b). Dried eluates were resuspended in 250 µL phosphate-buffered saline with gelatin. We used absolute ethanol (5% of resuspension volume) to aid in resuspension of steroids (Newman et al., 2008a). 100 µL (×2) was assayed for DHEA. Of the remaining 50 µL resuspension, 30 µL was removed, brought up 65  to 100 µL with phosphate-buffered diluent provided with the corticosterone RIA, and 50 µL (×2) was assayed for corticosterone. The lowest points on the corticosterone and DHEA standard curves were 3.12 pg corticosterone and 2 pg DHEA (per tube). For corticosterone, intra-assay variation was 4.1%, and inter-assay variation was 8.0% (low control) and 6.8% (high control) (n = 10 assays). For DHEA, intra-assay variation was 5.9%, and inter-assay variation was 10.1% (low control) and 7.9% (high control) (n=12 assays). In replicates of the “unspiked” plasma and brain tissue pools (see above), variability was low. For corticosterone, the coefficient of variation was 4.08% for plasma and 4.16% for brain (n=6 replicates each). For DHEA, the coefficient of variation was 8.93% for plasma and 6.95% for brain (n=6 replicates each). For corticosterone, all water blanks were nondetectable (< 3.12 pg; n = 12). For DHEA, 11 of 12 water blanks were nondetectable (< 2 pg) and one was slightly above the lowest standard. Thus, the steroid measurement protocol functions reliably for both plasma and brain tissue.  Statistics Nondetectable samples (below the lowest point on the standard curve) were set to zero, as previously done for small tissues (Schmidt and Soma, 2008). Data were corrected for recovery and transformed (log (x +1)) to reduce heteroscedasticity prior to analysis with JMP IN 5.1 (SAS, Cary, NC). For plasma, the effects of season and stress were tested using a two-factor ANOVA and Tukey’s honestly significant difference (HSD) post hoc tests. For brain tissue, we used mixeddesign three-factor analysis of variance (ANOVA) to test the effects of season, stress, and brain region on corticosterone and DHEA levels. Season and stress were between-subject factors and brain region was a within-subject factor. Significant interactions were broken down using twofactor ANOVA tests within each season to assess the effects of stress and brain region. If the 66  two-factor ANOVA revealed significant differences, we used Tukey’s honestly HSD tests for post hoc comparisons. For peripheral tissues, the effects of season and stress on steroid levels were examined using two-factor ANOVA tests, and significant differences were analyzed using Tukey’s HSD post hoc tests. Steroid levels in plasma and two specific brain regions (cmTel and Hp) were directly compared using mixed-design two-factor ANOVA tests, in which season was a between-subject factor and sample type (plasma or brain tissue) was a within-subject factor. The septum (in the cmTel) and Hp readily bind tritiated corticosterone (McEwen et al., 1968) and are sensitive to restraint stress (Goodson et al., 2004). Furthermore, both regions contain steroidogenic enzymes (Hojo et al., 2004; Soma et al., 2004; Tsutsui et al., 2006). Lastly, in the cmTel, corticosterone and DHEA were detectable in most samples, and steroid concentrations were similar to those in other brain regions (except the Hp). The ratio of corticosterone to DHEA was calculated in jugular plasma, cmTel and Hp. This ratio has been informative in studies of patients with stress-related psychiatric diseases (Ritsner et al., 2004). The corticosterone/DHEA ratio was calculated by dividing the corticosterone concentration (in pg/mL) by the DHEA concentration (in pg/mL). We presented a similar analysis for brachial and jugular plasma (Newman et al., 2008b), although in that study we presented log-transformed data. We considered results significant for p ≤ 0.05. Data are presented as mean ± SEM.  RESULTS Jugular corticosterone To examine the effects of season and stress on jugular corticosterone, we used a two-factor ANOVA (Fig 4.1). There were significant main effects of season (F2,67 = 148.89, p < 0.0001) and stress (F1,67 = 299.02, p < 0.0001), and the interaction between season and stress was 67  significant (F2,67 = 109.90, p < 0.0001). Post hoc tests revealed that baseline corticosterone was highest during the breeding season and similarly low during the molt and nonbreeding season (Tukey’s HSD, p < 0.05). Stress increased jugular corticosterone in all seasons, but the effect of stress was greatest during the breeding season and similar during the molt and nonbreeding seasons (Tukey’s HSD, p < 0.05).  Brain corticosterone In a mixed-design three-factor ANOVA examining the effects of season, stress, and brain region on corticosterone, the main effects of stress and region were significant but there was no significant main effect of season (Fig 4.1, Table 4.1). Also, the season × region, stress × region, and season × stress × region interactions were significant (Table 4.1). To break down the three-way interaction, we used a mixed-design two-factor ANOVA within each season. During the breeding season, there was a significant main effect of stress on corticosterone (F1,100 = 115.87, p < 0.0001). There was no effect of region (F5,100 = 0.92, p = 0.48) and no interaction between stress and region (F5,100 = 0.45, p = 0.81). During the molt, there was a significant interaction between stress and region (F5,64 = 7.96, p < 0.0001). Post hoc tests revealed that stressed corticosterone was higher than baseline corticosterone levels in all brain regions except in the Hp, where stressed corticosterone were significantly lower than baseline corticosterone levels (Tukey’s HSD, p < 0.05 in all cases). These data suggest that stress reduces corticosterone in the Hp during molt. During the nonbreeding season, there was a significant effect of stress (F1,105 = 21.13, p < 0.0001), but no effect of region (F5,105 = 0.88, p = 0.50) or interaction between stress and region (F5,105 = 0.65, p = 0.66).  68  Jugular DHEA We used a two-factor ANOVA to examine the effects of stress and season on jugular DHEA (Fig 4.2). There was a significant main effect of season (F2,67 = 4.61, p < 0.002), no main effect of stress (F1,67 = 2.08, p = 0.15), but the interaction between season and stress was significant (F2,67 = 3.83, p = 0.03). Post hoc tests revealed that baseline DHEA levels were lower during the molt than during the breeding and nonbreeding seasons and that stress significantly increased jugular DHEA during the molt (Tukey’s HSD, p < 0.05).  Brain DHEA In a mixed-design three-factor ANOVA examining the effects of season, stress, and brain region on DHEA, there was a significant main effect of region but no effects of stress or season (Fig 4.2, Table 4.1). The Hp had higher levels of DHEA than other regions (Tukey’s HSD, p < 0.05).  Corticosterone and DHEA concentrations in jugular plasma versus brain We used mixed-design two-factor ANOVA tests to examine the effects of season and sample type (plasma vs. cmTel vs. Hp) on corticosterone levels at baseline and after stress. At baseline, there was a significant main effect of season and an interaction between season and sample type (Fig 4.3A, Table 4.2). During the breeding and nonbreeding seasons, baseline corticosterone concentrations were greater in plasma than in cmTel or Hp (Tukey’s HSD, p < 0.05). During the molt, there was no difference in baseline corticosterone concentrations in plasma, cmTel and Hp (Tukey’s HSD, p > 0.05). After stress, the main effects of season and sample type were significant, as was the interaction between season and sample type (Fig 4.3B, Table 4.2). During the breeding and nonbreeding seasons, stressed corticosterone concentrations were greater in plasma than in cmTel or Hp (Tukey’s HSD, p < 0.05). During the molt, stressed 69  corticosterone concentrations were greater in plasma than in Hp (Tukey’s HSD, p < 0.05), but levels in cmTel were not different from either plasma or Hp (Tukey’s HSD, p > 0.05). We used the same approach to examine the effects of season and sample type on DHEA levels in plasma, cmTel and Hp. At baseline and after stress, there was a significant main effect of sample type only (Fig 4.3CD, Table 4.2). At baseline, DHEA concentrations were significantly lower in plasma than in cmTel, and DHEA concentrations were highest in the Hp (Tukey’s HSD, p < 0.05). After stress, DHEA concentrations were significantly lower in plasma than in Hp, and DHEA concentrations in cmTel were intermediate and not different from either plasma or Hp (Tukey’s HSD, p < 0.05).  Ratio of corticosterone to DHEA in jugular plasma versus brain At baseline and after stress, we used a mixed-design two-factor ANOVA to examine the effects of season and sample type on the corticosterone/DHEA ratio (Fig 4.4). At baseline, there was no effect of season (F2,81 = 0.60, p = 0.55), but there was a significant effect of sample type (F2,81 = 12.48, p < 0.0001). The corticosterone/DHEA ratio was greater in plasma than in cmTel or Hp (Tukey’s HSD, p < 0.05). After stress, the main effects of season and sample type were significant (F2,68 = 7.50, p < 0.002; F2,68 = 51.54, p < 0.0001, respectively), as was the interaction between season and sample type (F4,68 = 7.72, p < 0.0001). The corticosterone/DHEA ratio was greater in plasma than in cmTel or Hp, and in plasma, the ratio was greater during the breeding season (Tukey’s HSD, p < 0.05).  Effects of season and stress on steroids in peripheral tissues We used two-factor ANOVAs to examine the effects of season and stress on corticosterone concentrations in peripheral tissues (Table 4.3). In the adrenal glands, there were no effects of season or stress. In the gonads, there was a significant effect of season, with lowest 70  corticosterone concentrations during the non-breeding season. Note that the adrenals in all seasons and the gonads in the nonbreeding season were small, and high steroid concentrations may be, in part, a result of the small tissue mass. In both liver and pectoral muscle, there was an interaction between season and stress, and stress increased corticosterone levels to a greater extent during the breeding season. We also used two-factor ANOVAs to examine the effects of season and stress on DHEA concentrations in peripheral tissues (Table 4.4). In the adrenals and gonads, there were no effects of season or stress on DHEA levels. Again, the adrenals and regressed gonads were small, possibly affecting calculated steroid concentrations. In liver, DHEA levels tended to change seasonally (p = 0.06). Overall high DHEA concentrations in liver suggest that the liver synthesizes DHEA, as in developing rats (Katagiri et al., 1998). In pectoral muscle, DHEA levels were low and changed seasonally.  DISCUSSION Our results reveal dramatically different regulation of steroids in brain tissue versus plasma. First, corticosterone levels were up to 10× lower in brain than in plasma, whereas DHEA levels were up to 5× higher in brain than in plasma. Second, we found strong seasonal changes in baseline corticosterone and DHEA levels in plasma but not in brain. Third, acute stress increased corticosterone levels in plasma but decreased corticosterone levels in the Hp during molt. Also during molt, acute stress increased DHEA levels in jugular plasma but had no effect on DHEA levels in the brain. This is the first study to measure 1) corticosterone or DHEA levels in the brain of adult songbirds and 2) seasonal changes in corticosterone or DHEA levels in the brain of any species. Moreover, our results are from wild animals sampled under natural conditions.  