. : 2 , 32.x. .. ., gamflfa. ..MMWW... . . 33. I . n. .3: .1. 3); «MM . i: .ilvu...“ ubfihé. man. “rt-luau... a .. h hfidilr :(t.....ul: nun: $47.15 23:33.3? Elihu mars LIBRARY I Michigan State 9 University $191507» This is to certify that the dissertation entitled SEX, SURGES, AND CIRCADIAN RHYTHMS: THE TIMING OF REPRODUCTIVE EVENTS IN A DIURNAL RODENT presented by Megan M. Mahoney has been accepted towards fulfillment of the requirements for the PH.D degree in ZOOLOGY AND ECOLOGY, EVOLUTIONARY BIOLOGY AND BEHAVIOR PROGRAM $4M Major Professor’s Signature “ff/a: Date MSU is an Affirmative Action/Equal Opportunity Institution PLACE IN RETURN BOX to remove this checkout from your record. TO AVOID FINES return on or before date due. MAY BE RECALLED with earlier due date if requested. DATE DUE DATE DUE DATE DUE 6/01 cJClRC/DateDuosz-pJS SEX, SURGES AND CIRCADIAN RHYTHMS: THE TIMING OF REPRODUCTIVE EVENTS IN A DIURNAL RODENT By Megan M. Mahoney A DISSERTATION Submitted to Michigan State University in partial fulfillment of the requirements for the degree of DOCTOR OF PHILOSOPHY Department of Zoology and Ecology, Evolutionary Biology and Behavior Program 2003 ABSTRACT SEX, SURGES AND CIRCADIAN RHYTHMS: THE TIMING OF REPRODUCTIVE EVENTS IN A DIURNAL RODENT By Megan M. Mahoney Rhythms in the timing of reproductive events are reversed in diurnal and nocturnal rodents, but little is known about the neural mechanisms underlying these differences. I examined these issues by comparing the diurnal murid rodent Arvicanthis niloticus (grass rat) to nocturnal Rattus norvegicus (lab rat). In the first set of experiments, I examined the hypothesis that differences in the timing of estrous events in diurnal and nocturnal species are due to differences in rhythms in responsiveness to steroid hormones. I found that steroids were able to induce a rise in activity of neurons containing gonadotropin releasing hormone (GnRH) at only one time of day, which was 12 hours apart in grass rats and lab rats. These temporal patterns persisted in both species when they were housed in constant darkness for five days suggesting that an endogenous circadian clock drives the rhythms in responsiveness to hormones. Secondly, I determined whether cells within the suprachiasmatic nucleus (SCN), the site of the principle mammalian circadian clock, project to neuroendocrine and steroid sensitive cells in grass rats, and whether these pathways differed from those of lab rats. Anterograde tract-tracing in grass rats revealed that both GnRH and estrogen receptor (ER) containing cells appear to receive input from the SCN, as has been seen in nocturnal rodents. I then found that arginine vasopressin and vasoactive intestinal polypeptide, two neuropeptides made in the SCN, were contained in axon terminals that contacted GnRH neurons, as is also the case in lab rats and hamsters. In the third set of experiments, I examined the hypothesis that inverted rhythms in the timing of estrus-related behaviors in diurnal and nocturnal rodents are due to differences in rhythms in sensitivity to steroid hormones. Pairs of males and hormone-primed females were tested for mating at four different times of day. I found that grass rats had a daily rhythm in sexual behavior that was 12 hours out of phase relative to that seen in nocturnal rodents. Specifically, both the lordosis quotient and rate of copulation were relatively low at zeitgeber time (ZT) 17 and then rose to a peak at ZT 23. I also observed a bimodal rhythm in male mounting behavior that peaked at ZT 11 and 23. Taken together, these results indicate that steroid-primed grass rats and lab rats are similar with respect to the temporal relationship among estrous-related events, but that the timing of these events relative to the lightzdark cycle is dramatically different. These differences appear to be due to rhythms in responsiveness to steroid hormones whereas the structure of the neural pathways communicating temporal information from the SCN to cells within the reproductive axis appear to be the same. The mechanisms underlying the differences in rhythms of sensitivity to steroids might involve temporal patterns of signals emitted by the SCN and/or patterns of sensitivity of neural targets to these signals. _ _ ___._,____7‘_ ._fl __ _. For Bill iv Acknowledgements First and foremost I would like to thank my advisor, Dr. Laura Smale. She has been generous, patient, kind and helpful to me, both professionally and personally. I would also like to thank my committee members Drs. Antonio Nunez, Lynwood Clemens