<<

Neural Action of Androgens in the Brain Clock

By

Lindsay Coome

A thesis submitted in conformity with the requirements for the degree of Master of Arts Graduate Department of Psychology University of Toronto

© Copyright by Lindsay Coome, 2013 Neural Action of Androgens in the Suprachiasmatic Nucleus Brain Clock

Lindsay Coome

Master of Arts

Graduate Department of Psychology University of Toronto 2013

Abstract

The suprachiasmatic nucleus (SCN) of the hypothalamus is the of a master circadian clock that is critical in the temporal organization of circadian activity. The SCN coordinates the rhythmic secretion of gonadal hormones, and in turn, reproductive hormones may act on their receptors within the SCN to alter circadian function. Using transgenic mice that over-express (AR) only in neurons, the current study investigated the influence of neural AR on the function of the SCN. In particular, it addressed the effects of androgens on circadian behaviours as well as physiological responses to light within the SCN by measuring

Fos response after a phase-shifting light pulse. It was found that transgenic mice demonstrate a smaller increase in Fos expression in response to a light pulse than do wildtypes. Interpretations of our findings, including the possible functional significance of AR within the SCN, are discussed.

ii Acknowledgements I would first like to thank my supervisor, Dr. Ashley Monks, for his wisdom, guidance, and tremendous patience throughout the . His support and encouragement have allowed me to grow as a scientist and as a person. I would also like to thank my subsidiary advisor Dr. Joel Levine, for allowing me to benefit from his and expertise, and Dr. Robert Gerlai, for joining my thesis committee and showing genuine interest in my work. I would like to thank all the members of the Monks and Holmes labs, particularly Ashlyn Swift-Gallant, for her friendship and unwavering belief in my abilities. Finally, I’d like to thank my parents for a lifetime of love and encouragement, without which none of this would be possible.

iii Table of Contents

Abstract ...... ii

Acknowledgements ...... iii

List of Tables ...... vi

List of Figures ...... vii

Introduction ...... 1 1.1 Overview of Circadian Rhythms ...... 1 1.2 The Circadian System ...... 1 1.2.1 The SCN as the Master Clock ...... 1 1.2.2 The Molecular Basis of the Clock ...... 3 1.2.3 Organization of the SCN ...... 3 1.2.4 Diffusible and Neural Signals ...... 4 1.3 Circadian Rhythms and Hormones ...... 6 1.3.1 Circadian Regulation of Neuroendocrine Secretions ...... 6 1.3.2 Endocrine Influences on the Circadian System ...... 6 1.3.3 The Influence of Ovarian Hormones on Circadian Behaviour ...... 6 1.3.4 The Influence of Androgens on Circadian Behaviour ...... 7 1.4 The Role of AR in the SCN ...... 10 1.5 The Study ...... 12

Methods ...... 13 2.1 Animals and Housing ...... 13 2.2 Experimental Design ...... 14 2.3 Gonadectomy (GDX) and Hormone Replacement ...... 14 2.3.1 Steroid Implants ...... 14 2.3.2 Gonadectomy ...... 15 2.4 Behavioural Testing ...... 15 2.5 Neural Activation Following Exposure to Light ...... 15 2.6 c-Fos Immunohistochemistry ...... 16

iv 2.7 Androgen Receptor Immunohistochemistry ...... 16 2.8 Data Analysis ...... 17 2.8.1 Brain Analyses ...... 17 2.8.2 Behavioural Analyses ...... 17 2.8.3 Statistical Analyses ...... 18

Results ...... 18 3.1 Somatic Measures ...... 18 3.2 Behaviour ...... 19 3.3 Neural Activation ...... 19

Discussion...... 20

References ...... 26

Tables ...... 33

Figure Captions ...... 37

Figures ...... 38

v List of Tables Table 1. Number of animals per condition for behavioural studies. Table 2. Number of animals per condition for analysis of neural activation. Table 3. Group Means and Standard Error of the Means (SEM) for number of Fos-ir cells. Table 4. Group Means and Standard Error of the Means (SEM) for number of beam breaks.

vi List of Figures Figure 1. Total number of beam breaks recorded over 5 days. Figure 2. Number of Fos-ir cells in bilateral SCN. Figure 3. Androgen receptor immunoreactive cells of Nestin-AR mutants and wildtype littermates.

vii 1

Introduction 1.1 Overview of Circadian Rhythms To ensure optimal function, animals have evolved to restrict many behaviours, including locomotor activity, feeding, and reproductive behaviours, to specific temporal niches. Much of this temporal variation in behaviour is controlled by biological clocks. Although many of the observed rhythms in physiology and behaviour are mediated by internal timing mechanisms, other aspects of rhythmicity are sensitive to environmental time cues, such as the daily light-dark . Daily rhythms that are driven by external signals in the environment are called diurnal rhythms, and disappear under constant environmental conditions. Rhythms that are endogenously driven are referred to as circadian rhythms, and persist in the absence of external cues. Under constant conditions devoid of any external time cues, it has been found that circadian rhythms continue to be expressed with a that approximates, but is rarely identical to, 24 (Pittendrigh, 1960). Some animals have free-running internal clocks with a period longer than 24 hours, while others, such as mice, free-run with a period shorter than 24 hours. The fact that the circadian period under free-running conditions is not exactly 24 hours suggests that the circadian pacemaker can be synchronized to external time cues (Arble, Copinschi, Vitaterna, Van Cauter, and Turek, 2011). Light is the primary environmental time cue, or “zeitgeber”, that synchronizes biological rhythms, however other zeitgebers include temperature, food availability, noise, and social cues (Bruce, 1960). Internal clocks must be adjusted on a daily basis to stay synchronized to local time, and this synchronization of the internal clock to a periodic cue in the environment is known as entrainment. In the absence of zeitgeber cues, circadian rhythms are said to be free- running.

1.2 The Circadian System 1.2.1 The SCN as the Master Clock The circadian system is necessary for the appropriate timing of physiological and behavioural processes, and this temporal organization is controlled by a master circadian clock located in the suprachiasmatic nuclei (SCN) of the hypothalamus. In mammals, photic information is transmitted from the retina to the SCN via the retinohypothalamic tract (RHT), 2 which is then relayed by the SCN to target sites in the brain (Moore and Lenn, 1972). The identification of the SCN as the master circadian pacemaker first began with the discovery that lesions of this nuclei produced arrhythmicity, suggesting that the SCN is necessary for circadian function (Moore and Eichler, 1972; Stephan and Zucker, 1972). Stephan and Zucker’s (1972) finding that bilateral lesions of the SCN permanently eliminated circadian rhythms in locomotor activity and drinking behaviour in rats provided particularly strong evidence for the SCN as the site of the circadian clock. It was then demonstrated that the SCN, when isolated in an island of hypothalamic tissue, maintained circadian rhythmicity in electrical activity, while rhythmicity in neural activity outside of the SCN was lost following the isolation surgery (Inouye and Kawamura, 1979). Such isolation studies suggested that the SCN imparts circadian rhythmicity on other brain structures through its neural connections (Weaver, 1998), though more recently it has been determined that diffusible signals from the SCN play a role in sustaining circadian rhythms in behaviour (Silver, LeSauter, Tresco, and Lehman, 1996). Further evidence that the SCN is an autonomous circadian oscillator came from studies of SCN tissue transplantation and examination of rhythmicity within SCN tissue slices maintained in vitro (Weaver, 1998). Several studies of the SCN in vitro demonstrated that the firing rates of neurons in brain slices of the SCN maintained a similar to that of intact animals, with high levels of activity observed during the subjective and low levels during the subjective night, and were in accordance with the light-dark cycle of the donor animal (Groos and Hendriks, 1982; Green and Gillette, 1982). In vitro studies of the SCN allowed researchers to determine which properties of the SCN were truly endogenous apart from the complex in vivo environment, such as afferents from other brain regions (Gillette, 1991). Studies involving transplantation of SCN tissue from one animal to another demonstrated that behavioural rhythmicity can be restored in rodents made arrhythmic by SCN lesions through the transplantation of fetal hypothalamic tissue containing the SCN (Lehman et al., 1987). The discovery of the tau mutant hamster subsequently made it possible to demonstrate that the circadian period of the clock can be transferred by transplantation of SCN tissue. The tau mutant hamster shows a short circadian period under conditions of constant darkness. In SCN-lesioned tau mutants and wild-type animals, reciprocal transplants of tissue from the SCN restored 3 rhythms with a period predicted by the genotype of the donor tissue, not the host (Ralph, Foster, Davis, and Menaker, 1990). Several lines of evidence, then, place the SCN at the top of the hierarchy of the circadian system.