71  Corticosterone levels were typically lower in brain than in plasma. However, the differences between plasma and brain concentrations varied with season, brain region and stress. During the breeding season, baseline and stressed corticosterone levels were up to 10× greater in plasma than in brain. The difference between plasma and brain was most pronounced for NCM at baseline, whereas after stress, the difference was similarly large for Hp, NCM and dTel. During the molt and nonbreeding season, the differences between plasma and brain were less pronounced, particularly at molt when corticosterone concentrations were only 2× greater in plasma. There are several possible explanations for these patterns. First, corticosteroid binding globulin (CBG) binds ~90% of corticosterone in plasma and may restrict its entry into brain tissue (Hammond, 1990; Breuner and Orchinik, 2002). We measured total (free + bound) corticosterone in the plasma, and our measures of corticosterone in brain may primarily reflect free corticosterone in plasma. Plasma CBG levels are reduced at molt (Bruener and Orchinik, 2001; Romero et al., 2006), which may reduce the difference between plasma and brain corticosterone levels. Second, brain levels of 11β-HSD type 2 (11β-HSD2), which converts corticosterone to inactive dehydrocorticosterone (Holmes and Seckl, 2006), could vary seasonally. Third, corticosterone synthesis in the brain could change seasonally and be upregulated at molt (Newman et al., 2008a). The rodent brain expresses the requisite enzymes (Davies and MacKenzie, 2003; Gomez-Sanchez et al., 2005), and neural 11β-hydroxylase expression is upregulated when systemic corticosterone levels are low (Ye et al., 2008). In contrast to corticosterone, DHEA levels were higher in brain than in plasma. During the breeding and nonbreeding seasons, DHEA levels were up to 3× higher in brain than in plasma, across brain regions and stress conditions. During the molt, baseline DHEA concentrations in Hp, NCM and dTel were 5 to 10× higher than plasma concentrations. Overall, DHEA levels were highest in the Hp, consistent with hippocampal synthesis of DHEA. In the male rat Hp, Hojo et al. (2004) detected the mRNA and activity of P450c17, which synthesizes 72  DHEA. P450c17 and other steroidogenic enzymes are also expressed in the avian brain, suggesting that birds have the capacity to synthesize DHEA de novo in the nervous system (London et al., 2006; Tsutsui et al., 2006). It is unlikely that circulating DHEA is sequestered in brain tissue, because seasonal changes in plasma DHEA were not reflected in the brain (in contrast to the liver and pectoral muscle). It is also unlikely that lipoidal or fatty acid esters of DHEA in the brain affected our measures of free DHEA, because our extraction protocol used 90% methanol for elution and this is selective for free steroids (Liere et al., 2004). Plasma corticosterone levels showed dramatic seasonal changes, in stark contrast to brain corticosterone levels. Baseline and stressed corticosterone levels in plasma were elevated during the breeding season and reduced during the molt and nonbreeding seasons, as in previous studies (Romero et al., 2002; Newman et al., 2008a). In other species, seasonal changes in plasma corticosterone levels are due in part to changes in adrenal responsivity to ACTH and pituitary responsivity to CRH and AVT (Romero, 2006). Remarkably, in the brain, there were small or no seasonal changes in baseline or stressed corticosterone levels. Seasonal regulation of plasma CBG levels may account for the lack of seasonal changes in brain corticosterone levels. In house sparrows (Passer domesticus), CBG and total corticosterone levels in plasma change in parallel across the seasons. Thus, estimated free corticosterone levels in plasma (predicted using the equation of Barsano and Baumann, 1989) remain constant across seasons (Breuner and Orchinik, 2001; Romero et al., 2006). If brain corticosterone levels are primarily determined by plasma free corticosterone levels, then the absence of seasonal changes in brain corticosterone levels may reflect the absence of seasonal changes in plasma free corticosterone levels. Thus, our empirical measurements of corticosterone in brain are consistent with estimates of free corticosterone in plasma. Like corticosterone, DHEA levels also changed seasonally in plasma. Baseline plasma DHEA levels are elevated during the breeding and nonbreeding seasons and reduced at molt. 73  DHEA levels in the brain, however, did not change seasonally. These data suggest that neural DHEA synthesis is not seasonally regulated or even up-regulated during seasons when peripheral DHEA synthesis is down-regulated. Synthesis of local steroids and systemic steroids can be regulated independently (Schmidt et al., 2008b). High DHEA levels in the brain may facilitate year-round territorial behavior (Soma et al., 2008). It is possible that brain DHEAsulfate (DHEA-S) changes seasonally. In the brain of an amphibian (Rana nigromaculata), levels of pregnenolone (PREG) remain constant across the seasons, while levels of PREG-S change seasonally (Takase et al., 1999). Future studies should examine DHEA-S levels in songbirds, as done in mammals (Schumacher et al., 2008). Acute stress increased corticosterone in plasma and most brain regions. In molting song sparrows, acute stress increases corticosterone levels to a greater extent in jugular plasma than in brachial plasma (Newman et al., 2008a), suggesting that stress rapidly stimulates brain corticosterone synthesis during molt. However, brain corticosterone levels after 30 min of restraint were not greater at molt than at other seasons. Neurosteroids show very rapid and transient fluctuations (Balthazart and Ball, 2006; Remage-Healy et al., 2008) due in part to rapid changes in steroid synthesizing and metabolizing enzymes. In addition, neurosteroids can passively diffuse or be actively transported into the bloodstream (Pariante, 2008). Future studies should examine earlier timepoints (e.g., 5 and 10 min; Croft et al., 2008). Alternatively, stress may increase corticosterone synthesis in a brain region not studied here. Surprisingly, stress decreased corticosterone levels in the Hp during molt. The Hp has the highest density of glucocorticoid receptors in the brain and is particularly vulnerable to stress (McEwen, 2001). One possible mechanism to reduce exposure of the Hp to glucocorticoids is via 11β-HSD2, which rapidly inactivates corticosterone to dehydrocorticosterone (Holmes and Seckl, 2006; Klusonova et al., 2008). Acute restraint stress (45 min) rapidly upregulates 11β-HSD2 activity in the rat placenta to protect the fetus from high 74  levels of maternal glucocorticoids (Welberg et al., 2005). Future work will determine whether 11β-HSD2 activity is rapidly upregulated by stress in the songbird Hp. During molt, decreased adrenal reactivity may be coupled with increased hippocampal 11β-HSD2 reactivity to protect the Hp from high levels of corticosterone. Acute stress increases jugular, but not brachial, DHEA levels in molting song sparrows (Newman et al., 2008b). These data suggest that stress also stimulates neural DHEA synthesis during molt, when peripheral DHEA synthesis is reduced. Here, we also found that stress increased jugular DHEA concentrations during molt. However, acute stress did not affect neural DHEA levels at molt or at other seasons. As with corticosterone, DHEA levels in jugular plasma may reflect DHEA that is synthesized in the brain but diffuses away before being rapidly metabolized. Alternatively, stress may increase DHEA synthesis in a region of the brain not examined here. Ongoing studies are examining the effects of season and stress on DHEA metabolism throughout the brain. In conclusion, this study of wild animals under natural conditions identified regional variation in steroid concentrations and dynamic changes with season and acute stress. There are pronounced differences in the regulation of corticosterone and DHEA in plasma and brain, especially during molt. These data are consistent with accumulating evidence that neurosteroids may function more like neuromodulators or neurotransmitters than hormones (Balthazart and Ball, 2006; Schmidt et al., 2008). Further, these results highlight the difficulties associated with using circulating steroid levels to infer local steroid levels within the brain. This is one of the first comprehensive comparisons between brain and systemic steroid concentrations, and the results lay the foundation for future work examining the cellular and molecular mechanisms of neurosteroid regulation.  75  TABLES Table 4.1 Effects of season, stress and brain region on corticosterone and DHEA in the brain. Three-way mixed-design ANOVA Corticosterone DHEA Variable df F ratio p df F ratio p Season 2,271 1.39 0.25 2,271 1.22 0.30 Stress 1,271 115.41 < 0.0001 1,271 0.15 0.70 Season x Stress 2,271 2.85 0.06 2,271 0.78 0.46 Region 5,271 6.67 4.18 0.0012 < 0.0001 5,271 Season x Region 10,271 1.93 10,271 1.08 0.38 0.04 Stress x Region 5,271 4.68 5,271 0.28 0.92 0.0005 Season x Stress x Region 10,271 2.21 10,271 1.09 0.37 0.02  76  Table 4.2 Effects of season and sample type on corticosterone and DHEA levels. Two-way mixed-design ANOVA Baseline Stressed Steroid and Variable df F ratio p df F ratio p Corticosterone Season 2,85 0.25 0.78 2,69 12.81 < 0.0001 Sample (plasma, cmTel, Hp) 1,85 14.19 < 0.0001 1,69 62.57 < 0.0001 Season x Sample 2,85 3.22 2,69 3.96 0.02 0.009 DHEA Season 2,85 0. 95 0.40 2,70 0.79 0.46 Sample (plasma, cmTel, Hp) 1,85 13.46 < 0.0001 1,70 3.03 0.03 Season x Sample 2,85 2.31 0.08 2,70 0.38 0.82  77  Table 4.3 Effects of season and stress on corticosterone levels in peripheral tissues. Corticosterone (ng/g) Liver  Muscle  Adrenal  Gonad  Baseline  12501.16 ± 5041.89 (100)  0.83 ± 0.34 (50)  3.95 ± 0.57 (100)  2.88 ± 0.65 (88)  Stressed  9118.41 ± 642.68 (100)  5.22 ± 1.49 (100)  19.08 ± 2.62 (100)  25.66 ± 3.21 (100)  Baseline  12582.35 ± 834.89 (100)  6.74 ± 6.74 (20)  2.99 ± 0.30 (100)  3.17 ± 0.92 (100)  Stressed  8748.86 ± 1254.60 (100)  7.52 ± 5.60 (33)  6.64 ± 0.30 (100)  11.32 ± 2.54 (83)  Baseline  17738.62 ± 635.11 (89)  0±0 (0)  4.08 ± 0.60 (100)  2.82 ± 1.06 (75)  Stressed  12276.43 ± 1978.01 (100)  0±0 (0)  9.32 ± 1.41 (100)  10.88 ± 2.03 (89)  Breeding  Molt  NonBreeding  Twofactor ANOVA  Variable  df  F ratio  p  df  F ratio  p  df  F ratio  p  df  F ratio  p  Season  2,45  1.88  0.17  2,41  5.66  0.008  2,45  7.99  0.0012  2,45  8.12  0.001  Stress  1,45  0.62  0.44  1,41  2.89  0.10  1,45  71.56  <0.0001  1,45  52.85  <0.0001  Season x Stress  2,45  0.29  0.75  2,41  1.63  0.21  2,45  4.86  0.012  2,45  8.28  0.001  Note: Numbers in parentheses indicate the percent of detectable samples.  78  Table 4.4. Effects of season and stress on DHEA levels in peripheral tissues.  DHEA (pg/g) Adrenal  Gonad  Liver  Muscle  Baseline  5099.55 ± 1210.15 (88)  1100.90 ± 207.68 (100)  2538.33 ± 343.48 (100)  796.86 ± 171.38 (100)  Stressed  7975.71 ± 1976.49 (78)  1647.39 ± 330.61 (100)  3203.88 ± 313.10 (100)  788.83 ± 160.32 (100)  Baseline  19380.27 ± 9924.75 (100)  2546.13 ± 1209.52 (60)  2477.42 ± 444.57 (100)  396.05 ± 107.30 (100)  Stressed  2215.62 ± 1105.11 (50)  3467.06 ± 1404.92 (67)  2146.65 ± 462.87 (100)  439.72 ± 62.81 (100)  Baseline  15793.61 ± 5138.04 (89)  42468.00 ± 7335.52 (57)  3550.77 ± 532.34 (100)  488.08 ± 103.88 (88)  Stressed  13255.68 ± 3447.49 (89)  53591.04 ± 18577.67 (88)  3622.00 ± 597.72 (100)  418.26 ± 93.76 (100)  Breeding  Molt  NonBreeding  Variable  df  F ratio  p  df  F ratio  p  df  F ratio  p  df  F ratio  p  Season  2,45  2.17  0.13  2,41  1.55  0.23  2,45  3.01  0.06  2,45  5.15  0.011  Stress  1,45  2.68  0.11  1,41  1.35  0.25  1,45  0.10  0.75  1,45  0.01  0.92  Season x Stress  2,45  2.66  0.08  2,41  0.55  0.58  2,45  0.44  0.64  2,45  0.09  0.91  Two-factor ANOVA  Note: Numbers in parentheses indicate the percent of detectable samples.  