and Kay Holekamp for their readiness to talk, write letters, respond to e-mails, and answer questions whenever I needed them. I have also had a lot of support from current and past lab mates Michael Schwartz, Joshua Nixon, Gladys Martinez, Russell Van Horn, Julie Blanchong, Teresa McElhinny, and Dr. Colleen Novak. Many other graduate student and post-doc friends have helped me including Cynthia Wei, Kalynn Schulz, Kaliris Salas- Ramirez, and Drs. Russell Romeo, Heather Richardson, Julia Zehr, Yu-Ping Tang, Sean Veney, Matthew Lovern, and David Parfitt. l have had the good fortune to be assisted by many talented undergraduate students over the years including Heather Ross, Betty Gubik, Joel Breen, Julie Harris, Sandra Rose, Cassie Castlebury, Jennifer Adams, Paden Ross, Nicole Timm, and Matthew Heintz as well as laboratory technicians Janaina Gamez and Anna Baumgras. Jane Venier has advised me often over the years. She also performed the radioimmunoassays in Dr. Cheryl Sisk’s lab that are described in Chapter 2. I would also like to thank Drs. Sisk and Juli Wade for their advice and contributions towards my career development. I have also had the good fortune to work with Sonya Lawrence, Tony D’Angelo, and Drs. Don Hall and Dick Hill. Lastly, I received financial assistance in the form of research assistantships from NIMH ROI-MHO53433 to Dr. Smale. and grants from Sigma Xi Grants-in-Aid of Research Program, Graduate Women in Science Vessa Notchev Fellowship, NIMH-Society for Behavioral Neuroendocrinology, and Society for Research on Biological Rhythms. The following units at Michigan State University also provided me with funds: the Graduate School, College of Natural Science, Department of Zoology, and Program in Ecology Evolutionary Biology and Behavior. vi TABLE OF CONTENTS LIST OF TABLES ................................................................................................. ix LIST OF FIGURES ................................................................................................ x Key to abbreviations: .......................................................................................... xiii Chapter 1 General Introduction .............................................................................................. 1 The LH surge and the circadian system ............................................................ 5 Arvicanthis niloticus: a diurnal murid rodent ...................................................... 6 Overview of chapters ......................................................................................... 7 Chapter 2 Circadian regulation of gonadotropin-releasing hormone neurons and the preovulatory surge in luteinizing hormone in diurnal and nocturnal rodents .......... 9 Introduction ........................................................................................................ 9 Materials and methods: ................................................................................... 12 Animals ........................................................................................................ 12 lmmunocytochemistry .................................................................................. 13 Patterns of change in LH and in GnRH+ cells in grass rats ......................... 14 GnRH+ and Fos+ in grass rats and lab rats in a lightzdark cycle ................. 15 GnRH+ and Fos in grass rats and lab rats in constant darkness ................. 16 Statistical analysis ........................................................................................ 16 Results ............................................................................................................ 17 Patterns of change in LH and in GnRH+ cells in grass rats ......................... 17 GnRH+ and Fos+ in grass rats and lab rats in a lightzdark cycle ................. 20 GnRH+ and Fos+ in grass rats and lab rats in constant darkness ............... 22 Discussion ....................................................................................................... 22 Chapter 3 Projections from the suprachiasmatic nucleus of grass rats to gonadotropin releasing hormone and estrogen receptor containing cells ................................. 28 Introduction ...................................................................................................... 28 Materials and methods .................................................................................... 34 Animals ........................................................................................................ 