1.2.2 The Molecular Basis of the Clock The discovery of genetic that affect clock function in Drosophilia melanogaster (Konopka and Benzer, 1971) had a tremendous impact on our understanding of the genetic basis for circadian behaviour and subsequent investigation has led to the identification of the cellular mechanisms of clock in mammals. Briefly, circadian rhythms are generated at the cellular level via an autoregulatory negative feedback loop that takes approximately 24 hours. Two proteins, CLOCK and BMAL1, bind to each other in the cytoplasm forming heterodimers (Kondratov et al., 2003). These heterodimers are able to translocate back into the nucleus and interact with the E-box components of the promoter regions of Period (Per) and (Cry) genes. The binding of the CLOCK-BMAL1 heterodimer to the E-boxes of these genes functions to drive the transcription of three Period (Per1, Per2, and Per3) as well as two Cryptochrome (Cry1 and Cry2) genes (Reppert and Weaver, 2002). The resulting proteins, PER and CRY, build up in the cytoplasm of the cell over the course of the day and when at high enough levels, form heterodimers with each other. These PER-CRY heterodimers, similar to the CLOCK-BMAL1 heterodimers, are able to translocate back into the nucleus. Once in the nucleus, the PER-CRY heterodimers interact with and inhibit the CLOCK-BMAL1 heterodimer, leading to the subsequent inhibition of Period and Cryptochrome transcription (Ko and Takahashi, 2006).

1.2.3 Organization of the SCN Although the circadian system has often been characterized as consisting of three components, input, pacemaker, and output, more recent studies point to the functional and anatomical complexity of the circadian clock. The SCN is a heterogeneous structure, and can be divided into two main anatomical subdivisions, a dorsomedial “shell” and a ventrolateral “core”. Evidence for this distinction comes from the discovery that only some SCN cells possess 4 oscillatory capacity, and not all cells receive photic input directly from the retina (Antle and Silver, 2005). It is now known that the core of the SCN receives direct innervation from the retina (Bryant, LeSauter, Silver, and Romero, 2000; Tanaka et al., 1997), and cells within the core express clock genes Per1 and Per2 in response to a light pulse (Hamada, LeSauter, Venuti, and Silver, 2001; Yan and Okamura, 2002). It has also been demonstrated that cells within the SCN core do not show rhythmic expression of Per1 and Per2 and lack rhythmic electrical activity (Hamada, Antle, and Silver, 2004; Jobst and Allen, 2002; Karatsoreos, Yan, LeSauter, and Silver, 2004). By contrast, in the SCN shell, Per1 and Per2, and Per3 oscillate detectably on a circadian basis but are not inducible by light (Hamada et al., 2001). The SCN core and shell differ in regard to peptidergic content as well. Arginine vasopressin (AVP)-containing cells comprise the shell of the SCN, whereas the core has several cell types (Antle and Silver, 2005; Abrahamson and Moore, 2001; Moore, Speh, and Leak, 2002). The core primarily contains cells producing vasoactive intestinal polypeptide (VIP) or gastrin-releasing peptide (GRP) (Moore, Speh, and Leak, 2002). In mice, the GRP-expressing cells in the core of the SCN are inducible by light (Karatsoreos, Yan, LeSauter, and Silver, 2004). It has been suggested that the core communicates photic information to the oscillators of the SCN shell, which then produce coordinated output to other regions of the hypothalamus (Ruggiero and Silver, 2010). However, SCN lesions that destroy the ventrolateral core region but largely spare the shell result in complete loss of rhythmicity, including rhythms in locomotor activity, hormone secretion, body temperature, and heart rate (LeSauter and Silver, 1999; Kriegsfeld, LeSauter, and Silver, 2004). Thus, despite having no rhythmic clock gene expression, it appears that the core is essential for producing a daily rhythmic output signal from the SCN. It is largely still unclear how the light- sensitive and rhythmic compartments of the SCN interact to communicate information to other brain regions and the rest of the body (Butler et al., 2009).

1.2.4 Diffusible and Neural Signals As previously mentioned, transplantation of SCN tissue to an SCN-lesioned host restores circadian rhythms in behaviour, including rhythms in locomotor activity, gnawing, and drinking (LeSauter and Silver, 1994; Lehman et al., 1987; Silver, Lehman, Gibson, Gladstone, and 5

Bittman, 1990). Previous work had indicated that neural efferents were necessary to sustain circadian rhythmicity (Inouye and Kawamura, 1979). However, although some neural connections may be established between the donor SCN tissue and the host brain, subsequent studies suggested that the transplants produce diffusible signals as well. SCN transplants implanted into the third ventricle of an SCN-lesioned host restore behavioural rhythms despite varied attachment sites and efferent connections to the host tissue (LeSauter and Silver, 1994; Lehman et al., 1987; Silver, LeSauter, Tresco, and Lehman, 1996). More definitively, it was found that locomotor rhythms are still restored even when donor tissue is encapsulated before transplant, preventing neural outgrowth while allowing the diffusion of humoral signals (Silver et al., 1996). One candidate for a diffusible signal is prokineticin-2 (PK2), a protein expressed rhythmically in the SCN with receptors present throughout all major SCN target areas (Cheng et al., 2002). PK2-deficient mice exhibit disrupted locomotor rhythms as well as a reduction in other physiological and behavioural rhythms (Li et al., 2006). Another possible candidate diffusible signal is transforming growth factor-alpha (TGF-α). TGF-α is also expressed rhythmically in the SCN, and has been implicated in regulating activity levels in multiple species (Kramer et al., 2001; Olofsson and de Bono, 2008). However, more research is necessary to determine the role of these diffusible signals in transmitting information from the SCN. Although a diffusible signal is sufficient to sustain behavioural rhythmicity, endocrine rhythms require neural output from the SCN. Studies using knife cuts severing SCN efferents have been shown to abolish endocrine rhythms, and endocrine rhythms are not restored by SCN transplants to lesioned animals (Hakim, DeBernardo, and Silver, 1991; Meyer-Bernstein et al., 1999; Silver et al., 1996; Nunez and Stephan, 1977). Neural output from the SCN has been well- studied using tract-tracing techniques, and projections have been found in both the core and the shell of the SCN (Kriegsfeld, Leak, Yackulic, LeSauter, and Silver, 2004). Projections have been found to target brain regions that contain neuroendocrine cells producing hypothalamic-releasing hormones (Butler, Kriegsfeld, and Silver, 2009). Additionally, the SCN projects to the pineal gland via a multisynaptic pathway (Klein, 1985; Klein et al., 1983). Thus, the SCN may exert control over neuroendocrine cell populations directly via neural projections, which then may have widespread effects throughout the body (Butler, Kriegsfeld, and Silver, 2009). 6

1.3 Circadian Rhythms and Hormones 1.3.1 Circadian Regulation of Neuroendocrine Secretions Daily rhythms have been observed in the secretion of nearly all hormones (Arble et al., 2011). The importance of circadian rhythms in endocrine processes has been known since Everett and Sawyer (1950) determined that the LH surge that occurs during a critical period during the afternoon of proestrus is necessary to induce ovulation later that night. When activity in the hypothalamus is blocked by barbiturate anesthetics in females before certain critical hours during proestrus, the LH surge is delayed by 24 hours rather than the of the anesthetic sedation (Butler et al., 2009). This work demonstrated a role for the circadian system in the daily pulse of LH release, and it was later determined that ablation of the SCN blocks the daily surge in LH (Gray, Södersten, Tallentire, and Davidson, 1978). Thus, the SCN are important for the timing of gonadotropin secretion, which in turn may affect the timing of gonadal hormone secretion and the onset of reproductive behaviours (Buijs, Van Eden, Goncharuk, and Kalsbeek, 2003; Van Der Beek, Horvath, Wiegant, Van Den Hurk, and Buijs, 1998). Successful reproduction requires the temporal coordination of many hormone-dependent physiological and behavioural processes, and the circadian system ensures the appropriate timing of these processes.

1.3.2 Endocrine Influences on the Circadian System While it is well established that the SCN controls the timing of endocrine secretions, hormones may in turn feed back to the SCN and affect circadian rhythmicity. It has been suggested that the main function of hormonal influence on circadian rhythms is to “fine-tune” the temporal release of gonadal hormones, forming a neuroendocrine feedback loop (Kriegsfeld and Silver, 2006). Hormonal influences on behavioural and physiological rhythms may provide insight into the function of the circadian clock.

1.3.3 The Influence of Ovarian Hormones on Circadian Behaviour Ovarian hormones may have pronounced effects on daily activity cycles. Cycling female hamsters show a spontaneous phase advance during estrus, called “scalloping”. This pattern of 7 earlier onset of activity occurs when estradiol levels are at their highest, and can be eliminated by ovariectomy. Further, continuous treatment of ovariectomized animals with estradiol shortens the period of activity onset compared to ovariectomized controls (Morin, Fitzgerald, and Zucker, 1977), suggesting a direct effect of estrogen on the circadian clock. Estradiol may also affect the consolidation of activity in females. Ovariectomized hamsters exposed to constant light showed splitting of the daily activity phase into two separate bouts approximately 12 hours apart (Morin, 1980). Replacement with capsules containing estradiol benzoate prevents this activity splitting in females. By contrast, castration does not produce rhythm desynchrony in males, and estradiol benzoate administration via capsule implant had no effect on the incidence of rhythm desynchronies (Morin and Cummings, 1982). Taken together, these findings suggest that estrogens facilitate the synchrony of circadian rhythms in females but not males. Evidence that estrogen may act directly on the SCN comes from the fact that α and β estrogen receptors have been found in the SCN in several mammalian species including humans, macaques, and rats (Kruijver and Swaab, 2002; Gundlah et al., 2000; Su, Qiu, Zhong, and Chen, 2001). However, distribution throughout the SCN is sparse (Shughrue, Komm, and Merchenthaler, 1996), and it is possible that estrogen exerts its effects on the circadian clock indirectly. Several studies have shown that cells in the preoptic area, amygdala, bed nucleus of the stria terminalis (BNST), and arcuate nuclei possess a large number of estrogen receptor-alpha (ERα) and project directly to the SCN, while the SCN does not provide direct input to these structures (de la Iglesia, Blaustein, and Bittman, 1999). Thus, the effects of estradiol on the circadian pacemaker may be direct or indirect.