79  FIGURES Figure 4.1 Effects of season and acute stress on corticosterone in jugular plasma and brain.  100 50  Molt  50  Brain corticosterone (ng/g)  100  10 5  200 150 100 50  100 100  Jugular Plasma  50 100  38 100  75 100  29  78  25 100  50 100  40 100  0  100  100 100  40  0  40 100  60 100  44 100  56 100  100 100  22  63  22 78  22  rDien  cDien  cmTel  20 15 10 5 0  100 100  Nonbreeding  Plasma corticosterone (ng/mL)  Molt Nonbreeding  15  0  150  0  20  100 100  200  0 Plasma corticosterone (ng/mL)  Breeding  150  Brain corticosterone (ng/g)  200  0  Baseline Stressed  25  Brain corticosterone (ng/g)  Breeding  Plasma corticosterone (ng/mL)  250  20 15 10 5 0  Hp  NCM  78  dTel  Figure 4.1 Effects of season and stress on corticosterone levels in plasma (left) and in brain (right). Corticosterone was regulated by season in jugular plasma but not in brain. Acute stress increased corticosterone in plasma and brain, except during molt, when stress decreased corticosterone levels in Hp. rDien: rostral diencephalon; cDien: caudal diencephalon; cmTel: central medial telencephalon; Hp: hippocampus; NCM: caudal medial nidopallium; dTel: dorsal telencephalon. Numbers below bars indicate the percent of detectable samples.  80  1200  4800  1000  4000  Breeding  800 600 400  Brain DHEA (pg/g)  Plasma DHEA (pg/mL)  Breeding  Figure 4.2 Effects of season and acute stress on DHEA levels in plasma and brain  200  Molt  600 400 200  100 89  100 100  100 100  71 100  86  89  100 100  60  83  80  83  100 100  100 83  60 100  100 100  89  89  100 100  100 100  100 100  89  rDien  cDien  cmTel  Hp  3200 2400 1600 800 0  100 100  1000  4000 Nonbreeding Brain DHEA (pg/g)  Plasma DHEA (pg/mL)  Nonbreeding  1600  4000  800  800 600 400 200 0  2400  0  100 100  1000  0  3200  800  Brain DHEA (pg/g)  Plasma DHEA (pg/mL)  Molt  0  Baseline Stressed  3200 2400 1600 800  100 100  Jugular Plasma  0  78  NCM  100 89  dTel  Figure 4.2 Effects of season and stress on DHEA levels in plasma (left) and in brain (right). DHEA was regulated by season in jugular plasma but not in brain. Acute stress increased plasma DHEA during the molt. Acute stress did not affect DHEA levels in brain. Overall, DHEA levels were highest in the Hp. Numbers below bars indicate the percent of detectable samples.  81  Figure 4.3 Comparison of corticosterone and DHEA in plasma and brain  B 200  20 15  150  10 5  100 0  50  0 Breeding  C  Stressed corticosterone (ng/mL or ng/g)  Baseline corticosterone (ng/mL or ng/g)  A  Jugular Plasma cmTel Hp 150  100  50  0  Molt Nonbreeding  D  4000  3000  2000  1000  0  Breeding  Molt Nonbreeding  Breeding  Molt Nonbreeding  5000  Stressed DHEA (pg/mL or pg/g)  5000  Baseline DHEA (pg/mL or pg/g)  200  4000  3000  2000  1000  0 Breeding  Molt Nonbreeding  Figure 4.3 Baseline and stressed corticosterone (A, B) and DHEA (C,D) in plasma, cmTel, and Hp across seasons. Baseline and stressed corticosterone levels were higher in plasma than in cmTel or Hp. Inset in 3A depicts baseline corticosterone levels on an expanded y-axis scale. Baseline and stressed DHEA levels were lower in plasma than in cmTel or Hp.  82  Figure 4.4 Effects of season and acute stress on corticosterone / DHEA ratio  A Baseline corticosterone / DHEA  30  Plasma cmTel Hp  25 20 15 10 5 0 Breeding  Molt  Nonbreeding  Breeding  Molt  Nonbreeding  B Stressed corticosterone / DHEA  300 250 200 150 100 50 0  Figure 4.4 Effects of season and stress on the baseline (A) and stressed (B) corticosterone / DHEA ratio in jugular plasma, cmTel and Hp. Baseline and stressed plasma corticosterone / DHEA ratios were greater than the ratios in cmTel and Hp. Baseline plasma ratios did not differ significantly across seasons, but the stressed plasma ratio was greater during the breeding season than during the molt and nonbreeding season.  83  REFERENCES Apostolova, G., Schweizer, R.A.S., Balazs, Z., Kostadinova, R.M., Odermatt, A., 2005. Dehydroepiandrosterone inhibits the amplification of glucocorticoid action in adipose tissue. 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Pregnenolone, pregnenolone sulfate, and cytochrome P450 side-chain cleavage enzyme in the amphibian brain and their seasonal changes. Endocrinology 140, 1936-1944. Thoeringer, C.K., Sillaber, I., Roedel, A., Erhardt, A., Mueller, M.B., Ohl, F., Holsboer, F., Keck, M.E., 2007. The temporal dynamics of intrahippocampal corticosterone in response to stress-related stimuli with different emotional and physical load: An in vivo microdialysis study in C57BL/6 and DBA/2 inbred mice. Psychoneuroendocrinolo 32, 746-757. Torres, J.M., Ortega, E., 2003. DHEA, PREG and their sulphate derivatives on plasma and brain after CRH and ACTH administration. Neurochem Res 28, 1187-1191. Tramontin, A.D., Brenowitz, E.A., 2000. Seasonal plasticity in the adult brain. Trends Neurosci 23, 251-258. Tsutsui, K., Matsunaga, M., Miyabara, H., Ukena, K., 2006. Neurosteroid biosynthesis in the quail brain: a review. J Exp Zoolog A Comp Exp Biol 305A, 733-742. Welberg, L. A. M., Thrivikraman, K.V., Plotsky, P.M., 2005. Chronic maternal stress inhibits the capacity to up-regulate placental 11β-hydroxysteroid dehydrogenase type 2 activity. J Endocrinol 186, 7-12. Wingfield, J.C., Hahn, T.P., 1994. Testosterone and territorial behaviour in sedentary and migratory sparrows. Anim Behav 47, 77-89. Ye, P., Kenyon, C.J., MacKenzie, S.M., Nichol, K., Seckl, J.R., Fraser, R., Connell, J.M.C., Davies, E., 2008. Effects of ACTH, dexamethasone, and adrenalectomy on 11β-hydroxylase (CYP11B1) and aldosterone synthase (CYP11B2) gene expression in the rat central nervous system. J Endocrinol 196, 305-311.  88  5  EFFECTS OF CORTICOSTERONE AND DHEA ON NEURAL PLASTICITY IN THE ADULT SONGBIRD BRAIN 4  INTRODUCTION Neurogenesis continues during adulthood across vertebrates. In mammals, cell proliferation is high in the subventricular zone (SVZ) and dentate gyrus, from which neuronal progenitor cells migrate to the olfactory bulb and granular layer of the hippocampus, respectively (AlvarezBullya et al., 2002; Cameron et al., 1993). In birds, neuronal progenitor cells migrate from the ventricular zone (VZ) to many areas throughout the telencephalon (Goldman, 1998; AlvarezBullya and Kirn, 1997; Nottebohm, 2002). Songbirds have high rates of neuronal turnover in nuclei of the song control system (Nottebohm, 1981; Alvarez-Buylla and Kirn, 1997). The steroidal milieu in the brain influences the production and survival of new neurons during adulthood (Galea, 2008). Glucocorticoids are potent steroidal modulators of adult neurogenesis and are secreted by the adrenal glands following activation of the hypothalamicpituitary-adrenal (HPA) axis by various stressors. In adult rats, corticosterone treatment in vivo decreases neuronal proliferation and survival in the dentate gyrus (Gould et al., 1992; Karishma and Herbert, 2002; Wong and Herbert, 2004). In adult zebra finches (Taeniopygia guttata), corticosterone treatment in vitro decreased cell proliferation in the VZ (Katz et al., 2008). However, in adult chickadees (Poecile atricapillus), corticosterone treatment in vivo did not affect hippocampus size or cell number, or cell proliferation in the VZ (Pravosudov and Omanska, 2005). Effects of corticosterone treatment on HVC (HVC used as proper name, not an acronym) size and recruitment of new cells into HVC in adult songbirds have not been reported.  4  A version of this chapter will be submitted for publication. Newman, A.E.M., MacDougall-Shackleton, S.A., An, Y.S., Kriengwatana, P., Soma, K.K. Effects of corticosterone and DHEA on neural plasticity in the adult songbird brain.  89  Dehydroepiandrosterone (DHEA) is an androgen precursor that, like glucocorticoids, is increased by acute stress or ACTH in the brain (Corpechot et al., 1981; Torres and Ortega, 2003) or plasma (Oberbeck et al., 1998; Arvat et al., 2000), depending on the species examined. DHEA has various effects on the brain (Maninger et al., 2009) and, for example, increases HVC size in adult song sparrows (Soma et al., 2002). DHEA has been described as a native antiglucocorticoid in the nervous system (Kalimi et al., 1994; Maninger et al., 2009). Specifically, DHEA prevents corticosterone-induced translocation of stress-activated protein kinase 3 to the cell nucleus in rat hippocampal cells in vitro (Kimoinides et al., 1999). In vivo, corticosterone suppresses the recruitment of new cells into the adult male rat dentate gyrus, and this corticosterone effect is prevented by DHEA treatment (Karishma and Herbert, 2002). Together, these studies suggest that DHEA is regulated by stress and modulates the effects of glucocorticoids on the brain. However, because rats have very low levels of circulating DHEA, the effects of high DHEA doses in this species are difficult to interpret. Unlike rats, songbirds have relatively high levels of circulating DHEA (Soma and Wingfield, 2001; Hau et al., 2004; Newman et al., 2008b). In song sparrows (Melospiza melodia), DHEA levels in jugular plasma that is exiting the brain, but not in brachial plasma, are affected by acute restraint stress (Newman et al., 2008b). Also, DHEA levels are much higher in song sparrow brain than plasma, and DHEA concentrations are highest in the hippocampus (Newman et al., in review), a brain region particularly sensitive to glucocorticoids (McEwen, 2001). Together with studies in rodents, these data raise the hypothesis that DHEA reduces the effects of glucocorticoids on the brain. Here, we treated adult male song sparrows with corticosterone and/or DHEA and assessed the effects on gross neuroanatomy and recruitment of new cells into HVC, hippocampus and the rest of the telencephalon.  90  METHODS Experiment 1: Effects of corticosterone and DHEA on neuroanatomy and new cell recruitment Subjects Subjects were wild adult male song sparrows (n = 36 total) captured at the Queens University Biological Station (44°34’N, 76°19.5’W) (n = 12) or near the University of Western Ontario (43°0.5’N, 81°16.8’W) (n = 24) (June 4th – July 6th, 2007). Subjects were housed in individual cages in 4 separate colony rooms (n=9 per room) with ad libitum access to water, food (Mazuri maintenance diet + white millet) and grit and on kept a natural photoperiod. Subjects were then gradually shifted to a short day photoperiod of 8L:16D and maintained on 8L:16D for at least one month prior to steroid treatments. We randomly assigned subjects to one of four treatment groups: empty implant + empty implant (control), corticosterone implant + empty implant (corticosterone), DHEA implant + empty implant (DHEA), or corticosterone implant + DHEA implant (corticosterone + DHEA). Steroid treatments started from November 3rd to December 7th, 2007, depending on the colony room. Hereafter, we refer to the day of steroid implantation as day 1 (Fig 5.1A). On day 1, subjects were anaesthetized (1.5% isoflurane, 2 L/min O2) and given silastic implants (see below) subcutaneously on the back. Implants were inserted through a small incision in the skin, which was then sealed with veterinary adhesive (Nexaband S/C).  