34 Surgeries ...................................................................................................... 34 Tissue processing and analysis ................................................................... 35 Results ............................................................................................................ 43 Single labeled BDA ...................................................................................... 43 ER+ cell distribution, BDA fibers and ER+ cells ........................................... 48 BDA fibers and GnRH+ cells ........................................................................ 55 vii AVP+ and VIP+ fiber distribution .................................................................. 55 AVP+ and VIP+ contacts on GnRH+ neurons .............................................. 58 Discussion ....................................................................................................... 64 Chapter 4 A daily rhythm in mating behavior and progestin receptor expression in a diurnal murid rodent Arvicanthis niloticus ........................................................................ 69 Introduction ...................................................................................................... 69 Materials and methods: ................................................................................... 71 Sexual behavior ........................................................................................... 72 Progesterone receptors ............................................................................... 74 Results ............................................................................................................ 78 Sexual behavior ........................................................................................... 78 Progesterone receptors .................................................................... 82 Discussion ....................................................................................................... 84 Chapter 5 Conclusion .......................................................................................................... 91 Chapter summary ............................................................................................ 91 What do these data mean? Where do I go from here? .................................... 92 References .......................................................................................................... 94 viii LIST OF TABLES Table 3.1. Antibodies and sera used for immunocytochemistry. Note that BDA was already biotinylated and thus did not require antibodies. ............................. 38 Table 3.2. Definitions for abbreviations used in this report ................................. 39 Table 3.3. Percentage of GnRH+ cells contacted by BDA labeled fibers. GnRH+ cells were analyzed in tissue from 3 individuals with iontophoretic BDA injections in the SCN. Abbreviations can be found in Table 3.2 ......................................... 57 Table 3.4. Percent of GnRH+ neurons contacted by AVP+ and VIP+ fibers in the hypothalamus of grass rats (mean +SEM) .......................................................... 61 Table 3.5. Summary of data indicating the location and density of BDA labeled fibers, VIP+ and AVP+ fibers, and ER-I- and GnRH+ neurons in grass rats. Scale of density ranges from least dense (+) to most dense (++++). A dashed line indicates that labeling was not detected. Fiber and cell densities reflect the range for a given type of labeling and are not directly comparable across substances. See Table 3.2 for abbreviations. .................................................... 63 LIST OF FIGURES Images in this dissertation are presented in color. Figure 1.1. A schematic diagram of the hypothalamic-pituitary-gonadal axis. Immediately prior to the LH surge estradiol secreted from the ovaries acts as a signal that stimulates GnRH release into the portal blood system. From there, GnRH reaches the anterior pituitary and induces secretion of LH. ....................... 2 Figure 2.1. The average percent (:SEM) of GnRH cells that contained Fos in grass rats. Hormone injections occurred at ZT 19 (arrow). Dots represent the value for each individual (n=5/time point). Zeitgeber time 0 = lights-on. ............. 18 Figure 2.2. The average LH concentration (:SEM) in plasma of grass rats sacrificed at various times of day. See legend of Figure 2.1 for detail. .............. 