1.3.4 The Influence of Androgens on Circadian Behaviour In males, testosterone has been shown to affect the period, precision, and consolidation of activity rhythms. Castration of mice lengthens the free-running period of locomotor activity, reduces running wheel activity at the beginning of the active period, and stimulates it at the end of the active period (Daan, Damassa, Pittendrigh, and Smith, 1975). In male hamsters, long- exposure to short day lengths leads to a decrease in testicular size, levels of plasma testosterone, and copulatory behaviour (Ellis and Turek, 1983). After castration or testicular regression in 8 hamsters, the precision of the onset of locomotor activity is lost, total amount of daily locomotor activity is reduced, and the length of the daily active period is increased (Morin and Cummings, 1981; Ellis and Turek, 1983). Testosterone replacement reverses the effects of castration. Testosterone can be aromatized to estradiol, and it has been suggested that this is how testosterone exerts its effects on circadian rhythms (Kriegsfeld and Silver, 2006). Testosterone aromatized to estradiol may then act directly within the SCN or indirectly through sites that communicate with the SCN (Shughrue, Scrimo, Lane, Askew, and Merchenthaler, 1997; de la Iglesia et al., 1999). For instance, testosterone’s effects on activity levels might occur through its conversion to estradiol. Castration significantly reduces overall levels of running-wheel activity in rats, and conversion of testosterone to estradiol may be responsible for stimulating activity levels (Roy and Wade, 1975). Roy and Wade (1975) compared aromatizable and non- aromatizable androgens with estradiol benzoate (EB) in their ability to induce wheel-running in castrated male rats. While it was found that the non-aromatizable androgen, dihydrotestosterone (DHT), had no effect on wheel-running, animals given the aromatizable testosterone (T) did show increased running levels. Similarly, EB was found to be most effective in stimulating running, and is nearly 100 more effective than T in increasing running-wheel activity in rats. More recent evidence suggests that non-aromatized testosterone may affect circadian behaviours in mice as well. Katsoreos et al. (2007) implanted capusles bearing T, DHT, or an empty capsule in castrated mice. Gonadectomized mice demonstrated lengthened period, reduced precision of activity onset, and eliminated activity bout at the onset of subjective night. Replacement with either T or DHT restored these measures to precastration levels, suggesting that these effects of testosterone are mediated by androgen receptors (ARs). Although androgens may affect circadian behaviour at multiple sites, precision and period are believed to be controlled by the SCN directly (Daan and Oklejewicz, 2003), suggesting that testosterone may act directly through ARs within the SCN (Karatsoreos and Silver, 2007). ARs have been discovered in the SCN of several species, including humans, ferrets, and rhesus monkeys (Kashon, Arbogast, and Sisk, 1998; Rees and Michael, 1982; Michael and Rees, 1982; Fernández-Guasti, Kruijver, Fodor, and Swaab, 2000). Katsoreos et al. (2007) have found that, 9 in the mouse, ARs are highly localized to the ventrolateral core of the SCN, with fibre tracts exiting dorsally from the nucleus; only sparse staining was observed in the dorsomedial shell. Additionally, the majority of AR cells within the SCN were found to coexpress GRP, whereas very few coexpress AVP or VIP. If the site of AR action is specific to the core of the SCN, this finding would be important in understanding the effects of androgens on SCN circuits (Karatsoreos et al., 2007). AR expression is sexually dimorphic in both humans and rodents (Fernández-Guasti et al, 2000; Iwahana, Karatsoreos, Shibata, and Silver, 2008), and ARs are more highly expressed in males than in females. In the SCN of females, too, AR is localized within the cells of the core. Gonadectomy in males reduces AR levels within the SCN to levels seen in intact females (Karatsoreos et al., 2007), while testosterone replacement restores AR to levels seen precastration. Ovariectomy in females does not affect AR levels in the SCN (Iwahana, Karatsoreos, Shibata, and Silver, 2008). The function of ARs within the SCN is unclear, although it has been suggested that androgens serve to alter the responses of the SCN to light. Indeed, castration reduces the FOS expression to a phase-shifting light pulse, and DHT replacement restores levels to those seen in intact animals (Karatsoreos et al., 2007). More recently, it has been found that gonadectomy affects the induction of clock genes within the SCN (Karatsoreos, Butler, LeSauter, and Silver, 2011). After a phase-advancing light pulse, gonadectomy increased light induced mPer1 but not mPer2 expression compared to intact animals. After a light pulse at a phase delay point, gonadectomy decreased light induced mPer2 but did not affect mPer1. Gonadectomized animals treated with DHT showed a pattern of molecular responses to light similar to intact animals. The finding that almost all GRP cells within the SCN contain AR may also be significant to our understanding of AR action within the SCN. GRP is known to be involved in phase resetting by photic stimuli. The GRP receptor (GRPr) is expressed principally in the rhythmic shell of the SCN. Mice lacking the gene for this receptor display blunted phase shifts, and light- induced expression of Per1 and Per2 in the SCN is decreased, consistent with the finding that gonadectomy reduces FOS expression after photic stimulation (Karatsoreos, Romeo, McEwan, and Silver, 2006; Aida et al., 2002; Karatsoreos et al., 2007). In addition to its role in the SCN 10 response to light, GRP plays a role in the synchronization of SCN oscillator networks as well as within the SCN (Karatsoreos et al., 2007). Karatsoreos et al. (2007) have suggested that the loss of coherence of locomotor activity after gonadectomy could indicate that androgenic feedback to GRP cells may be necessary for the coordination of individual oscillators in the SCN. Finally, gonadectomy alters both synaptic protein expression and astrocyte morphology within the SCN (Karatsoreos et al., 2011). Astrocytes are prevalent throughout the SCN and can affect neural connectivity (Van den Pol, 1980; Oliet, Piet, Poulain, and Theodosis, 2004). After castration, glial fibrillary acidic protein increased, while decreases were seen in the expression of the synaptic proteins synaptophysin and postsynaptic density 95; DHT replacement restored levels to those seen pre-castration (Karatsoreos, 2011). Changes in astrocytes and synaptic proteins may alter oscillator coupling, which in turn may affect rhythm precision (Hagenauer and Lee, 2011). This model is consistent with the findings that androgens enhance rhythm precision, and castration decreases the precision of free-running locomotor rhythms (Karatsoreos et al., 2007). Thus, growing evidence shows that androgens may alter the structure and function of the SCN, and therefore circadian behaviour. More research is needed to understand the role of SCN AR in endocrine function and determine whether it serves a purpose for behaviour beyond that of neuroendocrine feedback (Hagenauer and Lee, 2011).

1.4 The Role of AR in the SCN In sum, alterations in levels of androgens have been shown to affect circadian responses, and sex differences in these responses have been documented as well. Although some of these circadian responses are known to be SCN-dependent, the site of hormone action remains poorly understood. Recent work shows clear evidence of androgens altering SCN organization and function (Hagenauer and Lee, 2011; Silver and Karatsoreos, 2007). However, it remains to be directly tested whether AR action within the SCN itself can account for these observed effects. It is possible that the effects of androgens on other tissues are indirectly responsible for the observed changes in SCN physiology. Previous work suggests that androgens affect various circadian output rhythms, such as entrained phase, period, rhythm precision, sensitivity to phase shifts, and amplitude of activity (Gorman and Lee, 2002; Karatsoreos and Silver, 2007; Daan et 11 al., 1975). Entrained phase, phase shifting in response to environmental time cues, precision and period are all believed to be regulated by the SCN. Many of these rhythm properties are associated with the function of the core of the SCN (Hagenauer and Lee, 2011), where AR is highly localized in mice (Karatsoreos et al., 2007). Therefore, it has been proposed that androgens act directly on these AR-containing cells within the SCN. It has also been argued that, because the effects of castration on behaviour are seen only after 5-7 days, classical steroid receptor action, rather than rapid non-genomic effects of androgens, is responsible for these changes (Karatsoreos et al., 2011). Furthermore, androgen administration has been shown to affect aspects of SCN physiology and function, such as photic induction of clock genes and c- Fos, as well as glial cell proliferation and synaptic protein expression (Karatsoreos et al., 2011; Karatsoreos et al., 2007). It seems probable that androgenic effects directly on the SCN are responsible for these changes in SCN physiology, which in turn are causing the observed effects on behaviour. Although an SCN site of action is most parsimonious, the SCN receives input from other androgen sensitive tissues, namely the retina and dorsal raphe nucleus. So, it is possible androgen effects on these tissues are causing the observed changes in SCN physiology. By contrast, amplitude of activity is thought to be controlled by the medial preoptic area (MPOA), not the SCN (Pittendrigh and Daan, 1976; Daan and Oklejewicz, 2003). Previous research has shown that testosterone but not DHT restores overall amount of activity in gonadectomized mice. It is thought that testosterone’s effect on total activity in mice may be through its conversion to estradiol, with the MPOA as the site of action. Thus, the effects of testosterone on circadian rhythms can be SCN-dependent or independent, through action directly on AR or by aromatization to estradiol. Finally, sex differences in AR expression within the SCN have been found, with greater AR expression found in males than in females (Iwahana et al., 2008). These sex differences disappear after gonadectomy, and are restored following treatment with testosterone, although treatment with the same dose of hormone produces a greater response in males than in females. In the absence of circulating androgens, male and female mice have different temporal patterns of activity. It has been hypothesized that androgens acting on AR receptors in the SCN allow males and females to display similar patterns of behaviour (Iwahana et al., 2008), which might facilitate reproduction in the wild. 12