Steroid Treatment Implants were made from silastic tubing (for corticosterone: i.d. 1.47mm, o.d. 1.96mm; for DHEA: i.d. 0.76mm, o.d. 1.65mm). For corticosterone, silastic implants with effective lengths of 10-15mm have been used to elevate circulating corticosterone levels in similarly-sized songbirds (Astheimer et al., 2000; Breuner & Hahn, 2003; Martin et al., 2005). Thus we used an effective length of 12mm. Implants were packed with crystalline corticosterone (Sigma) and 91  sealed with liquid silastic. Previous studies have shown that it is necessary to facilitate corticosterone release from silastic implants by making a small hole in the implant (Silverin, 1998; Astheimer et al., 2000). We used a 22 gauge needle to poke one hole in each corticosterone implant at the end of the effective length (i.e. not through the liquid silastic plug). For DHEA, we used an effective length of 7mm, which elevates plasma DHEA levels in song sparrows within the physiological range (Soma et al., 2002). Implants were packed with crystalline DHEA (Steraloids, Newport, RI, USA) and sealed with liquid silastic.  BrdU Injections & Blood Collection On day 4 and day 5, all subjects received 3 intramuscular injections of bromo-deoxyuridine (BrdU) in 0.1M phosphate-buffered saline (PBS; pH = 7.4) (100 µL volume; 0.015g/mL BrdU in saline). We used a dose of 65 mg/kg, similar to previous songbird studies (Hoshooley et al., 2007). Injections were administered at 0900, 1300, and 1700 on day 4 and day 5. All baseline blood samples were collected from the brachial vein within 3 min of entering the room. Three days prior to implantation (day -2), a baseline blood sample (~100 µL) was collected. On days 8 and 22, a baseline blood sample (~100 µL) was collected, and subjects were then restrained in an opaque paper bag for 30 min. After restraint, another blood sample was collected (~100 µL). On day 29, a final baseline blood sample (~100 µL) was collected prior to sacrifice. Blood was kept on ice until centrifuged, and plasma was stored at 20°C. After blood collection on day 29, subjects were deeply anaesthetized with ketamine and xylazine and transcardially perfused with heparinized saline followed by buffered 4% paraformaldehyde (pH = 8.5). Brains were removed from the skull, postfixed in 4% paraformaldehyde for 24hr, and then cryoprotected in 30% sucrose until saturated (~48hr). Brains were then frozen on pulverized dry ice and stored at -80°C. 92  Immunocytochemistry Brains were sectioned in the coronal plane at 30µm thickness on a cryostat. Two sets of brain sections were collected into PBS for immunochemistry to detect BrdU- and NeuNimmunoreactivity, and a third set was collected into cryoprotectant and stored at -20°C. To detect BrdU immunoreactivity (BrdU+ cells; Fig 5.2), free-floating sections were washed in PBS and then immersed in 2N HCL for 40 min. Sections were washed in 0.1 M sodium borate (with 0.5% HCl) for 10 min and then washed in PBS. Sections were washed in 0.3% H2O2 for 30min and then washed in PBS. Sections were blocked in 10% normal goat serum (Vector) for 30 min and then exposed to the anti-BrdU antibody (mouse anti-BrdU/IdU, Caltag MD5000, 1:500 in PBS with 0.3% Triton) for 20 hr at room temp. This antibody has been used previously in chick brain at a 1:500 dilution (Carnahan et al., 1994), and in pilot studies, we tested a range of dilutions on song sparrow brain sections. In pilot studies, we also tested other BrdU antibodies (from BD Pure, BD Pharmingen, Chemicon, Caltag) and chose the antibody that gave minimal background staining and maximal BrdU-labelled cells. Next, sections were washed in PBS with 0.1% Triton and then exposed to the biotinylated secondary antibody (1:250 in PBS with 0.3% Triton; goat anti-mouse IgG, Vector) for 1hr. Sections were washed in PBS with 0.1% Triton, and then treated with avidin-biotin-peroxidase reagent (Vector Elite ABC kit) for 1 hr. BrdU labelling was visualized by exposing the sections to a diaminobenzidine solution (Sigma FastDAB). Sections were then washed thoroughly in PBS, mounted on glass slides, dehydrated through a series of graded ethanols, cleared in solvent (Harleco Neo-Clear, EMD Chemicals) and coverslipped using Neo-mount (EMD Chemicals). To detect NeuN immunoreactivity (Fig 5.2), sections were processed similarly; however, sections were not exposed to HCl or sodium borate and were exposed to anti-NeuN primary antibody (mouse antiNeuN, Chemicon MAB377, 1:2000 in PBS with 0.3% Triton) for 20 hr at room temp. As a negative control, 20 sections from different subjects were processed for BrdU and NeuN 93  staining with the primary antibody omitted. Nuclear (for BrdU) and cytoplasmic (for NeuN) staining were absent in these sections.  Neuromorphometry and Cell Counting All brain measurements were made blind to the treatment group. To obtain estimates of total telencephalon volume, slides with NeuN-labelled sections were scanned into a computer using a high-resolution (2400 dpi) flat bed scanner. The telencephalon area was measured from every 12th section using NIH ImageJ software (Wayne Rasband, National Institutes of Health, Bethesda, MD); volume was calculated by multiplying the area of each section by the sampling interval (360 µm). The left and right sides of the telencephalon did not differ (left: 251.4 ± 3.9 mm3; right: 254.5 ± 3.7 mm3), as in previous studies (e.g. Tramontin et al., 2000). To estimate HVC and hippocampus volume, we captured images from every NeuN-labelled section containing the HVC (90 µm apart, ~12 sections per subject) and from every 12th section containing the hippocampus (360 µm apart, ~20 sections per subject) using a Leica Digital CCD camera mounted on a Leica DM5000 B light microscope through a 10× objective lens. Leica Application Suite software was used to make area measurements of HVC or hippocampus in one hemisphere; volume was calculated by multiplying the HVC or hippocampus area of each section by the sampling interval (90 µm and 360µm). Values were then multiplied ×2 to obtain an estimate of total volume (both hemispheres). To estimate the number of HVC neurons, we used the unbiased stereology optical fractionator method (Glaser et al., 2007) on sections labelled for NeuN. In one hemisphere, cells were counted in every 9th section that contained HVC (i.e. 270µm apart). Under a 20× objective, we used the Leica Application Suite software driving a motorized stage to cover HVC with a 270 × 270 µm sampling grid. Then, under a 63× oil objective, the section was moved to the first square in the sampling grid. At each point in the sampling grid (270µm apart), the 94  counting frame was 30 × 30 µm. We measured the thickness of the tissue section by focusing on its top and bottom edges in 3 counting frames per section. We did not count cells in the top and bottom 1 µm of the section (guard zones) except in a few cases where the section was ≤ 7 µm thick, in which case we counted all of the cells. Due to shrinkage during processing, the section thickness was 7.7 ± 0.3 µm (range 6-11 µm). We focused through the section by 1 µm intervals and counted all of the cells that came into focus within the 5 µm thick x 30 µm2 threedimensional slab. The total population of cells in one half of HVC was then estimated using the following formula:  Total population = n x (1/ssf) x (1/asf) x (1/hsf)  (1)  where n = total number of cells counted, ssf = section sampling frame (i.e. 1/9 as we counted every 9th section), asf = area sectioning frame (302µm/2702µm), hsf = optical dissector height (5µm)/tissue section thickness. The total was multiplied ×2 to obtain an estimate of total HVC neuron number (both hemispheres). To estimate the number of BrdU+ cells in HVC, in the hippocampus, along the ventricular zone, and in the telencephalon (not including HVC and hippocampus), BrdU+ cells were counted exhaustively in one hemisphere of the brain. Exhaustive sampling was used rather than a stereological approach because BrdU+ cells were relatively infrequent, and BrdU+ cell nuclei were small relative to the section thickness. In such cases, exhaustive sampling is more accurate than sampling from a small subset of sections, and the risk of sampling bias is low when objects are small compared to section thickness. For all 4 regions, BrdU+ cells were counted in all sections containing HVC (90 µm apart). In the rest of the brain (rostral and caudal to HVC), BrdU+ cells were counted in every 12th section (360 µm apart). The number of  95  counted BrdU+ cells was then multiplied by the section interval (i.e. 12 or 3) and subsequently multiplied ×2 to obtain an estimate for both hemispheres.  Experiment 2: Effects of corticosterone implants on plasma corticosterone In Experiment 1, plasma corticosterone concentrations were not significantly elevated on day 8 in the corticosterone group (see Results). Thus, to clarify the initial effects of the corticosterone implants on plasma corticosterone levels, we used a second group of subjects to measure plasma corticosterone concentrations during the first week of treatment (Fig 5.1B). Treatment of the two subject groups was similar; however, subjects in Experiment 2 were not injected with BrdU.  Subjects Wild adult male song-sparrows in non-breeding condition (n=12) were captured in August 2008 from around Guelph, Canada (43°33’N, 80°15’W) and the University of Western Ontario. Subjects were maintained in captivity under identical conditions (photoperiod, housing, food) as the subjects in Experiment 1. On day 1, subjects received corticosterone-filled silastic implants (prepared exactly as in Experiment 1, n=6) or empty silastic implants (n=6) subcutaneously on the back. Three subjects in the control group and one subject in the corticosterone group either continued or began molting feathers during the experiment, and these molting subjects were excluded because molt dramatically alters stress physiology in song sparrows and other songbirds (Romero, 2006; Newman et al., 2008b and in revision).  Blood collection Three days prior to implantation (day -3), a baseline blood sample (~50µL) was collected from the brachial vein within 3 min of entering the room. Baseline blood samples were also collected on days 2, 3, 4 and 5 (~25µL on each day). On days 8 and 22, a baseline blood sample (~50µL) 96  was collected, and another blood sample (~50µL) was collected after 30 min of restraint (as in Experiment 1). On day 29, a final baseline blood sample was collected. Blood was kept on ice until centrifuged, and plasma was stored at -20°C. After blood collection on day 29, individuals were sacrificed by rapid decapitation (within 3 min of disturbance), and the brain was rapidly dissected, frozen on dry ice, and stored at -80°C.  Steroid Measurement (Experiments 1 & 2) Plasma corticosterone levels were measured without an extraction step, and plasma DHEA levels were measured after extraction with dichloromethane (as described in Newman et al., 2008a). We used sensitive and specific radioimmunoassays to measure corticosterone (ImmuChem 07-120103, MP Biomedicals, Orangeburg, NY) and DHEA (DSL 8900, Diagnostic Systems Laboratories, Webster, TX). These assays have been validated extensively for use with song sparrow plasma (Newman et al., 2008a,b; Newman and Soma, in press). The lowest points on the corticosterone and DHEA standard curves were 3.12pg corticosterone and 2pg DHEA (per tube). For corticosterone, intra-assay variation was 5.9%, and interassay variation was 9.7% (low control) and 7.3% (high control) (n = 5 assays). Recovery of 75pg of exogenous corticosterone from a pool of song sparrow plasma (n = 6 replicates) was 89%. For DHEA, intra-assay variation was 9.5%, and interassay variation was 5.2% (low control) and 11.7% (high control). Recovery of 50pg of exogenous DHEA from a pool of song sparrow plasma (n = 6 replicates) was 143%. Plasma concentrations were corrected for recovery.  