19 Figure 2.3. GnRH cell activity in steroid primed grass rats and lab rats housed in a 12:12 lightzdark cycle. Zeitgeber time 0 = lights-on. ........................................ 21 Figure 2.4. Percent of GnRH cells that were active in steroid primed grass rats and lab rats housed in constant darkness for 5 days before perfusion. .............. 23 Figure 3.1. A-C) Photomicrographs depicting IDA injection sites in the SCN of 3 individual grass rats. The boxed area in A is enlarged in D. D) Note the dense staining within the injection site and that labeled fibers become more visible at the borders of the site. E) An example of BDA labeled fibers in the LS taken at 200x magnification. F) An example of retrogradely labeled cells in the LS. Arrows indicate retrogradely labeled cells. Abbreviations are found in Table 3.2 .......................................................................................................................... .44 Figure 3.2. A series of line drawings based on 3 animals with BDA injections into the SCN. Black lines indicate BDA labeled fibers. In all pictures, the injection site was on the left side. Note that labeling was heavier on the side ipsilateral to the injection site. Abbreviations are noted in Table 3.2. ..................................... 45 Figure 3.3. Photomicrographs depicting representative BDA labeled fibers in animals with an injection of BDA within the SCN. The dashed line in picture C indicates the ventral edge of the tissue. Dashed lines in B, E, and F outline structures. See Table 3.2 for abbreviations ........................................................ 46 X Figure 3.4. Photomicrographs depicting A) ER+ cells in the AVPV of an estradiol primed grass rat. B) Low power view of patterns of overlap between BDA labeled fibers (blue) and ER+ cells (brown) in the PeVN (left side of image). Note that as ER+ staining decreases so does the density of BDA staining. Abbreviations are found in Table 3.2 ................................................................... 49 Figure 3.5. A series of line drawings at 5 levels from rostral (1) to caudal (5) depicting the distribution of ER+ and GnRH+ cells and AVP+ and VIP+ fibers in the hypothalamus of grass rats. In figures of ER+ and GnRH+ each black dot represents 1 cell. Pictures of GnRH+ cell distribution are modified from McElhinny et al. (1999). See Table 3.2 for abbreviations. .................................. 50 Figure 3.6. Photomicrographs depicting BDA fibers (blue staining) overlapping with ER+ cells (brown staining). Arrows indicate places where BDA fibers are in close apposition with ER+ cells. .......................................................................... 53 Figure 3.7. A, B) Photomicrographs depicting the pattern of overlap between BDA labeled fibers (blue) and GnRH-I- fibers (brown) in female grass rats. C-G) BDA labeled fibers contacting GnRH+ cells. Arrows indicate putative contacts. ............................................................................................................................ 56 Figure 3.8. Photomicrographs depicting AVP+ fibers (blue) contacting GnRH+ cells (brown) in the hypothalamus of grass rats. Pictures in C and D are the same image taken at different planes of focus. Arrows indicate putative contacts. .............................................................................................................. 59 Figure 3.9. Photomicrographs depicting VIP+ fibers (blue) contacting GnRH+ cells (brown) in a steroid primed female grass rat. Arrows indicate putative contacts. .............................................................................................................. 60 Figure 4.1. An example of PR+ cells in the ventrolateral hypothalamus in an estradiol-treated female grass rat. The box (200 x 400 pm) in the figure indicates the region in which PR+ cells were counted. ARC = arcuate nucleus. ............... 77 Figure 4.2. Average rates of sexual behaviors (1 SEM) in 15 pairs of grass rats tested at 4 different times of day. Bars with different letters over them are significantly different from one another, p<0.05. Zeitgeber time O=lights-on. ..... 79 xi Figure 4.3. Lordosis quotient (number of lordosis responses/number of mounting attempts) for 15 individual grass rats. Animals were grouped into two types of sexual responsiveness on the basis of their behavior at ZT 17. Animals in the “low” group exhibited an LQ below 50% at ZT 17 (grey dots or bars); remaining individuals were placed in the “high” group (black dots or bars). Zeitgeber time 0=lights-on. A) LQ as a function of time for each individual. B) Average lordosis quotient (_+_ SEM) for the two groups of responders. ............................................ 