1.5 The Present Study Transgenic mice that selectively express androgen receptors provide a useful model for the study of the site of hormone action on circadian responses. The present study will compare mice that overexpress AR in neurons only to mice that have normal levels of androgen expression. Using a Cre-lox system, we have developed a transgenic mouse that selectively overexpresses AR only where Cre is present. Cre recombinase, originally derived from bacteriophage P1, is an enzyme that recombines pairs of short nucleotide sequences called loxP sites, excising the sequences flanked by the two loxP sites. The use of the Cre/loxP recombination system allows for the development of transgenic models whose transgene expression is limited to specific tissues or developmental stages (Wilson and Kola, 2001). Mice that express Cre under the promoter of Nestin, a nervous system-specific promoter, allow for a specific transgene to be transcribed only in neurons (Tronche et al., 1999). The AR26 transgene is an androgen receptor transgene with 26 polyglutamine repeats, which falls within the normal range. AR26 is driven by the cytomegalovirus (CMV) promoter, a strong promoter that drives transgene expression everywhere in the organism. The AR26 transgene has a neo stop sequence surrounded by loxP sites, which prevents transcription of the transgene on its own. However, when Nestin-Cre mice are crossed with AR26 mice, the neo stop sequence is excised in the nervous system, allowing AR to be overexpressed in neurons only (Nestin-AR). The present study was designed to test the hypothesis that neural ARs regulate the circadian system and to evaluate how features of the circadian system that are known to be androgen-dependent interact with our overexpression model by manipulating hormone levels in these animals. Because androgens have been implicated in modulating the SCN’s response to light, we explored whether altering AR expression in neurons only affects c-Fos expression within the SCN in response to a light pulse. c-Fos is a marker of neural activity, and differences in the number of c-Fos immunoreactive cells between mutants and wildtypes would suggest that expression of AR in neurons may be sufficient to alter the SCN’s response to light. Alternatively, if no differences are seen between Nestin-AR and wildtype mice, it is possible that AR in non- neural tissues is necessary to alter light-induced Fos expression within the SCN. By examining 13 gonadectomized animals and animals with DHT replacement, we also provide a test of whether any observed effects on the circadian system are dependent on circulating androgens, and whether such effects of androgens are due to their direct action on ARs, or by aromatization to estradiol and subsequent action on estrogen receptors (ERs). Finally, after gonadectomy, male mice show a decrease in overall amount of activity, and this response is restored by testosterone replacement. We examined whether altered expression of neural ARs affects locomotor activity in mice by measuring overall daily activity. Mutant and wild-type males received castration surgery and either DHT or replacement. Intact Nestin-AR and wildtype males will also be compared in order to characterize the circadian system in this transgenic model. Behavioural rhythms under entrained conditions will be measured, as well as c-Fos expression in response to a light pulse. In keeping with previous findings, it is predicted that gonadectomy will result in altered circadian behaviours, including decreased levels of overall activity. In response to a 30- light pulse during the early night (at ZT13.5-14), it is expected that, consistent with previous results, castration (GDX) will lead to blunted Fos expression. It is further predicted that increased sensitivity to AR in Nestin-AR mice will result in increased Fos response to a light pulse. By comparing physiological responses to light in these mice, such as Fos response, we will add to current knowledge about the contribution of neural AR action on SCN function, which is important for expanding our current understanding of the mechanisms of androgenic action on SCN circuits.

Methods 2.1 Animals and Housing Littermate wildtype and Nestin-AR transgenic animals were generated in our colony by mating mice heterozygous for Nestin-Cre and AR26 on a C57Bl ⁄ 6J background. Genotyping was performed by PCR amplification using tail samples. For Nestin-AR genotyping, primers (forward) AGGATCTCCTGTCATCTCACCTTGCTCCTG and (reverse) AAGAACTCGTCAAGAAGGCGATAGAAGGCG were used to detect the AR26 transgene, and primers (forward) GCGGTCTGGCAGTAAAAACTATC and (reverse) 14

GTGAAACAGCATTGCTGTCACTT were used to detect the Nestin-Cre transgene. Animals were housed with a 12-h light, 12-h dark cycle (12:12 LD) with a dim red light (<1 lux) allowing for animal maintenance. Animals were housed singly in clear polypropylene cages containing corn cob bedding and given ad libitum access to food and water. All procedures were conducted in accordance with the guidelines of the University Animal Care Committee at the University of Toronto.

2.2 Experimental Design A total of 72 experimental animals were used, and all subjects were included in both the behaviour testing and immunohistochemistry (IHC) portions of the experiment. Groups for behaviour testing consisted of GDX (gonadectomized), GDX + DHT (gonadectomized with DHT replacement), and INT (gonadally intact) animals, for both Nestin-AR and wildtype mice (n = 12 per group). Investigation of Fos response to a light pulse included these six groups, as well as matching control groups not receiving a light pulse (n = 6 per group). Behaviour and tissue from 72 animals were collected, but due to tissue quality and missing data, not all animals were included in analyses of behaviour and IHC. All 72 animals were included in analyses of physiological measures. For a list of groups and their sizes, see Tables 1-2.

2.3 Gonadectomy (GDX) and Hormone Replacement 2.3.1 Steroid Implants Dihydrotestosterone (DHT) implants were prepared to mimic physiological levels of hormones as described previously by Karatsoreos et al. (2011). SILASTIC capsules (inner diameter 1.02 mm, outer diameter 2.16 mm; Dow Corning Corp., Midland, MI) were filled with 10 mm of crystalline DHT, sealed with medical adhesive (Factor II, Inc, Lakeside, AZ), and allowed to dry overnight. Capsules were primed in a 37°C 0.9% saline bath for 36 h before implantation. 15

2.3.2 Gonadectomy Mice were deeply anesthetized under inhalant anaesthesia (1-2% isoflurane) and gonadectomized by abdominal incision and removal of both testes. Following removal of the testes, the incision was closed with wound clips. Immediately after castration, animals were implanted with DHT or empty capsules. Animals were allowed to recover for at least 10 d before experimental manipulation.

2.4 Behavioural Testing For behavioural experiments, animals were individually housed in polypropylene cages placed in a frame equipped with two arrays of 16 × 16 infrared photo-beams. Boxes were separated by opaque screens to prevent mice from seeing each other. Arrays were connected to a via an interface that detects photo-beam interruptions (beam breaks) to record activity levels. Locomotor activity in the form of beam breaks was recorded in 10-min bins over the course of 4 days under 12:12 LD conditions. A dim red light (<1 lux) allowed for daily animal maintenance, and sound was masked by a white-noise generator (91 spl).

2.5 Neural Activation Following Exposure to Light On the sixth day of behaviour testing, animals were transferred to an experimental room and exposed to a 30-min light pulse (LP) between ZT13.5-14 (800 lux). Animals were then returned to darkness for 60 min and then killed. Control animals (non light pulse, NLP) were also moved from the behaviour testing room to a separate experimental room but were not exposed to light. Sixty after the end of the light pulse, animals were weighed, then deeply anesthetized (Avertin, 40mg/100g ip) under a dim red light, their heads covered with a lightproof hood, and transcardially perfused with phosphate-buffered saline (PBS) followed by 4% paraformaldehyde in 0.1 M phosphate buffer, pH 7.3. Blood and tails were collected, and anogenital distance, seminal vesicle weight, testes weight (for intact animals) measurements were taken from the animals. Brains were extracted and post-fixed in 4% phosphate-buffered paraformaldyde for 2 hours and then cryoprotected in 20% sucrose in phosphate buffer overnight 16 prior to sectioning. Brains were then blocked (cerebellum and olfactory bulbs removed) and sectioned on a freezing microtome at 35 µm into four series. Sections were stored in cryoprotectant at -20°C until ICC.