Statistics The effects of corticosterone treatment and DHEA treatment on telencephalon, HVC and hippocampus volume, HVC neuron number, and BrdU+ cell numbers were assessed using 2factor ANOVAs and Tukey’s honestly significant difference (HSD) post hoc tests. The effects 97  of steroid treatments and treatment duration (sampling day) on plasma corticosterone and DHEA levels were analyzed using 2-factor mixed-design ANOVAs in Experiment 1, where treatment duration was a within-subjects factor and treatment group was a between-subjects factor (as defined in O’Rourke et al., 2005). A 2-factor mixed-design ANOVA was also used to analyze the effect of acute restraint stress on plasma steroid levels, where stress was a withinsubjects factor and treatment group was a between-subjects factor. The effects of stress were measured on day 8 and day 22. In Experiment 2, due to small sample sizes, the effect of corticosterone implants on plasma corticosterone levels was analyzed for each treatment day using Welch’s t-tests. The effects of acute restraint stress on treatment days 8 and 22 were calculated using difference scores (stress – baseline). The effect of corticosterone treatment on the difference scores was analyzed on day 8 and 22 using Welch’s t-tests. For both experiments, data were log transformed, if necessary to achieve homogeneity of variance. All data were analyzed with JMP IN 5.1 (SAS, Cary, NC). Data are presented as mean ± SEM.  RESULTS Neural attributes HVC. Corticosterone and DHEA treatment had significant effects on HVC (Table 5.1). For HVC volume (Fig 5.3A), number of HVC neurons (Fig 5.3B), and the number of BrdU+ cells in HVC (Fig 5.3C) there were main effects of both corticosterone and DHEA treatment, but there was no interaction effect. Corticosterone significantly decreased HVC volume, and DHEA significantly increased HVC volume Telencephalon. For the volume of the entire telencephalon, there were no main effects of corticosterone treatment or DHEA treatment and no interaction (Fig 5.4A). For telencephalon BrdU+ cell number, there was no main effect of corticosterone, but there was a 98  main effect of DHEA and a significant interaction effect. BrdU+ cells were more abundant in the telencephalon of the DHEA group than all other groups (Tukey’s HSD, p < 0.05; Fig 5.4B). DHEA treatment increased recruitment of new cells into the telencephalon, but only in the absence of corticosterone treatment. Stated another way, corticosterone treatment had no effect on recruitment of new cells into the telencephalon, except in the presence of DHEA treatment. Hippocampus. For hippocampus volume, there were no main effects of corticosterone or DHEA, and no interaction effect (Table 5.1, Fig 5.5A). For hippocampal BrdU+ cell number, there was no main effect of corticosterone, but there was a main effect of DHEA, and a significant interaction effect (Table 5.1; Fig 5.5B). Similar to the telencephalon (see above), BrdU+ cells were more abundant in the hippocampus of the DHEA group than all other groups (Tukey’s HSD, p < 0.05). Lateral ventricle. For BrdU+ cell number along the lateral ventricle, there was a main effect of corticosterone, no main effect of DHEA, and a significant interaction effect (Table 5.2). BrdU+ cells were more abundant along the lateral ventricle in the DHEA group than all other groups (Tukey’s HSD, p < 0.05; Fig 5.6), as in the telencephalon and hippocampus.  Plasma steroid levels Baseline plasma corticosterone levels. In Experiment 1, the first measurement of baseline plasma corticosterone after implantation was on day 8 (Fig 5.1A) and baseline plasma corticosterone levels (Table 5.2) were not significantly affected by treatment (F3,138 = 0.99, p = 0.40) or sampling day (F3,138 = 1.87, p = 0.14) and there was no interaction (F9,138 = 1.25, p = 0.27). In Experiment 2 however, baseline plasma corticosterone levels were significantly higher in the corticosterone group than controls on day 2 and day 3 and tended to be higher on day 4 (Table 5.3). Baseline plasma corticosterone levels were not significantly different on subsequent days, similar to results from Experiment 1. 99  Baseline plasma DHEA levels. In Experiment 1, for baseline plasma DHEA levels (Table 5.4), there was a significant interaction between treatment and sampling day (F9,138 = 3.46, p = 0.0008). Post hoc tests revealed that plasma DHEA levels were higher in the DHEA and corticosterone + DHEA groups on days 8, 22 and 29, relative to control and corticosterone groups. Even in DHEA treated groups, plasma DHEA levels were within the physiological range for this song sparrow population (1.2 ng/mL to 17 ng/mL). Stressed plasma corticosterone levels. In Experiments 1 and 2, the effect of acute restraint stress was measured on day 8 and day 22. In Experiment 1 on day 8, there was a significant interaction between stress and treatment (F3,61 = 4.50, p = 0.007). Post hoc tests revealed that acute restraint stress increased plasma corticosterone levels in all groups except the corticosterone group. In Experiment 1 on day 22, there was a significant interaction between stress and treatment (F3,67 = 2.96, p = 0.04) . Post hoc tests revealed that acute restraint stress increased plasma corticosterone levels in all groups and that stressed plasma corticosterone levels were greater in the controls than the corticosterone group. In Experiment 2 on day 8, the stress-induced increase in plasma corticosterone levels was significantly greater in controls than the corticosterone group (t = 3.14, p = 0.04). Similarly, in Experiment 2 on day 22, the stressinduced increase in plasma corticosterone levels was significantly greater in controls than the corticosterone group (t = 2.59, p = 0.05). Stressed plasma DHEA levels. For stressed plasma DHEA levels on day 8 and day 22, there was a main effect of treatment (Day 8: F3,67 = 51.29, p < 0.0001; Day 22: F3,67 = 24.96, p < 0.0001) but no effect of stress and no interaction effect. Post hoc tests revealed that on both days, plasma DHEA levels were greater in the DHEA and corticosterone + DHEA groups compared to the control and corticosterone groups.  100  DISCUSSION This study is the first to report the combined effects of corticosterone and DHEA on adult songbird brains. Corticosterone treatment decreased HVC volume, decreased HVC BrdU+ cell number, and tended to decrease HVC neuron number. In contrast, DHEA treatment increased HVC volume, increased HVC BrdU+ cell number, and increased HVC neuron number. Corticosterone and DHEA had additive effects on HVC volume and HVC BrdU+ cell number, but in other regions, corticosterone and DHEA had interactive effects. For example, along the lateral ventricle, in the hippocampus, and in the telencephalon, DHEA treatment increased BrdU+ cell number only in the absence of corticosterone treatment.  Corticosterone and DHEA effects on plasma steroids In Experiment 1, corticosterone treatment did not increase baseline plasma corticosterone levels on days 8, 22, or 29. In recent studies on birds, corticosterone implants increased baseline plasma corticosterone levels only during the first 2-5 days of treatment (Angelier et al., 2007; Muller et al., 2008). Thus, in Experiment 2, we examined the effect of corticosterone implants during the first 2-5 days of treatment. Baseline plasma corticosterone levels were elevated in the corticosterone group on day 2 and day 3. On day 4, baseline plasma corticosterone tended to be higher in the corticosterone group (p = 0.06), but plasma corticosterone levels in controls began to increase, likely due to repeated daily blood collection. On day 5, baseline plasma corticosterone levels in the controls and corticosterone group were not different. On day 8, plasma corticosterone levels in both groups had declined and remained similar. By day 8, the corticosterone implants may have stopped releasing corticosterone, suppressed endogenous corticosterone secretion, and/or increased hepatic clearance of circulating corticosterone. Consistent with the idea that corticosterone was reduced via negative feedback mechanisms, acute restraint stress on day 8 did not increase plasma corticosterone levels in the 101  corticosterone group in Experiment 1 or 2. Similarly, in developing kestrels (Falco tinnunculus), plasma corticosterone levels did not increase in response to acute restraint stress on day 8 after corticosterone implantation, in contrast to controls (Muller et al., 2008). It thus seems likely that chronic treatment with exogenous corticosterone suppresses endogenous corticosterone secretion. DHEA treatment significantly increased plasma DHEA levels throughout the course of Experiment 1 (DHEA was not measured in Experiment 2). Baseline plasma DHEA levels before implantation ranged from 1.2 ng/mL to 17 ng/mL, and thus DHEA treatment produced levels within the physiological range. Interestingly, plasma DHEA levels were much higher in this eastern population of song sparrows than western populations (Soma et al., 2002; Goodson et al., 2005; Newman et al., 2008b). Despite population differences in baseline plasma DHEA levels, DHEA implants increased baseline plasma levels by 2-3× and DHEA levels in treated subjects were within the physiological range for their respective population.  Corticosterone and DHEA effects on neural morphometry and neuron number Corticosterone treatment decreased HVC volume by ~ 25% and tended to decrease HVC neuron number. In contrast, corticosterone did not affect hippocampus volume or overall telencephalon volume. These results suggest that HVC is particularly sensitive to glucocorticoid exposure in songbirds. Also, zebra finches treated with corticosterone during early development have reduced HVC volume in adulthood (Buchanan et al., 2004). Moreover, stressors during early development, such as parasite infection and food restriction, have been shown to result in reduced HVC volume in adulthood (Spencer et al., 2005; MacDonald et al., 2006). How corticosterone affects HVC volume remains unclear. Glucocorticoid receptors are expressed in the songbird brain (Breuner and Orchinik, 2001; Katz et al., 2008), but whether they are expressed in HVC is unknown. These effects of corticosterone on HVC are similar to the 102  effects of corticosterone on the rat dentate gyrus, where corticosterone decreases neuron survival (Sapolsky et al., 1985; Karishma and Herbert, 2002). In captive song sparrows, DHEA treatment increased HVC volume by ~35%. In wild non-breeding song sparrows, DHEA treatment (for 2 weeks) appears to have slightly greater effects on HVC growth (Soma et al., 2002). In the present study, DHEA did not affect hippocampus volume or telencephalon volume. DHEA treatment also increased the HVC neuron number by ~26%. Although there is no known classical intracellular steroid receptor for DHEA (Widstrom and Dillon, 2004), DHEA is converted to androgens and estrogens in the adult song sparrow brain (Pradhan et al., in review) and these potent metabolites of DHEA can bind to androgen and estrogen receptors in HVC (Tramontin et al., 2003). Treatment with testosterone or estradiol increases HVC volume in song sparrows and white-crowned sparrows (Soma et al., 2004; Tramontin et al., 2003). Estradiol treatment increases HVC neuron number in canaries (Serinus canaria) (Hidalgo et al., 1995). HVC volume and neuron number show dramatic seasonal changes in adult song sparrows and are maximal during breeding, when plasma levels of testosterone and estradiol are maximal (Smith et al., 1997; Soma et al., 1999). Thus the effects of DHEA may be mediated through its androgenic and estrogenic metabolites. Corticosterone and DHEA treatments had opposite effects of HVC volume and neuron number, and with regard to HVC volume, corticosterone and DHEA had additive effects, rather than interactive effects. Corticosterone and DHEA could have additive effects on HVC volume by acting via separate mechanisms or by converging on the same mechanism. For example, corticosterone and DHEA could regulate different neurotrophic factors. Brain derived neurotropic factor (BDNF) increases neuron survival in HVC (Rasika et al., 1994), and in rat hippocampus, BDNF is decreased by corticosterone (Jacobsen and Mork, 2006) but not affected by DHEA (Gubba et al., 2004; Pinnock et al., in press). Alternatively, corticosterone and DHEA may converge on the same mechanism, such as the phosphatidylinositol-3 kinase /Akt 103  (PI3/Akt) pathway, which when phosphorylated, decreases neuronal apoptosis. In cultured rat neurons, corticosterone reduces phosphorylation of Akt and increases apoptosis (Nitta et al., 2004), while DHEA increases phosphorylation of Akt and decreases apoptosis (Zhang et al., 2002). Future studies should quantify corticosterone and DHEA induced changes in apoptosis.  