80 Figure 4.4. Average number (_+_ SEM) of PR+ cells in the ventrolateral hypothalamus in female grass rats. Estradiol benzoate treated females received injections on days 1 and 2 and were sacrificed at day 3, at the same time. Control females did not receive any treatment. Bars with different letters over them are significantly different from one another, p<0.05. ZT O = lights-on ....... 83 Figure 4.5. Average number (:SEM) of PR+ cells in the ventrolateral hypothalamus in female grass rats. Intact females were implanted with EB- containing silastic capsules. Bars with different letters over them are significantly different from one another, p<0.05. ZT 0 = lights-on. ......................................... 85 xii Key to abbreviations: Abbreviation Definition 3V 3rd ventricle ac anterior commisure AHA anterior hypothalamic area ANOVA Analysis of variance ARC arcuate nucleus AVP arginine vasopressin AVPV anteroventralgortion of the periventricular nucleus BDA biotinylated dextran amine BNST bed nucleus of the stria terminalis D3V dorsal portion of 3rd ventricle DAB diaminobenzidine 088 diagonal band of Broca DD constant darkness EB estradiol benzoate ER estrogen receptor-oz f fornix GnRH gonadotropin releasing hormone hDBB horizontal portion of 088 HPG hypothalamic-pituitary-gonadal icv intracerebroventriculag im intramuscularly LH luteinizing hormone LPOA lateral portion of POA LQ lordosis quotient LS lateral septum LSPV lower sub paraventricular zone LSV ventral portion of LS LV lateral ventricle mPOA medial portion of POA MS medial septum NGS normal goat serum NHS normal horse serum oc optic chiasm ot optic tract OVLT organum vasculosum of the lamina terminalis P progesterone PBS phosphate buffered saline PeVN periventricular nucleus POA preoptic area PR progestin receptors PVN paraventricular nucleus of the hypothalamus xiii Key to abbreviations: Abbreviation Definition PVT paraventricular thalamus RCH retrochiasmatic area so subcutaneous SCN suprachiasmatic nucleus SEM standard error of the mean SON supraoptic nucleus TX triton-X vDBB vertical portion of 088 VIP vasoactive intestinal polypeptide VLH ventrolateral hypothalamus VMH ventromedial hypothalamus VMPO ventromedial portion of POA ZT zeitgeber time xiv Chapter 1 General Introduction The research described in this dissertation is aimed at investigating the neuronal mechanisms that regulate the timing of reproductive events in a diurnal rodent, Arvicanthis niloticus. Specifically, I have focused on the timing of mating and of the ovulatory surge of luteinizing hormone (LH). In this introductory chapter I will provide 1) an overview of how the hypothalamic-pituitary-gonadal (HPG) axis functions to regulate the secretion of LH, 2) how the circadian timing system may interact with the HPG system to regulate the timing of reproductive events in female rodents, 3) a rationale for why I use a diurnal rodent model, and 4) the questions and experiments that will be addressed in each chapter of my dissertation. The hypothalamic-pituitary-gonadal axis and the LH surge Critical reproductive events associated with the mammalian estrous cycle such as mating, ovulation and preovulatory changes in hormone secretion depend upon the precise temporal coordination of functions occurring at different levels of the HPG axis (Figure 1.1). This integration requires: 1) a steroid- hormone sensitive system which responds to the increasing levels of estradiol released from the maturing follicles, 2) Gonadotropin releasing hormone (GnRH) neurons that stimulate the release of LH from the anterior pituitary, and 3) a precise timing mechanism which can coordinate the release of GnRH from a diffuse population of cells (van der Beek, 1996). +/- f hypothalamus Anterior pituitary Figure 1.1. A schematic diagram of the hypothalamic-pituitary—gonadal axis. Immediately prior to the LH surge estradiol secreted from the ovaries acts as a signal that stimulates GnRH release into the portal blood system. From there, GnRH reaches the anterior pituitary and induces secretion of LH. 2 Much of our knowledge regarding the neuroendocrine changes associated with the estrous cycle is based on studies of laboratory rodents including Rattus norvegicus (“lab rats”) and Mesocricetus auratus (golden hamsters). During the follicular phase of the estrous cycle, developing ovarian follicles release estradiol, which inhibits gonadotropin secretion through a negative feedback