2.6 c-Fos Immunohistochemistry One of the four series was processed for c-fos immunohistochemistry to evaluate neuronal activation patterns in response to a light pulse. Tissue was washed three times for 5 min each in PBS, followed by a 90-minute incubation in blocking solution at room temperature (4%

NGS with 0.3% TritonX and 0.3% H2O2 in PBS). Tissue was then rinsed in PBS for 3 x 5 minutes and incubated in c-Fos primary antibody (1:500 rabbit anti-c-fos polyclonal antibody;

Santa Cruz Biotechnology, Inc., cat # sc-52) in blocking solution without H2O2 at 4°C for approximately 24 hours. Tissue was rinsed in PBS for 3 x 5 minutes, then incubated for 90 minutes at room temperature in a secondary antibody (goat anti-rabbit 1:200; Vector Laboratories) in PBS with 0.3% TritonX and 2% NGS. Tissue was rinsed again in PBS for 3 x 5 minutes, then incubated at room temperature for 90 minutes in avidin-biotin complex (ABC, Vector Laboratories). Sections were washed again for 3 x 5 minutes in PBS, and then placed in a diaminobensizine reaction to visualize c-Fos immunoreactivity. Tissue was then mounted onto gelatin-coated slides, and dehydrated and coverslipped using Permount (Fisher Scientific, Mississauga, ON, Canada). All 12 experimental groups were stained together to eliminate inter- run variability. c-Fos ICC included a no-primary control for each animal, and staining was found to be absent for controls.

2.7 Androgen Receptor Immunohistochemistry To confirm that Nestin-AR mutants overexpress AR, androgen receptor immunoreactive cells were compared between a small subset of animals. Tissue was washed for 6 x 5 minutes in PBS, followed by 3 x 10 minute incubations in 0.3% hydrogen peroxide. Following 3 x 5 minute washes in PBS, sections were incubated in 10% normal goat serum (NGS) in PBS with 0.3% TritonX100 and 0.1% pig gelatin (PBS-GT) for 45 minutes at room temperature. Tissue was then incubated for approximately 48 hours in androgen receptor primary solution made in 4% NGS 17 and PBS-GT at 4°C (AR antibody 1:100 concentration; Epitomics 3165-1). Following incubation in the primary antibody, tissue was washed 3 x 5 minutes in PBS, then incubated in goat anti- rabbit biotinilated (GARB) secondary solution made in 10% NGS in PBS (3.1ul of GARB in 10mL of solution; Vector laboratories BA-1000). Sections are then washed 3 x 5 minutes in PBS then incubated in ABC in PBS (Vector laboratories PK-3100). Sections are washed again, 3 x 5 minutes in PBS, then placed in a diaminobensizine reaction for 2-8 minutes. Tissue was then mounted onto pig gelatin coated slides, dehydrated, and coverslipped using permount (Fisher brand SP15-1000). Qualitative analyses performed comparing Nestin-AR and wildtype littermates demonstrate that Nestin-AR males have more AR-ir cells (see Figure 3).

2.8 Data Analysis 2.8.1 Brain Analyses Analyses were performed on slides coded to conceal the genotype and experimental condition of the animal. Images were taken at 400x magnification on an Olympus BX-51 compound light with a mounted Olympus D70 CCD camera attached to a computer. The microscope was allowed to stabilize for 15 min prior to use. Photographs were taken of either plate 35 or 36 for each animal. Two photographs, one of the left and right side of the same section, were taken for each animal. Automated counts for the left and right side of one section were generated independently using ImageJ software from NIH (Bethesda, Maryland, USA), and then a total for each animal was computed from the bilateral measurements. See table 3 for group means and standard errors for number of Fos-ir cells counted for genotype and hormone condition.

2.8.2 Behavioural Analyses Behavioural analyses were performed on 5 consecutive days of data at least 10 d after any manipulation. Amount of daily activity was calculated as the number of beam breaks per day, averaged over 5 days. Refer to table 4 for group means and standard errors for the total number of beam breaks recorded. 18

2.8.3 Statistical Analyses Behavioural measurements, seminal vesicle weights, anogenital distance, and body weight were analyzed using two-way analyses of variance (ANOVAs) (hormone condition-by- genotype). Testes weight was analyzed using a one-way ANOVA with genotype as a factor. Fos- ir cells in the SCN were analyzed by a three-way ANOVA (genotype-by-hormone condition-by- light condition) using start time of light pulse or sham light pulse as a covariate. Post-hoc comparisons using Fisher’s LSD were performed to investigate main effects or interactions for the ANOVAs. Planned comparisons were performed using Student’s t-tests. Results were considered significant at p < 0.05.

Results 3.1 Somatic Measures at time of sacrifice was used as a covariate for all physiological analyses. A two-way analysis of variance (ANOVA) revealed a significant effect of genotype, F(1,65) = 19.814, p < . 001, on body weight, with WT animals weighing more than TG mice. A main effect of hormone treatment was found for body weight, F(2,65) = 6.218, p < .01. Post-hoc analyses found that GDX mice weighed significantly less than both INT and DHT-treated mice, p < .01. No differences in body weight were found between DHT and INT animals. No significant interaction between hormone treatment and genotype was found, F(2,65) = 0.444, ns. Significant differences were found for seminal vesicle weights between hormone treatments, F(2,65) = 72.131, p < .001. The seminal vesicles of GDX mice weighed significantly less than those of both DHT and INT mice, p < .001. Seminal vesicle weights of DHT and INT mice also differed, with seminal vesicles of INT animals weighing significantly more than those of DHT mice, p < .01. No main effect of genotype, F(1,65) = 1.117, ns, or interaction between genotype and hormone treatment, F(2,65) = 2.689, ns, was found. A significant effect of hormone treatment was found for anogenital distance (AGD), F(2,65) = 18.472, p < .001. Further comparisons indicated that INT animals had an AGD that was significantly larger than both GDX and DHT animals, p < .001, and that the AGD of DHT 19 animals was larger than that of GDX animals, p < .05. No effect of genotype, F(1,65) = 0.382, ns, or interaction between genotype and hormone treatment, F(2,65) = 0.608, ns, was found for AGD. No effect of genotype was found on testes weight for INT animals, F(1,21) = 0.023, ns. Taking body weight into account did not produce differences between TG and WT animals on any physiological measures.

3.2 Behaviour Age at time of testing and testing cohort were used as covariates for all ANOVAs. A significant main effect of hormone treatment on overall activity was found, F(2,60) = 3.632, p < . 05. Post-hoc analyses revealed that GDX mice have significantly lower levels of overall activity than both INT and DHT groups, p < .05. No significant difference in activity levels was found between INT and DHT animals. No significant effect of genotype on activity was found, F(1,60) = 0.21, ns. An interaction between genotype and treatment was not found to be significant, F(2,60) = 2.964, ns. Planned comparisons found no differences for WT between GDX and INT, t(21) = 0.056, ns, GDX and DHT, t(21) = 0.216, ns, or DHT and INT, t(20) = 0.135, groups. For Nestin-AR mice, no differences were found between INT and DHT groups, t(20) = 0.501, ns. However, GDX Nestin-AR mice were found to have significantly lower levels of activity than both INT, t(20) = 2.703, p < .05, and DHT-treated mice, t(22) = 2.373, p < .05.

3.3 Neural Activation Time of light pulse administration was used as a covariate for all ANOVAs. A significant main effect of lighting condition was found, F(1,52) = 59.350, p < .001, indicating that animals exposed to a light pulse show a higher Fos response than animals that are not exposed to light. There was no main effect of genotype, F(1,52) = 0.161, ns, or hormone treatment, F(2,52) = 0.499, ns, on c-Fos immunoreactivity. A significant interaction between genotype and lighting condition was found for Fos response, F(1,52) = 5.504, p < .05, indicating that Fos response differs between TG and WT 20 animals depending on lighting conditions. WT animals displayed less Fos activity than TG in the NLP condition and more activity than TG in the LP condition, indicating that the increase in Fos expression after exposure to a light pulse is greater for WT than for TG animals. A significant interaction between hormone treatment and lighting condition was also found for Fos response, F(2,52) = 4.197, p < .05. No significant interaction was found between genotype, lighting condition, and hormone treatment, F(2,52) = 0.145, ns. For WT GDX animals, planned comparisons indicated a significant difference between LP and NLP conditions, t(8) = 5.463, p < .01. Similarly, LP and NLP conditions were found to differ for TG GDX animals, t(10) = 4.047, p < .01. Significant differences were also found between LP and NLP conditions for both WT INT, t(7) = 3.103, p < .05, and TG INT, t(10) = 2.298, p < .05, groups. However, while LP and NLP conditions differed for WT DHT animals, t(10) = 2.661, p < .05, no difference between lighting conditions was found for TG DHT animals, t(8) = 1.167, ns, indicating that exposure to light does not significantly increase Fos expression in these mice.