Corticosterone and DHEA effects on BrdU+ cell numbers As with HVC volume and neuron number, corticosterone decreased the BrdU+ cell number in HVC, whereas DHEA increased the BrdU+ cell number in HVC. The change in BrdU+ cell number was more dramatic than the change in HVC volume or neuron number. Corticosterone decreased BrdU+ cells in HVC by ~60%, and DHEA increased BrdU+ cell number in HVC by ~80%. Again, the effects of corticosterone and DHEA on BrdU+ cell number in HVC were additive and could be mediated by similar mechanisms mentioned above. In contrast to effects in HVC, the effects of corticosterone and DHEA on BrdU+ cell number along the lateral ventricle, in the hippocampus, and rest of the telencephalon were not additive. In these 3 regions, corticosterone treatment alone did not affect BrdU+ cell number, but corticosterone did prevent the DHEA-induced increase in BrdU+ cell number. These results differ from previous studies in which corticosterone reduced BrdU+ cells in the rodent dentate gyrus (Cameron and Gould, 1994; Wong and Herbert, 2004) and lateral ventricle of the songbird (Katz et al., 2008). Note that in Experiment 1, we used multiple BrdU injections over 2 days, which likely increased endogenous corticosterone levels in all groups, including those not treated with corticosterone. In Experiment 2, repeated handling and blood collection during days 2-5 increased plasma corticosterone levels in the control group. Elevated endogenous plasma corticosterone levels in the control group may have obscured the effects of exogenous corticosterone on BrdU+ cell number. DHEA treatment alone may have been sufficient to increase BrdU+ cell number even during periods of elevated endogenous corticosterone, 104  however, exogenous corticosterone treatment inhibited the effect of DHEA. It is also possible that these areas of the avian brain, unlike HVC, may not be sensitive to corticosterone treatment itself. In the chickadee hippocampus, corticosterone treatment did not affect volume or BrdU+ cell number (Pravasudov and Omanska, 2005), even though glucocorticoid receptors are expressed in the songbird hippcampus (Hodgson et al., 2007). In the absence of exogenous corticosterone, DHEA could increase BrdU+ cell number by decreasing local levels of corticosterone via modulation of 11β-hydroxysteroid dehydrogenase (11β-HSD) isozymes. This may be consistent with the observation of increased cell proliferation in the rodent brain after adrenalectomy (Cameron and Gould, 1994). For example, 11β-HSD type 1 mediated regeneration of active glucocorticoids from inactive metabolites is reduced when the concentrations of 7-hydroxy DHEA derivatives are elevated as these derivatives are the preferred substrate for 11β-HSD1 (Muller et al., 2006 ab). In peripheral tissues, DHEA also modulates 11β-HSD by decreasing expression of 11β-HSD type 1 (Apostolova et al., 2005) and increasing the activity of 11β-HSD type 2 (Balazs et al., 2008). Alternatively, DHEA may act on neurotransmitter systems. In adult male rats, DHEA potentiates fluoxetine induced cell proliferation in the dentate gyrus and this effect is abolished by corticosterone treatment (Pinnock et al., in press). Also, DHEA or its metabolites might also increase expression of several growth factors which regulate neurogenesis (Cameron et al., 1998). In canaries, testosterone increases vascular endothelial growth factor (VEGF) and BDNF to support the migration of new neurons (Louissaint et al., 2002). Although corticosterone treatment alone did not affect BrdU+ along the lateral ventricle, in the hippocampus and in the telencephalon, corticosterone treatment prevented the DHEAinduced increase in BrdU+ cell number. Corticosterone could blunt the effects of DHEA on BrdU+ cell number by locally inactivating DHEA or its metabolites. For example, in rodents, expression of sulfotransferase 2, an enzyme that converts free hydroxysteroids (DHEA, 105  testosterone, estradiol) to their inactive sulphated forms, is induced by dexamethasone, a potent synthetic glucocorticoid (Runge-Morris et al., 1999; Gong et al., 2008). Alternatively, corticosterone might decrease DHEA metabolism by acting on 3β-hydroxysteroid dehydrogenase (3β-HSD) activity. However, 3β-HSD inhibition did not potentiate corticosterone-induced decreases in cell proliferation in the zebra finch brain (Katz et al., 2008), but corticosterone may decrease 3β-HSD over the long-term and influence cell survival. Future studies are required to determine if chronic corticosterone and DHEA treatment alter DHEA metabolism and/or inactivation in the songbird brain.  DHEA as a corticosterone antagonist? Interestingly, none of the data presented here provide evidence that DHEA inhibits the actions of corticosterone in the songbird brain. Further, DHEA treatment did not decrease baseline or stressed plasma corticosterone levels, thus, DHEA does not appear to decrease secretion or increase systemic clearance of corticosterone. This differs from other studies in which DHEA and estradiol limit the actions of glucocorticoids by regulating 11β-HSDs in peripheral tissues (Jamieson et al., 1999; Gomez-Sanchez et al., 2003; Apostolova et al., 2005; Balazs, 2008). Further work is needed to understand the action of DHEA on 11β-HSD in the songbird brain; first to determine if DHEA modulates 11β-HSD activity in the brain and if so, to determine if DHEA-induced changes to 11β-HSD activity in the brain result in changes in local glucocorticoid levels.  Conclusions In conclusion, this study of wild-caught songbirds demonstrated independent effects of corticosterone and DHEA on the volume, neuron number and recruitment of new cells into HVC. We show for the first time, that corticosterone treatment during adulthood decreases cell 106  survival and recruitment in the songbird HVC. Elsewhere in the avian brain, DHEA increased BrdU+ cells only in the absence of corticosterone suggesting that corticosterone can interfere with the action of DHEA. These data are in contrast to previous claims that DHEA inhibits glucocorticoid action and provide evidence that glucocorticoids may moderate the action of DHEA.  107  TABLES  Table 5.1 Effects of steroid treatment on NeuN measurements and BrdU+ cell number 2-factor ANOVA Tel Volume Corticosterone DHEA Corticosterone × DHEA Tel BrdU+ Cell Number Corticosterone DHEA Corticosterone × DHEA HVC Volume Corticosterone DHEA Corticosterone × DHEA HVC Neuron Number Corticosterone DHEA Corticosterone × DHEA HVC BrdU+ Cell Number Corticosterone DHEA Corticosterone × DHEA Hp Volume Corticosterone DHEA Corticosterone × DHEA Hp BrdU+ Cell Number Corticosterone DHEA Corticosterone × DHEA Ventricle BrdU+ Cell Number Corticosterone DHEA Corticosterone × DHEA  df  F-ratio  p  1,30 1,30 1,30  1.16 0.003 0.06  0.29 0.96 0.82  1,26 1,26 1,26  0.72 4.36 6.50  0.41 0.05 0.02  1,31 1,31 1,31  9.41 14.85 0.24  0.005 0.0006 0.63  1,31 1,31 1,31  6.14 15.69 1.32  0.02 0.0005 0.26  1,25 1,25 1,25  5.80 13.00 2.49  0.02 0.002 0.13  1,28 1,28 1,28  1.07 0.32 0.06  0.31 0.58 0.81  1,26 1,26 1,26  3.29 5.90 4.61  0.08 0.02 0.04  1,26 1,26 1,26  7.99 1.42 18.05  0.01 0.25 0.0003  Note: Tel: Telencephalon, Hp: Hippocampus  108  Table 5.2 Effects of steroid treatments on plasma corticosterone levels (ng/ml) (Experiment 1) Treatment Group Control Corticosterone DHEA Corticosterone + DHEA  Day -2 Baseline 6.4 ± 1.5 6.3 ± 0.9 7.9 ± 2.3 7.2 ± 1.5  Day 8 Baseline 6.5 ± 1.7 11.2 ± 6.7 9.3 ± 1.5 7.1 ± 1.6  Sampling Day Day 8 Day 22 Stressed Baseline 58.1 ± 28.3 6.7 ± 1.8 18.7 ± 1.6 6.7 ± 1.0 36.7 ± 4.7 6.7 ± 0.6 30.4 ± 4.5 7.6 ± 1.9  Day 22 Stressed 74.0 ± 24.9 23.3 ± 5.0 33.7 ± 4.0 27.8 ± 4.6  Day 29 Baseline 6.8 ± 1.5 4.5 ± 0.8 5.6 ± 0.9 8.7 ± 2.3  109  Table 5.3 Effects of corticosterone treatment on plasma corticosterone levels (ng/ml) (Experiment 2) Sampling Day Day -3  Day 2  Day 3  Day 4  Day 5  Day 8  Day 22  25.3 ±11.3  Day 8 (Change w/ stress) 171.1 ±38.5  Control (n = 3)  11.6 ±3.3  10.7 ±3.6  53.4 ±14.4  134.8 ±10.6  192.1 ±14.6  CORT (n = 5)  7.3 ±1.3  101.4 ±20.6  152.5 ±16.9  175.1 ±14.2  191.6 ±8.5  31.3 ±4.8  -8.9 ±4.6  Treatment Group  Day 29  18.5 ±8.4  Day 22 (Change w/ stress) 23.4 ±6.1  39.1 ±7.5  -10.0 ±10.4  51.0 ±38.7  8.2 ±2.0  t  0.89  3.57  4.23  2.38  0.03  1.51  3.14  1.89  2.59  1.01  p  0.47  0.04*  0.01*  0.06  0.98  0.20  0. 04*  0.17  0.05*  0.37  Note: asterisks indicate a significant difference between groups.  110  Table 5.4 Effects of steroid treatments on plasma DHEA levels (ng/ml) (Experiment 1)  Treatment Group Control Corticosterone DHEA Corticosterone +DHEA  Day -2 Baseline 3.4 ± 1.3 2.2 ± 1.1 2.1 ± 1.3 3.8 + 1.2  Day 8 Baseline 1.4 ± 0.5 1.4 ± 0.3 8.7 ± 1.8 12.9 ± 1.9  Sampling Day Day 8 Day 22 Stressed Baseline 1.0 ± 0.5 2.6 ± 1.3 0.7 ± 0.3 1.1 ± 0.2 12.6 ± 1.8 10.2 ± 1.9 13.5 ± 2.5 15.4 ± 3.7  Day 8 Stressed 0.8 ± 0.3 0.7 ± 0.1 11.0 ± 2.5 20.6 ± 5.5  Day 29 Baseline 1.0 ± 0.3 2.6 ± 1.8 7.6 ± 1.5 10.3 ± 3.1  111  FIGURES  4  5  8  sa cr i fic  30 mi nb lee d  22  29  TREATMENT DAY -3  1 2 3 4 5  8  sa cr i fic ble ed &  30 mi nb lee d  Ba se lin e  PROCEDURE  Ba se lin e&  B  Ba se lin eb Im lee pla d Ba nta se tio Ba line n se b Ba line leed se ble Ba line ed se ble lin e b ed lee d Ba se lin e& 30 mi nb lee d  e  1  Ba se lin eb lee d&  TREATMENT DAY -2  Ba se lin e&  PROCEDURE  Ba se lin e Im pla blee d nta tio n  A  Br dU Br Inje dU cti Inj ons ec tio 3x/d ns a Ba 3x y se /da lin y e& 30 mi nb lee d  e  Figure 5.1 Timeline for experiments 1 and 2  22  29  Figure 5.1 Timelines for experiment 1 (A) and experiment 2 (B). Experiment 1 and 2 were similar, however BrdU was injected 3× per day on days 4 and 5 in experiment 1. In experiment 2, no BrdU was injected, and baseline blood samples were collected on days 2-5. After day 5, the timelines for experiments 1 and 2 were identical.  112  Figure 5.2 Representative immunocytochemical staining for BrdU and NeuN  Figure 5.2 Representative immunocytochemical staining for BrdU and NeuN. A) BrdU+ cells in the lateral ventricular zone; dorsal is up and medial is to the left (scale bar 25 µm), B) BrdU cells along the lateral ventricular zone (scale bar 25 µm), C) NeuN+ cells in HVC (scale bar 250 µm); the dorsal edge of HVC is defined by the lateral ventricle and the ventral edge is indicated by arrows. Dorsal is up and medial is to the left and D) NeuN+ cells in HVC (scale bar 25 µm).  