loop. At proestrous, estradiol levels eventually peak and this ovarian signal switches from an inhibitory to stimulatory factor. Estradiol acts on the hypothalamus, causing GnRH to be released into the portal circulatory system through which it reaches the anterior pituitary (reviewed in Freeman, 1994). GnRH then acts on gonadotropes in the pituitary to trigger LH release. Thus, during proestrous the surge in LH is driven by a surge in GnRH released from neurons within the hypothalamus. Further evidence from lab rats and hamsters indicates that the increase in steroid secretion from the ovaries induces a rise in GnRH mRNA (Levine and Ramirez, 1982; Petersen et al., 1995; Porkka-Helskanen et al., 1994; Wang et al., 1995) and a rise in the number of GnRH cells that contain Fos (Doan and Urbanski, 1994; Lee et al., 1992; Lee et al., 1990a; Lee et al., 1990b). Fos, a protein product of the c-fos gene, has been found to be a reliable indicator of cellular activity (Hoffman et al., 1994; Hoffman et al., 1990; Wang et al., 1995). Ovariectomized (OVXed) lab rats do not have a spontaneous LH surge due to the removal of their endogenous estrogens. However, administration (either by injections or capsules) of Estradiol (E) or a combination of E and progesterone in laboratory rodents (e.g. lab rats, mice) can stimulate an increase in GnRH, GnRH mRNA, GnRH neurons containing Fos, and LH secretion that resemble normal proestrous events (Bronson and Vom Saal, 1979; Jimenez-Linan and Rubin, 2001 ; Lee et al., 1990b; Legan et al., 1975; Levine and Ramirez, 1982; Wu et al., 1992). While coordination between estrogens and GnRH neurons is required to trigger the LH surge, a large body of evidence suggests that GnRH neurons lack estrogen receptor-a (ER-a; Herbison and Theodosis, 1992; Lehman and Karsch, 1993; Shivers et al., 1983; Warembourg et al., 1998). These data, from a variety of species, suggest that these cells do not receive information on ovarian status directly from circulating estrogens. Retrograde tracing in lab rats has revealed that ER-a positive neurons in the hypothalamus and caudal brainstem project to the vicinity of GnRH cells in the preoptic area, suggesting a possible anatomical substrate for integration of steroid signals with the GnRH cell system (Simonian etaL,1999) Recently, however, controversial reports have identified ER-a in both an immortalized GnRH cell line and in GnRH cells present in tissue slices of the lab rat preoptic area (Butler et al., 1999; Roy et al., 1999; Shen et al., 1998; Skynner et al., 1999). Additionally, ER-B, a second estrogen receptor isoform has recently been identified, and its mRNA and protein have been found in GnRH cells in vivo and in vitro (Hrabovszky et al., 2000; Roy et al., 1999; Skynner et al., 1999). Although these data are intriguing, further investigations are needed to determine whether these steroid receptors in GnRH neurons play a role in mediating the LH surge. The LH surge and the circadian system In many rodents, the LH surge is regulated by the endogenous circadian system. The rhythm regulating the LH surge free-runs in constant conditions (Alleva et al., 1971; Takeo, 1984), is phase-locked to the lightzdark cycle (Fitzgerald and Zucker, 1976; Stetson and Gibson, 1977), and is delayed a full circadian cycle following a single injection of barbiturates (Everett and Sawyer, 1950). OVXed lab rats implanted with estradiol capsules have a daily proestrous-like surge, indicating that there is a daily signal for the initiation of LH release. This circadian signal appears to operate in conjunction with the high levels of circulating estradiol that are reached at proestrous to stimulate the LH surge once every 4 or 5 days in intact lab rats. The SCN, the site of the primary circadian clock in mammals, plays an important role in the temporal coordination of these events in nocturnal laboratory rodents. Lab rats and hamsters with bilateral SCN lesions lack a preovulatory LH surge and a consistently functional estrous cycle, and do not exhibit an estradiol induced LH surge (Gray et al., 1978; Kawakami et al., 1980; Meyer-Bernstein et al., 1999; Weigand and Terasawa, 1982). Fetal SCN transplants do not restore these endocrine rhythms, indicating that synaptic inputs are critical for mediating these estrous-related events (Meyer-Bernstein et al., 1999). The SCN may communicate temporal information though direct neural connections to cells within the HPG system. Light and electron microscopy reveal SCN efferents on GnRH and ER-a containing cells in lab rats and