Discussion Analysis of physiological measures of androgen production found that GDX animals have significantly lower body weight, lower seminal vesicle weight, and smaller anogenital distance than both INT and DHT-treated animals, confirming the efficacy of the hormone treatment. As expected, animals exposed to a half- light pulse at the onset of nighttime activity demonstrated higher levels of Fos expression than animals not exposed to light. Contrary to expectations, however, we found a greater increase in Fos expression after light exposure in WT than Nestin-AR mice. Moreover, for Nestin-AR animals treated with DHT, it was found that exposure to a light pulse does not significantly increase Fos-ir over the levels of unmanipulated animals. Behavioural analyses revealed that castration had an effect on overall activity levels, with GDX animals displaying lower levels of activity than DHT or INT mice. In particular, while 21 hormone treatment did not significantly alter locomotor activity in WT mice, GDX in Nestin-AR mice led to lower overall activity when compared to INT and DHT-treated counterparts. The results of the present study provide further evidence of a role for androgens in modulating the function of the SCN, in particular its response to light. Interestingly, Nestin-AR mice showed less of an increase in Fos expression after exposure to light than did WT mice. Because previous research has demonstrated that castration reduces FOS response after a light pulse, we predicted that increased sensitivity to AR would lead to higher levels of FOS expression after light exposure. Overexpression of a WT protein has been shown to exert toxic effects in several animal models, and therefore it is possible that AR is having toxic effects on neurons in these mice, resulting in smaller cell nuclei, greater cell death, or other abnormalities in the SCN. However, in this case one might expect lower levels of activity overall in the SCN of Nestin-AR mice regardless of light exposure, which the present study did not find. Investigation of neuroanatomical differences between Nestin-AR and WT SCNs would provide insight as to whether this hypothesis is valid. Conversely, it is possible that overexpression of AR leads to more baseline activity in the SCN generally, although it is not clear why this would not lead to a greater level of Fos expression in both LP and NLP conditions. Examining neural activity in nearby hypothalamic nuclei might shed light on whether AR overexpression leads to increased activity more generally. The present results suggest a possible developmental effect of increased androgen sensitivity on the SCN, as reduced Fos expression after exposure to a light pulse was found for Nestin-AR mice without taking hormone treatment into consideration. This apparent reduction in sensitivity to light might be a compensatory mechanism in response to increased sensitivity to androgens throughout development. As previously mentioned, almost all GRP cells in the SCN contain AR (Karatsoreos et al., 2007) and it has been suggested that gonadal steroid feedback onto retinorecipient GRP cells may be required to coordinate individual SCN oscillators. It is possible that inappropriate levels of androgenic feedback, regardless of direction, impair the function of these cell populations, as previous research has suggested a role for these neurons in the SCN response to light, synchronization of SCN oscillator networks, and intra-SCN communication (Karatsoreos et al., 2007). research can address this question by 22 examining whether the differences in Fos expression observed here between Nestin-AR and WT mice are specific to the core of the SCN, and in particular by looking at the peptidergic content of the Fos expressing cells in these animals. It is also interesting to note that Nestin-AR animals treated with DHT did not show significantly higher levels of Fos expression when exposed to light, suggestive of an activational effect of androgens. It is not clear why this effect is seen in DHT-treated Nestin-AR mice and not INT mice, however it is important to note that DHT has higher levels of biological activity at androgen receptors in the brain than T (Martini, 1982). Furthermore, in Nestin-ARKO mice (with AR knocked-out in the nervous system only), knock- out males were found to have slightly increased circulating levels of T when compared to wildtype males (Raskin et al., 2009; Juntti, 2010). It is therefore possible that androgen production in intact Nestin-AR mice is lower than that of WT mice, however this is not reflected in the seminal vesicles weights of these animals. Regardless, if Nestin-AR mice treated with DHT have the highest levels of androgenic activity on SCN AR, the lack of significant response to a phase delaying light pulse supports the idea that inappropriate levels of AR activity in the SCN alter the SCN’s response to light. It would also be of interest to investigate whether free-running period, precision of activity onset, activity bout length (α), and early evening activity bout differ between WT and Nestin-AR mice. As mentioned previously, precision and period are thought to be controlled by the SCN directly (Pittendrigh and Daan, 1976), and are affected by castration. Several rodents are seasonal breeders, including some mice, voles, and hamsters. In hamsters, short day-length induced testicular regression results in loss of precision of onset of activity and cohesion of the daily activity bout (Morin and Cummings, 1981), and leads to decreased behavioural sensitivity to the activational effects of gonadal hormones (Morin and Zucker, 1978). While the gonadal axis of the mouse does not respond to day-length cues, mice do exhibit seasonal changes in reproduction in response to change in the availability of specific nutrients as well as seasonal variation in temperature (Bronson, 1979). Furthermore, the SCN of the responds to changes in day length (Inagaki et al., 2007), and it has been found that mice may retain latent photoperiodic responses (Nelson, 1990). In light of these findings, it has been suggested that androgenic feedback to the SCN plays a functional role in reproductive 23 behaviours (Iwahana et al., 2008; Hagenauer and Lee, 2011; Karatsoreos et al., 2007). The circadian system can act to facilitate behaviours such as mating and social coordination; gonadal hormones can signal changes in the social and physical environment of an animal, which can in turn influence the circadian system (Hagenauer and Lee, 2011). For example, testosterone has been found to suppress circadian responses to social cues, specifically olfactory stimuli, in male and female rodents (Jechura et al., 2003). In male mice, the role of androgens may be to promote overlapping temporal niches in the patterns of daily activity between male and female mice (Iwahana et al., 2008). It has been demonstrated previously (Iwahana et al., 2008; Karatsoreos et al., 2007) that castration eliminates the onset bout of nightly activity in mice, whereas both female and intact male mice both show running wheel activity at the start of night. This effect of castration may reduce the possibility of male and female mice meeting in the wild. The work of Daan et al. (1975) and Karatsoreos et al. (2007) has demonstrated that the early evening bout of activity is androgen dependent, which is complemented by our own findings and previous work showing that castration affects Fos expression to a phase delaying light pulse (i.e. at the onset of nighttime activity). Indeed, many examples exist of sex differences in hormonal conditions acting to reduce sex differences in behaviour (De Vries and Boyle, 1998), and androgens acting on SCN AR might function to allow males and females to show similar temporal patterns of behaviour (Iwahana et al., 2008). Looking at these measures will not only be helpful in determining what behavioural outcomes are associated with the observed effects seen in Nestin- AR mice, but also in determining what function neural AR has in controlling reproductive behaviours. An unexpected finding of this study was that GDX Nestin-AR mice display significantly lower levels of overall activity than either DHT or INT Nestin-AR mice. Overall activity level in males is thought to be modulated largely by estradiol acting on target receptors in the medial preoptic area (Ogawa et al., 2003; Morgan et al., 2004; Fahrbach et al., 1985), as few estrogen receptors have been found in the SCN (Shughrue et al., 1997; Shughrue and Merchenthaler, 2001). However, several studies have found evidence that replacement with DHT, which cannot be aromatized to estrogen, does increase total daily activity above levels observed in castrates (Karatsoreos et al., 2007; Iwahana et al., 2008), indicating some effect of androgens on overall 24 activity levels. The medial preoptic area (MPOA) has also been identified as a possible source of androgenic input to the SCN (Hagenauer and Lee, 2011), and it is possible that the effect seen in Nestin-AR mice is due to extra-SCN action of AR in the MPOA. Since administration of DHT has been shown to increase activity levels after castration, it is interesting to speculate whether increased AR sensitivity would result in higher levels of activity; on this view, lower levels of activity seen in GDX Nestin-AR animals may represent a compensatory mechanism by which Nestin-AR animals adjust to higher levels of AR action throughout development. It is interesting to note that INT Nestin-AR animals do show slightly higher levels of overall activity than wildtypes, however this difference is not significant. Further investigation is needed to determine whether this observed effect on activity level is SCN-dependent or independent, and how AR is acting to reduce activity levels in Nestin-AR GDX mice. The current results indicate that expression of AR in neurons alone may be sufficient to alter the SCN’s response to light. However, a role for non-neural tissues affecting the function of the SCN cannot be ruled out. Nestin-Cre is expressed only in neuronal and possibly glial precursor cells (Tronche et al., 1999). Future studies can address the question of whether AR in non-neural tissues can affect SCN function by comparing Nestin-AR mice to mice that overexpress AR globally. Additionally, although previous research has indicated the SCN as the likely site of action of AR, the SCN also receives input from several hormone-sensitive neural tissues, including the retina, intergeniculate leaflet, and the raphe nuclei (Karatsoreos and Silver, 2007; Hagenauer and Lee, 2011). While the intergeniculate leaflet has not been identified as a probable site of androgenic input to the SCN (Karatsoreos and Silver, 2007; Hagenauer and Lee, 2011), moderate levels of AR staining in the dorsal raphe nucleus (DRN) have been found for both male rats and mice (Shen et al., 2004). Previous research has shown evidence of ovarian hormones affecting the DRN’s response to light in female rats (Abizaid, 2005), however the effects of androgens on this nucleus have not been investigated. Another possible site of action for AR is the retina, which has been shown to possess mRNA for AR (Wickham et al., 2000). Nestin expression has been found in the neuronal and glial cells of the developing retina (Walcott and Provis, 2003), however levels of retinal AR have not been characterized in Nestin-AR mice. It would be of interest to see whether patterns of neural activation along input pathways to the 25