113  Figure 5.3 Effects of corticosterone and DHEA treatment on HVC A 1.4  HVC Volume (mm3)  1.2 1.0 0.8 0.6 0.4 0.2 0.0  8  8  8  8  Control  CORT  DHEA  Cort + DHEA  8  8  8  8  Control  CORT  DHEA  Cort + DHEA  6  7  7  6  Control  CORT  DHEA  Cort + DHEA  B  Neurons in HVC (x1000)  300 250 200 150 100 50 0  C 120  BrdU+ cells in HVC  100 80 60 40 20 0  Figure 5.3 Effects of corticosterone (Cort) and DHEA on HVC volume (A), HVC neuron number (B) and HVC BrdU+ cell number (C). Numbers in parentheses indicate sample sizes, and bars that share a letter are not significantly different. Note that in B, the decrease in NeuN+ cells with corticosterone treatment alone was nearly significant (p = 0.06).  114  Figure 5.4 Effects of corticosterone and DHEA treatment on the telencephalon  A  B 20  *  500 BrdU+ Cells in 3 Telencephalon (x10 )  Telencephalon Volume (mm3 )  600  400 300 200  15  10  5  100 0  (7)  (7)  (8)  (9)  Control  Cort  DHEA  Cort + DHEA  (7)  (7)  (7)  (6)  Control  Cort  DHEA  Cort + DHEA  0  Figure 5.4 The effects of corticosterone (Cort) and DHEA on telencephalon volume (A) and BrdU+ cell number in the telencephalon (excluding HVC, hippocampus and ventricular zone) (B). Numbers in parentheses indicate sample sizes, and the asterisk indicates a significant difference from all other treatment groups.  115  Figure 5.5 Effects of corticosterone and DHEA treatment on the hippocampus  A  B 2500 BrdU+ Cells in Hippocampus  Hippocampus Volume (mm3 )  20  15  10  5  0  (7)  (6)  (8)  (8)  Control  Cort  DHEA  Cort + DHEA  *  2000  1500  1000  500  (7)  (7)  (7)  (6)  Control  Cort  DHEA  Cort + DHEA  0  Figure 5.5 Effects of corticosterone (Cort) and DHEA on hippocampus volume (A) and hippocampal BrdU+ cell number (B). Numbers in parentheses indicate sample sizes, and the asterisk indicates a significant difference from all other treatment groups.  116  Figure 5.6 Effects of corticosterone and DHEA treatment on BrdU+ cells in the ventricular zone  BrdU+ cells along ventricle  4000  *  3000  2000  1000  (7)  (7)  (7)  (6)  Control  Cort  DHEA  Cort + DHEA  0  Figure 5.6 Effects of corticosterone (Cort) and DHEA on BrdU+ cell number along the lateral ventricle. Numbers in parentheses indicate sample sizes, and the asterisk indicates a significant difference from all other treatment groups.  117  REFERENCES Alvarez-Buylla, A., Kirn, J.R., 1997. Birth, migration, incorporation, and death of vocal control neurons in adult songbirds. J Neurobiol 33, 585-601. Alvarez-Buylla, A., Garcia-Verdugo, J.M., 2002. Neurogenesis in adult subventricular zone. J Neurosci 22, 629-634. Angelier, F., Clément-Chastel, C., Gabrielsen, G.W., Chastel, O., 2007. 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Territorial behaviour and hormones of pied flycatchers in optimal and suboptimal habitats. Anim Behav 56, 811-818. Smith, G.T., Brenowitz, E.A., Beecher, M.D., Wingfield, J.C., 1997. Seasonal Changes in Testosterone, Neural Attributes of Song Control Nuclei, and Song Structure in Wild Songbirds. J Neurosci 17, 6001-6010. Soma, K.K., Tramontin, A.D., Featherstone, J., Brenowitz, E.A., 2004. Estrogen contributes to seasonal plasticity of the adult avian song control system. J Neurobiol 58, 413-422. Soma, K.K., Wingfield, J.C., 1999. Endocrinology of aggression in the non-breeding season. In: Adams, N., Slotow, R. (Eds), Proceedings of the 22nd International Ornithological Congress, Durban. Birdlife of South Africa, Johannesburg, pp 1606-1620. Soma, K.K., Alday, N.A., Hau, M., Schlinger, B.A., 2004. Dehydroepiandrosterone metabolism by 3β-hydroxysteroid dehydrogenase/Δ5-Δ4 isomerase in adult Zebra Finch brain: Sex difference and rapid effect of stress. Endocrinology 145, 1668-1677. Soma, K.K., Wingfield, J.C., 2001. Dehydroepiandrosterone in songbird plasma: Seasonal regulation and relationship to territorial aggression. Gen Comp Endocrinol 123, 144-155. Soma, K.K., Wissman, A.M., Brenowitz, E.A., Wingfield, J.C., 2002. Dehydroepiandrosterone (DHEA) increases territorial song and the size of an associated brain region in a male songbird. Horm Behav 41, 203-212. Spencer, K.A., Buchanan, K.L., Leitner, S., Goldsmith, A.R., Catchpole, C.K., 2005. Parasites affect song complexity and neural development in a songbird. Proc Roy Soc B 272, 2037-2043. Torres, J.M., Ortega, E., 2003. DHEA, PREG and their sulphate derivatives on plasma and brain after CRH and ACTH administration. Neurochem Res 28, 1187-1191. Tramontin, A.D., Wingfield, J.C., Brenowitz, E.A., 2003. Androgens and estrogens induce seasonal-like growth of song nuclei in the adult songbird brain. J Neurobiol 57, 130-140. Tramontin, A.D., Hartman, V.N., Brenowitz, E.A., 2000. Breeding Conditions Induce Rapid and Sequential Growth in Adult Avian Song Control Circuits: A Model of Seasonal Plasticity in the Brain J Neurosci 20, 854-861. Widstrom, R.L., Dillon, J.S., 2004. Is There a Receptor for Dehydroepiandrosterone or Dehydroepiandrosterone Sulfate? Semin Reprod Med 289-298. Wong, E.Y.H., Herbert, J., 2004. The corticoid environment: a determining factor for neural progenitors' survival in the adult hippocampus. Eur J Neurosci 20, 2491-2498. Zhang, L., Li, B.s., Ma, W., Barker, J.L., Chang, Y.H., Zhao, W., Rubinow, D.R., 2002. Dehydroepiandrosterone (DHEA) and its sulfated derivative (DHEAS) regulate apoptosis during neurogenesis by triggering the Akt signaling pathway in opposing ways. Mol Brain Res 98, 58-66. 122  6  GENERAL DISCUSSION  SUMMARY In this thesis, I used a multidisciplinary approach to compare the effects of season and acute stress on corticosterone and DHEA levels in the brain and periphery and to examine the hypothesis that DHEA mitigates the effects of elevated glucocorticoids in the brain. These studies were conducted on wild, free-living, adult songbirds. Previous studies evaluating this hypothesis have used laboratory rodents (Kimoinides et al., 1999; Karishma and Herbert, 2002), which have very low levels of circulating DHEA and thus how these results apply to other species, including humans, is not clear. The studies described in this thesis are the first to examine the relationship between endogenous corticosterone and DHEA and among the first to comprehensively compare steroid concentrations in systemic circulation and brain. Further, I used wild subjects in their natural habitat which takes into account natural variation between individuals and seasons and facilitates an understanding of the neuroendocrinology of free living organisms (Fusani et al., 2005). Last, I present the first study to measure the effect of exogenous corticosterone on HVC in adult songbirds. In Chapter 2, I optimized a method to measure steroid levels in songbird plasma and brain and, in Chapters 3 and 4, I measured the effects of acute stress and season on systemic and local levels of corticosterone and DHEA. In chapter 3, I discovered that acute restraint stress affects corticosterone and DHEA levels in jugular plasma differently than in brachial plasma and that this effect of restraint was season-dependent. These results led me to examine the effects season and acute stress on corticosterone and DHEA levels directly in brain tissue (Chapter 4) where I found that the effects of acute stress were season and brain region specific. For corticosterone, acute stress increased jugular levels during all seasons but decreased corticosterone levels in the hippocampus during molt. For DHEA, acute stress increased DHEA 123  levels in jugular plasma during molt but did not affect brain DHEA levels in any season. Also, DHEA levels were up to 5× higher in brain than in jugular plasma and were highest in the hippocampus. Lastly, in chapter 5, I experimentally manipulated levels of corticosterone and DHEA, in vivo, to determine if 1) corticosterone was detrimental to cell survival or recruitment in the adult songbird brain and 2) if DHEA abolished the effects of corticosterone.  LIMITATIONS Chapters 3 and 4 examined wild song sparrows in their natural environment, and as Fernando Nottebohm said, “Unless you understand the needs, the habits, the problems of an animal in nature, you will not understand it at all...Take nature away and all your insight is in a biological vacuum” (Specter, 2001). However, unlike laboratory studies, field studies lack control of environmental variables. For example, our ability to detect seasonal differences in neurosteroids may have been reduced as there was sometimes large individual variation within a season. In chapter 3, individual variation in plasma DHEA and corticosterone concentrations was greatest during the breeding season. Within the breeding season, steroid levels change according to the stage of breeding (e.g., nest building, incubation, feeding young) (Wingfield and Hahn, 1994) but I was not able to control for this because it would have taken a significant amount of time to find and monitor nests. In the future, studies that focus on monitoring a wild population and collecting biological samples from individuals at consistent sub-stages would be a valuable contribution that could account for individual and within-season variation. Our work also focuses only on males. Wild female song sparrows are difficult to capture and exposing females to stress during the breeding season could have a severe impact on reproductive success because females invest significantly more in parental care than males. The conclusions drawn from the present data cannot be easily applied to females as there are marked sex differences in steroid concentrations in songbirds (e.g. Chin et al., 2008), in the enzymes 124  that regulate neurosteroid synthesis (Soma et al., 2004; Pradhan et al., 2008), and in neuroanatomy (Ball and Shackleton, 2001). Furthermore, the effects of stress may differ between males and females. For example, in the rat dentate gyrus, acute stress decreases cell proliferation in males, but not in females (Falconer and Galea, 2003). In Chapter 5, I found that corticosterone treatment affected neuroanatomy. However, administering corticosterone using silastic implants was challenging. A hole must be poked through the tubing or one end left unsealed to facilitate corticosterone release (Astheimer et al., 2000; Breuner and Hahn, 2003; Martin et al., 2005), which can result in the release of a large bolus followed by a tapering dose of hormone (Gray et al., 1990). Because of this, dosages can be difficult to control. I used silastic implants with a single 22 gauge needle hole, which was effective in increasing plasma corticosterone over the short-term, but inter-individual variation in plasma corticosterone levels were high. Osmotic minipumps have been used to successfully administer corticosterone in songbirds (Horton et al., 2007); however, this study only used the pumps for 7 days. Furthermore, the pumps are relatively large for small songbirds and, in a pilot study on song sparrows, I found that when implanted subcutaneously, the skin tore by day 7 and only 1 of the 4 pumps that were administered remained implanted after the 4 week period (the advertised release duration of these pumps was 4 weeks). Recently, French et al. (2007) reliably increased plasma corticosterone levels in lizards using an injectable gel that polymerizes under the skin, forming a slow release pellet. This technique appears to be promising and will soon be tested in songbirds. Daily corticosterone injections are not ideal for songbirds as repeated handling and needle punctures can increase endogenous corticosterone levels as I demonstrated in Chapter 5. The method of bromodeoxyuridine (BrdU) administration in Chapter 5 should also be refined to avoid repeated handling and injections. There is some debate about the optimal BrdU dose and administration schedule. BrdU is toxic and can decrease DNA stability and increase 125  the risk of mutations (Taupin, 2007). Doses of 30 to 150mg/kg have been used in adult and developing birds (Cao et al., 2002; Hilton et al., 2002; Hoshooley et al., 2007), and doses of 50 to 300mg/kg in adult rats do not seem to have physiological side effects (Cameron and McKay, 2001) . I chose multiple injections with a low BrdU dose (65mg/kg) in an attempt to increase the number of cells labelled over multiple mitotic cycles. However, increases in endogenous corticosterone levels after multiple injections could have thwarted these attempts, and one BrdU injection of a larger dose could be more effective (Cameron and McKay, 2001; Galea et al., 2008). Further experiments are required to optimize the BrdU dose and injection protocol in songbirds.  