SCN differ between WT and Nestin-AR mice to determine whether the observed effects on the SCN are direct or indirect. Ultimately, the present study contributes to the growing evidence of androgenic effects on the circadian system, in particular in modulating the SCN’s response to light. Transgenic animals that selectively express AR will provide a useful model to better understand the influence of androgens on circadian function and investigate a possible SCN site of action. 26

References Abrahamson, E. E., & Moore, R. Y. (2001). Suprachiasmatic nucleus in the mouse: retinal innervation, intrinsic organization and efferent projections. Brain research, 916(1), 172-191. Aida, R., Moriya, T., Araki, M., Akiyama, M., Wada, K., Wada, E., & Shibata, S. (2002). Gastrin-Releasing Peptide Mediates Photic Entrainable Signals to Dorsal Subsets of Suprachiasmatic Nucleus via Induction of Period Gene in Mice. Molecular pharmacology, 61(1), 26-34. Antle, M. C., & Silver, R. (2005). Orchestrating time: arrangements of the brain circadian clock. Trends in neurosciences, 28(3), 145-151. Arble, D. M., Copinschi, G., Vitaterna, M. H., Van Cauter, E., & Turek, F. W. (2011). Circadian Rhythms in Neuroendocrine Systems. Handbook of Neuroendocrinology, 271. Bruce, V. G. (1960). Environmental entrainment of circadian rhythms. In Cold Harbor Symposia on Quantitative Biology (Vol. 25, pp. 29-48). Cold Spring Harbor Laboratory Press. Bryant, D. N., LeSauter, J., Silver, R., & Romero, M. T. (2000). Retinal innervation of calbindin- D28K cells in the hamster suprachiasmatic nucleus: ultrastructural characterization. Journal of biological rhythms, 15(2), 103-111. Buijs, R. M., Van Eden, C. G., Goncharuk, V. D., & Kalsbeek, A. (2003). The biological clock tunes the organs of the body: timing by hormones and the autonomic nervous system. Journal of Endocrinology, 177(1), 17-26. Butler, M. P., Kriegsfeld, L. J., & Silver, R. (2009). Circadian regulation of endocrine functions. Hormones, brain, and behavior, 473-505. Cheng, M. Y., Bullock, C. M., Li, C., Lee, A. G., Bermak, J. C., Belluzzi, J., ... & Zhou, Q. Y. (2002). Prokineticin 2 transmits the behavioural circadian rhythm of the suprachiasmatic nucleus. Nature, 417(6887), 405-410. Daan, S., & Oklejewicz, M. (2003). The precision of circadian clocks: assessment and analysis in Syrian hamsters. international, 20(2), 209-221. 27

Daan, S., Damassa, D., Pittendrigh, C. S., & Smith, E. R. (1975). An effect of castration and testosterone replacement on a circadian pacemaker in mice (Mus musculus). Proceedings of the National Academy of , 72(9), 3744-3747. de la Iglesia, H.O., Blaustein, J.D., & Bittman, E.L. (1999). Oestrogen receptor-alpha- immunoreactive neurones project to the suprachiasmatic nucleus of the female Syrian hamster. J. Neuroendocrinol. 11(7):481–490. Ellis, G. B., & Turek, F. W. (1983). Testosterone and photoperiod interact to regulate locomotor activity in male hamsters. Hormones and behavior, 17(1), 66-75. Everett, J. W., & Sawyer, C. H. (1950). A 24-hour periodicity in the “LH-release apparatus” of female rats, disclosed by barbiturate sedation. Endocrinology, 47(3), 198-218. Fernández-Guasti, A., Kruijver, F. P., Fodor, M., & Swaab, D. F. (2000). Sex differences in the distribution of androgen receptors in the human hypothalamus. The Journal of comparative neurology, 425(3), 422-435. Gillette, M. U. (1991). SCN electrophysiology in vitro: rhythmic activity and endogenous clock properties. Suprachiasmatic Nucleus, the Mind’s Clock, 6, 125-143. Gray, G. D., Södersten, P., Tallentire, D., & Davidson, J. M. (1978). Effects of lesions in various structures of the suprachiasmatic-preoptic region on LH regulation and sexual behavior in female rats. Neuroendocrinology, 25(3), 174-191. Green, D. J., & Gillette, R. (1982). Circadian rhythm of firing rate recorded from single cells in the rat suprachiasmatic brain slice. Brain Research; Brain Research. Groos, G., & Hendriks, J. (1982). Circadian rhythms in electrical discharge of rat suprachiasmatic neurones recorded in vitro. Neuroscience letters, 34(3), 283-288. Gundlah, C., Kohama, S. G., Mirkes, S. J., Garyfallou, V. T., Urbanski, H. F., & Bethea, C. L. (2000). Distribution of (ERbeta) mRNA in hypothalamus, midbrain and temporal lobe of spayed macaque: continued expression with hormone replacement. Brain research. Molecular brain research, 76(2), 191. Hagenauer, M. H., & Lee, T. M. (2011). Time for Testosterone: The Suprachiasmatic Nucleus Gets Sexy. Endocrinology, 152(5), 1727-1730. 28

Hakim, H., DeBernardo, A. P., & Silver, R. (1991). Circadian locomotor rhythms, but not photoperiodic responses, survive surgical isolation of the SCN in hamsters. Journal of biological rhythms, 6(2), 97-113. Hamada, T., Antle, M. C., & Silver, R. (2004). Temporal and spatial expression patterns of canonical clock genes and clock-controlled genes in the suprachiasmatic nucleus. European Journal of Neuroscience, 19(7), 1741-1748. Hamada, T., LeSauter, J., Venuti, J. M., & Silver, R. (2001). Expression of Period genes: rhythmic and nonrhythmic compartments of the suprachiasmatic nucleus pacemaker. The Journal of Neuroscience, 21(19), 7742-7750. Inouye, S. T., & Kawamura, H. (1979). Persistence of circadian rhythmicity in a mammalian hypothalamic" island" containing the suprachiasmatic nucleus. Proceedings of the National Academy of Sciences, 76(11), 5962-5966. Iwahana, E., Karatsoreos, I., Shibata, S., & Silver, R. (2008). Gonadectomy reveals sex differences in circadian rhythms and suprachiasmatic nucleus androgen receptors in mice. Hormones and behavior, 53(3), 422-430. Jobst, E. E., & Allen, C. N. (2002). Calbindin neurons in the hamster suprachiasmatic nucleus do not exhibit a circadian variation in spontaneous firing rate. European Journal of Neuroscience, 16(12), 2469-2474. Karatsoreos, I. N., & Silver, R. (2007). Minireview: the neuroendocrinology of the suprachiasmatic nucleus as a conductor of body time in mammals. Endocrinology, 148(12), 5640-5647. Karatsoreos, I. N., Butler, M. P., LeSauter, J., & Silver, R. (2011). Androgens modulate structure and function of the suprachiasmatic nucleus brain clock. Endocrinology, 152(5), 1970-1978. Karatsoreos, I. N., Romeo, R. D., McEwen, B. S., & Silver, R. (2006). Diurnal regulation of the gastrin-releasing peptide receptor in the mouse circadian clock. European Journal of Neuroscience, 23(4), 1047-1053. Karatsoreos, I. N., Wang, A., Sasanian, J., & Silver, R. (2007). A role for androgens in regulating circadian behavior and the suprachiasmatic nucleus. Endocrinology, 148(11), 5487-5495. 29

Karatsoreos, I. N., Yan, L., LeSauter, J., & Silver, R. (2004). Phenotype matters: identification of light-responsive cells in the mouse suprachiasmatic nucleus. The Journal of neuroscience, 24(1), 68-75. Kashon, M. L., Arbogast, J. A., & Sisk, C. L. (1998). Distribution and hormonal regulation of androgen receptor immunoreactivity in the forebrain of the male European ferret. The Journal of comparative neurology, 376(4), 567-586. Klein, D. C. (1985). Photoneural regulation of the mammalian pineal gland. In Ciba Foundation symposium (Vol. 117, p. 38). Klein, D. C., Smoot, R., Weller, J. L., Higa, S., Markey, S. P., Creed, G. J., & Jacobowitz, D. M. (1983). Lesions of the paraventricular nucleus area of the hypothalamus disrupt the suprachiasmatic spinal cord circuit in the melatonin rhythm generating system. Brain Research Bulletin, 10(5), 647-652. Ko, C. H., & Takahashi, J. S. (2006). Molecular components of the mammalian circadian clock. Human molecular genetics, 15(suppl 2), R271-R277. Konopka, R. J., & Benzer, S. (1971). Clock mutants of melanogaster. Proceedings of the National Academy of Sciences, 68(9), 2112-2116. Kramer, A., Yang, F. C., Snodgrass, P., Li, X., Scammell, T. E., Davis, F. C., & Weitz, C. J. (2001). Regulation of daily locomotor activity and sleep by hypothalamic EGF receptor signaling. , 294(5551), 2511-2515. Kriegsfeld, L. J., & Silver, R. (2006). The regulation of neuroendocrine function: timing is everything. Hormones and behavior, 49(5), 557-574. Kriegsfeld, L. J., Leak, R. K., Yackulic, C. B., LeSauter, J., & Silver, R. (2004). Organization of suprachiasmatic nucleus projections in Syrian hamsters (Mesocricetus auratus): an anterograde and retrograde analysis. The Journal of comparative neurology, 468(3), 361-379. Kriegsfeld, L. J., LeSauter, J., & Silver, R. (2004). Targeted microlesions reveal novel organization of the hamster suprachiasmatic nucleus. The Journal of neuroscience, 24(10), 2449-2457. 30

Kruijver, F. P., & Swaab, D. F. (2002). Sex hormone receptors are present in the human suprachiasmatic nucleus. Neuroendocrinology, 75(5), 296-305. Lehman, M. N., Silver, R., Gladstone, W. R., Kahn, R. M., Gibson, M., & Bittman, E. L. (1987). Circadian rhythmicity restored by neural transplant. Immunocytochemical characterization of the graft and its integration with the host brain. The Journal of neuroscience, 7(6), 1626-1638. LeSauter, J., & Silver, R. (1994). Suprachiasmatic nucleus lesions abolish and fetal grafts restore circadian gnawing rhythms in hamsters. Restorative neurology and neuroscience, 6(2), 135-143. LeSauter, J., & Silver, R. (1999). Localization of a suprachiasmatic nucleus subregion regulating locomotor rhythmicity. The Journal of neuroscience, 19(13), 5574-5585. Li, J. D., Hu, W. P., Boehmer, L., Cheng, M. Y., Lee, A. G., Jilek, A., ... & Zhou, Q. Y. (2006). Attenuated circadian rhythms in mice lacking the prokineticin 2 gene. The Journal of neuroscience, 26(45), 11615-11623. Meyer-Bernstein, E. L., Jetton, A. E., Matsumoto, S. I., Markuns, J. F., Lehman, M. N., & Bittman, E. L. (1999). Effects of suprachiasmatic transplants on circadian rhythms of neuroendocrine function in golden hamsters. Endocrinology, 140(1), 207-218. Michael, R. P., & Rees, H. D. (1982). Autoradiographic localization of 3H-dihyrotestosterone in the preoptic area, hypothalamus, and amygdala of a male rhesus monkey. Life sciences, 30(24), 2087-2093. Moore, R. Y., & Eichler, V. B. (1972). Loss of a circadian adrenal corticosterone rhythm following suprachiasmatic lesions in the rat. Brain Research; Brain Research. Moore, R. Y., & Lenn, N. J. (1972). A retinohypothalamic projection in the rat. The Journal of comparative neurology, 146(1), 1. Moore, R. Y., Speh, J. C., & Leak, R. K. (2002). Suprachiasmatic nucleus organization. Cell and tissue research, 309(1), 89-98. Morin, L. P. (1980). Effect of ovarian hormones on synchrony of hamster circadian rhythms. Physiology & behavior, 24(4), 741-749. 31

Morin, L. P., & Cummings, L. A. (1981). Effect of surgical or photoperiodic castration, testosterone replacement or pinealectomy on male hamster running rhythmicity. Physiology & behavior, 26(5), 825-838. Morin, L. P., & Cummings, L. A. (1982). Splitting of wheelrunning rhythms by castrated or steroid treated male and female hamsters. Physiology & Behavior, 29(4), 665-675. Morin, L. P., Fitzgerald, K. M., & Zucker, I. (1977). Estradiol shortens the period of hamster circadian rhythms. Science (New York, NY), 196(4287), 305. Nunez, A. A., & Stephan, F. K. (1977). The effects of hypothalamic knife cuts on drinking rhythms and the estrus cycle of the rat. Behavioral biology, 20(2), 224-234. Oliet, S. H., Piet, R., Poulain, D. A., & Theodosis, D. T. (2004). Glial modulation of synaptic transmission: Insights from the supraoptic nucleus of the hypothalamus. Glia, 47(3), 258-267. Olofsson, B., & de Bono, M. (2008). Sleep: Dozy worms and sleepy flies. Current Biology, 18(5), R204-R206. Pittendrigh, C. S. (1960). Circadian rhythms and the circadian organization of living systems. In Cold Spring Harbor Symposia on Quantitative Biology (Vol. 25, pp. 159-184). Cold Spring Harbor Laboratory Press. Ralph, M. R., Foster, R. G., Davis, F. C., & Menaker, M. (1990). Transplanted suprachiasmatic nucleus determines circadian period. Science, 247(4945), 975-978. Rees, H. D., & Michael, R. P. (1982). Brain cells of the male rhesus monkey accumulate 3H- testosterone or its metabolites. The Journal of comparative neurology, 206(3), 273-277. Roy, E. J., & Wade, G. N. (1975). Role of estrogens in androgen-induced spontaneous activity in male rats. Journal of comparative and physiological psychology, 89(6), 573. Ruggiero, L., & Silver, R. (2010) Circadian and Circannual Rhythms and Hormones. In Wingfield, J.C. ed. Encyclopedia of Animal Behavior. Elsevier. Shughrue, P. J., Komm, B., & Merchenthaler, I. (1996). The distribution of estrogen receptor-β mRNA in the rat hypothalamus. Steroids, 61(12), 678-681. 32

Shughrue, P., Scrimo, P., Lane, M., Askew, R., & Merchenthaler, I. (1997). The distribution of estrogen receptor-β mRNA in forebrain regions of the estrogen receptor-α knockout mouse. Endocrinology, 138(12), 5649-5652. Silver, R., Lehman, M. N., Gibson, M., Gladstone, W. R., & Bittman, E. L. (1990). Dispersed cell suspensions of fetal SCN restore circadian rhythmicity in SCN-lesioned adult hamsters. Brain research, 525(1), 45-58. Silver, R., LeSauter, J., Tresco, P. A., & Lehman, M. N. (1996). A diffusible coupling signal from the transplanted suprachiasmatic nucleus controlling circadian locomotor rhythms. Nature, 382(6594), 810-813. Stephan, F. K., & Zucker, I. (1972). Circadian rhythms in drinking behavior and locomotor activity of rats are eliminated by hypothalamic lesions. Proceedings of the National Academy of Sciences, 69(6), 1583-1586. Su, J. D., Qiu, J., Zhong, Y. P., & Chen, Y. Z. (2001). Expression of estrogen receptor-alpha and - beta immunoreactivity in the cultured neonatal suprachiasmatic nucleus: with special attention to GABAergic neurons. Neuroreport, 12(9), 1955-1959. Tanaka, M., Hayashi, S., Tamada, Y., Ikeda, T., Hisa, Y., Takamatsu, T., & Ibata, Y. (1997). Direct retinal projections to GRP neurons in the suprachiasmatic nucleus of the rat. Neuroreport, 8(9), 2187-2191. Van den Pol, A. N. (1980). The hypothalamic suprachiasmatic nucleus of rat: intrinsic anatomy. The Journal of comparative neurology, 191(4), 661. Van Der Beek, E. M., Horvath, T. L., Wiegant, V. M., Van Den Hurk, R., & Buijs, R. M. (1998). Evidence for a direct neuronal pathway from the suprachiasmatic nucleus to the gonadotropin-releasing hormone system: Combined tracing and light and microscopic immunocytochemical studies. The Journal of comparative neurology, 384(4), 569-579. Weaver, D. R. (1998). The suprachiasmatic nucleus: a 25-year retrospective. Journal of biological rhythms, 13(2), 100-112. Yan, L., & Okamura, H. (2002). Gradients in the circadian expression of Per1 and Per2 genes in the rat suprachiasmatic nucleus. European Journal of Neuroscience, 15(7), 1153-1162. 33

Table 1. Number of animals per condition for behavioural studies. 34

Table 2. Number of animals per condition for analysis of neural activation. 35

Table 3. Group Means and Standard Error of the Means (SEM) for number of Fos-ir cells. 36

Table 4. Group Means and Standard Error of the Means (SEM) for number of beam breaks. 37

Figure Captions Figure 1. Mean (+/- SEM) number of beam breaks recorded over 5 days. * Indicates statistical significance (p < 0.05). DHT = gonadectomized animals treated with dihydrotestosterone. INT = unmanipulated intact animals. GDX = gonadectomized animals without androgen replacement. Figure 2. Mean (+/- SEM) number of Fos-immunoreactive cells in bilateral paired SCN. * Indicates statistical significance (p < 0.05). DHT = gonadectomized animals treated with dihydrotestosterone. INT = unmanipulated intact animals. GDX = gonadectomized animals without androgen replacement. Figure 3. Comparison of AR immunoreactive cells in wildtype and Nestin-AR littermates captured at 200x magnification. 38

Figure 1. Total number of beam breaks recorded over 5 days. 39

Figure 2. Number of Fos-ir cells in bilateral SCN. 40

Figure 3. Androgen receptor immunoreactive cells of Nestin-AR mutants and wildtype littermates.

Left SCN Right SCN

Nestin-AR

Wildtype