FUTURE DIRECTIONS There is accumulating evidence for local steroid synthesis in the brain and other organs (Schmidt et al., 2008). Thus, steroid concentrations in the brain and other target tissues (e.g. immune organs) can be dramatically different from those in plasma (Newman et al., 2008a; Schmidt and Soma, 2008; Newman and Soma in review). Further, it is possible that locally synthesized steroids are either rapidly metabolized or sulphated. For example, levels of pregnenolone sulphate, but not pregnenolone, change seasonally in the amphibian brain (Takase et al., 1999) and sulfotransferase expression is sensitive to glucocorticoids (Runge-Morris, 1999). The findings in this thesis can be extended by investigating the seasonal and stressinduced dynamics of DHEA metabolites such as estradiol and DHEA-sulphate. Currently, further optimizations are underway to refine the solid phase extraction method in order to separate free steroids from their sulphated derivatives and we are measuring the effects of season and acute stress on 3β-hydroxysteroid dehydrogenase, the enzyme which catalyzes the conversion of DHEA to androstenedione, in the song sparrow brain.  126  In Chapters 3 and 4, I found differential effects of season and acute restraint stress on steroid levels in brain and plasma. However, different types of stressors may have different effects on neurosteroid concentrations. For example, I will test the effects of chronic stress on corticosterone, DHEA and DHEA metabolites in plasma and brain. I will use an established system in which adult male song sparrows living under higher predator exposure have elevated baseline and stress-induced plasma corticosterone levels when compared with their low predator exposure counterparts (Clinchy et al., 2004). Secondly, I will also test the effect of social stress on corticosterone, DHEA and DHEA metabolites levels in plasma and the brain. Adult male song sparrows will be exposed to simulated territorial intrusion (STI), a social disruption that is known to affect systemic steroids in other songbirds (Landys et al., 2007). Both chronic and social stress may have effect season-specific effects, thus these two studies will be conducted across different seasons. In Chapters 4 and 5, I found evidence that acute stress or exogenous glucocorticoids may affect the 11β-hydroxysteroid dehydrogenase isozymes, which regulate the interconversion between active and inactive glucocorticoids. In Chapter 4, acute stress decreased corticosterone concentrations in the hippocampus during molt and it is possible that upregulation of 11β-HSD type 2, which rapidly inactivates corticosterone to dehydrocorticosterone (Holmes and Seckl, 2006; Klusonova et al., 2008), is a mechanism to protect the hippocampus from glucocorticoid exposure. Interestingly, acute stress rapidly upregulates 11β-HSD2 activity in the rat placenta to protect the fetus from high levels of maternal glucocorticoids (Welberg et al., 2005). Furthermore, DHEA is also known to upregulate 11β-HSD type 2 activity (Balazs, 2008) and decrease expression of 11β-HSD type 1 (Apostolova, 2005) and thus DHEA could also act to decrease local levels of glucocorticoids. In Chapter 5, after 4 days of exogenous corticosterone treatment, corticosterone levels in plasma were no longer elevated, perhaps due to increased 11β-HSD2 activity in peripheral tissues. These results raise the interesting possibility that 127  corticosterone and DHEA both modulate the expression and activity of 11β-HSD and future studies will examine the effects of exogenous steroids, season and stress on these isozymes in the brain. Last, the data described in chapter 5 indicate that corticosterone and DHEA have dramatic and opposite effects on HVC neuroanatomy in nonbreeding song sparrows. It will be important to determine if these effects have long-term consequences for behaviour and fitness. For example, do changes in HVC size and cell recruitment during the nonbreeding season affect singing behaviour? If so, does this variation in singing behaviour influence mate choice and reproductive success? Furthermore, would exposure to corticosterone and DHEA during the breeding season, when rates of neuronal turnover are lower than in the nonbreeding season (Wilbrecht and Kirn, 2004), have the similar effects on HVC? These questions pose interesting avenues for future research in the field and laboratory.  CONCLUSIONS I have found distinct effects of both acute stress and season on plasma and brain levels of corticosterone and DHEA. Our data support the hypothesis that DHEA is synthesized in the songbird brain, particularly in the hippocampus. Furthermore, I demonstrate that corticosterone and DHEA can have opposite, additive effects on cell proliferation and survival in the brain; and that in some cases, corticosterone inhibits the actions of DHEA on the survival and recruitment of new cells in the brain. Last, I have established a new model for studying the relationship between glucocorticoids and DHEA in the brain, and several psychiatric disorders, such as major depression, are associated with altered glucocorticoid and DHEA profiles. Thus, the use of this model may aid in understanding the role that glucocorticoids and DHEA play in the etiology and treatment of some psychiatric illnesses.  128  REFERENCES Apostolova, G., Schweizer, R.A.S., Balazs, Z., Kostadinova, R.M., Odermatt, A., 2005. Dehydroepiandrosterone inhibits the amplification of glucocorticoid action in adipose tissue. Am J Physiol Endocrinol Metab 288, 957-964. Astheimer, L.B., Buttemer, W.A., Wingfield, J.C., 2000. Corticosterone Treatment Has No Effect on Reproductive Hormones or Aggressive Behavior in Free-living Male Tree Sparrows, Spizella arborea. Horm and Behav 37, 31-39. Balazs, Z., Schweizer, R.A.S., Frey, F.J., Rohner-Jeanrenaud, F., Odermatt, A., 2008. DHEA Induces 11β-HSD2 by Acting on CCAAT/enhancer-Binding Proteins. J Am Soc Nephrol 19, 92-101. Ball, G.F., Macdougall-Shackleton, S.A., 2001. Sex differences in songbirds 25 years later: What have we learned and where do we go? Microsc Res Tech 54, 327-334. Breuner, C.W., Hahn, T.P., 2003/1. Integrating stress physiology, environmental change, and behavior in free-living sparrows. Horm and Behav 43, 115-123. Cameron, H.A., McKay, R.D.G., 2001. Adult neurogenesis produces a large pool of new granule cells in the dentate gyrus. J Comp Neurol 435, 406-417. Cao, J., Wenberg, K., Cheng, M., 2002. Lesion induced new neuron incorporation in the adult hypothalamus of the avian brain. Brain Res 943, 80-92. Clinchy, M., Zanette, L., Boonstra, R., Wingfield, J.C., Smith, J.N., 2004. Balancing food and predator pressure induces chronic stress in songbirds. Proc Biol Sci 271:2473-2479 Chin, E.H., Shah, A.H., Schmidt, K.L., Sheldon, L.D., Love, O.P., Soma, K.K., 2008. Sex differences in DHEA and estradiol during development in a wild songbird: Jugular versus brachial plasma. Horm Behav 54, 194-202. Falconer, E.M., Galea, L.A.M., 2003. 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Corticosterone, foraging behavior, and metabolism in dark-eyed juncos, Junco hyemalis. Gen Comp Endocrinol 79, 375-384. Hilton, L.S., Bean, A.G.D., Kimpton, W.G., Lowenthal, J.W., 2002. Interleukin-2 Directly Induces Activation and Proliferation of Chicken T Cells In Vivo. Journal of Interferon & Cytokine Res 22, 755-763. Holmes, M.C., Seckl, J.R., 2006. The role of 11β-hydroxysteroid dehydrogenases in the brain. Mol Cell Endrocrinol 248, 9-14. Horton, B.M., Long, J.A., Holberton, R.L., Intraperitoneal delivery of exogenous corticosterone via osmotic pump in a passerine bird. Gen Comp Endocrinol 152, 8-13. Hoshooley, J.S., Phillmore, L.S., Sherry, D.F., MacDougall-Shackleton, S.A., 2007. Annual cycle of the Black-Capped Chickadee: Seasonality of food-storing and the hippocampus. Brain Behav Evol 69, 161-168. Karishma, K.K., Herbert, J., 2002. Dehydroepiandrosterone (DHEA) stimulates neurogenesis in the hippocampus of the rat, promotes survival of newly formed neurons and prevents corticosterone-induced suppression. Eur J Neurosci 16, 445-453. Kimonides, V.G., Spillantini, M.G., Sofroniew, M.V., Fawcett, J.W., Herbert, J., 1999. Dehydroepiandrosterone antagonizes the neurotoxic effects of corticosterone and translocation of stress-activated protein kinase 3 in hippocampal primary cultures. Neuroscience 89, 429-436. Klusoňová, P., Kučka, M., Mikšík, I., Bryndová, J., Pácha, J., 2008. Chicken 11βhydroxysteroid dehydrogenase type 2: Partial cloning and tissue distribution. Steroids 73, 348355. Landys, M.M., Goymann, W., Raess, M., Slagsvold, T., 2007. Hormonal responses to malemale social challenge in the blue tit (Cyanistes caeruleus): Single-broodedness as an explanatory variable. Phsyiol Biochem Zool 80, 228-240. Martin II, L.B., Gilliam, J., Han, P., Lee, K., Wikelski, M., 2005. Corticosterone suppresses cutaneous immune function in temperate but not tropical House Sparrows, Passer domesticus. General and Comparative Endocrinology 140, 126-135. Pradhan, D.S., Yu, Y., Soma, K.K., 2008. Rapid estrogen regulation of DHEA metabolism in the male and female songbird brain. J Neurochem 104, 244-253. Runge-Morris, M., Wu, W., Kocarek, T.A., 1999. Regulation of Rat Hepatic Hydroxysteroid Sulfotransferase (SULT2-40/41) Gene Expression by Glucocorticoids: Evidence for a Dual Mechanism of Transcriptional Control. Mol Pharmacol 56, 1198-1206.  130  Schmidt, K.L., Pradhan, D.S., Shah, A.H., Charlier, T.D., Chin, E.H., Soma, K.K., 2008. Neurosteroids, immunosteroids, and the Balkanization of endocrinology. Gen Comp Endocrinol 157, 266-274. Schmidt, K.L., Soma, K.K., 2008. Cortisol and Corticosterone in the Songbird Immune and Nervous Systems: Local versus Systemic Levels during Development. Am J Physiol Regul Integr Comp Physiol 295, 103-110. Soma, K.K., Alday, N.A., Hau, M., Schlinger, B.A., 2004. Dehydroepiandrosterone metabolism by 3β-hydroxysteroid dehydrogenase/Δ5-Δ4 isomerase in adult Zebra Finch brain: Sex difference and rapid effect of stress. Endocrinology 145, 1668-1677. Specter, M., 2001. Rethinking the brain: How the songs of canaries upset a fundamental principle of science. The New Yorker July 23, 42-53. Takase, M., Ukena, K., Yamazaki, T., Kominami, S., Tsutsui, K., 1999. Pregnenolone, pregnenolone sulfate, and cytochrome P450 side-chain cleavage enzyme in the amphibian brain and their seasonal changes. Endocrinology 140, 1936-1944. Taupin, P., 2007. BrdU immunohistochemistry for studying adult neurogenesis: Paradigms, pitfalls, limitations, and validation. Brain Res Rev 53, 198-214. Welberg, L. A. M., Thrivikraman, K.V., Plotsky, P.M., 2005. Chronic maternal stress inhibits the capacity to up-regulate placental 11β-hydroxysteroid dehydrogenase type 2 activity. J Endocrinol 186, 7-12. Wilbrecht, L., Kirn, J.R., 2004. Neuron addition and loss in the song system: Regulation and function. Ann NY Acad Sci 1016, 659-683. Wingfield, J.C., Hahn, T.P., 1994. Testosterone and territorial behaviour in sedentary and migratory sparrows. Anim Behav 47, 77-89.  131  

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