hamsters (de la lglesia et al., 1995; van der Beek et al., 1997; Watson et al., 1995). Furthermore, these two cell types also project back to cells contained within SCN and the region surrounding this nucleus (de la lglesia et al., 1999; van der Beek, Wiegant et al., 1997). These data provide evidence for neural pathways that connect the circadian system, GnRH neurons, and ER-containing cells. Arvicanthis niloticus: a diurnal murid rodent The rhythms regulating the timing of estrous related events are reversed in diurnal and nocturnal rodents. In lab rats, the preovulatory LH surge, GnRH cell activation, and the onset of sexual receptivity occur at the beginning of the active period, which is around the time of lights-off (reviewed in Silver and Bittman, 1984). In several diurnal rodents including ground squirrels and degus, mating behavior occurs at the beginning and throughout their active period, but in this case it occurs during the light phase of the lightzdark cycle (Dobson and Michener, 1995; Michener, 1980; Labyak and Lee, 1995; Rossi and Lee personal communication). It is not clear how the neural mechanisms that regulate the timing of these events differ in diurnal and nocturnal species. This issue has been difficult to address in squirrels and degus for a variety of reasons. The research in this thesis focuses on a diurnal rodent, the unstriped Nile grass rat, Arvicanthis niloticus (“grass rat”). This murid species is closely related to lab rats and hamsters, nocturnal animals often used for endocrine and chronobiology studies. Grass rats are diurnal with respect to patterns in general activity, body temperature, wheel-running, and activity in the field (Blanchong and Smale, 2000; McElhinny et al., 1997). This species also exhibits a reversal in the rhythms controlling mating, the LH surge during the postpartum estrus period (PPE), and GnRH cell activation during the PPE when compared to those of lab rats (McElhinny et al., 1999). Much of the neuroanatomy of the SCN, as well as the GnRH cell distribution has been described in this species (Katona et al., 1998; Mahoney et al., 2001; McElhinny et al., 1999; Smale and Boverhof, 1999). For these various reasons, the grass rat represents an ideal model with which to elucidate the regulation and timing of reproductive events in a diurnal mammal. Overview of chapters The goals of this dissertation research were to: 1) elucidate possible mechanisms underlying rhythms in the timing of reproductive events in Arvicanthis niloticus and 2) determine whether, and how, these mechanisms differ in grass rats compared to nocturnal rodent species. Specifically, in the work described in chapter 2 I sought to determine whether there is a daily pattern of change in responsiveness to steroid hormones in the diurnal grass rat that is reversed when compared to that of nocturnal rodents. To examine this issue, | first determined that OVXed grass rats have a surge of LH and an associated increase in GnRH cell activation following appropriate steroid treatment. Using immunocytochemistry I then examined whether grass rats and lab rats have a rhythm in the timing of GnRH cell activation, whether it is reversed in these species, and whether the rhythm is endogenous. The experiments described in chapter 3 l characterized the pathways from the SCN to GnRH and ER-o cells in grass rats and compared them to those described earlier in nocturnal rodents. Anterograde tract-tracing was used to identify SCN efferents innervating these two cell types. I then used immunocytochemistry to determine whether arginine vasopressin (AVP) and vasoactive intestinal polypeptide (VIP), two peptides found in efferents of the SCN, contact GnRH cells in grass rats. A large body of evidence indicates that these peptides regulate the timing of the LH surge and sexual receptivity in female lab rats. I used immunocytochemistry to determine whether connections between GnRH neurons and VIP and/or AVP fibers differ between grass rats and Mbmm. In chapter 4 I sought to determine whether there is a rhythm in responsiveness of sexual behavior to steroid hormones in grass rats, and whether the rhythm is reversed compared to that seen in nocturnal rodents. To address this issue I analyzed the rates of sexual behaviors in paired grass rats tested at 4 different times of day. I then used immunocytochemistry to explore the possibility that the rhythm in behavioral responsiveness to steroids is associated with a rhythm in of progesterone receptors. Chapter 2: Circadian regulation of gonadotropin-releasing hormone neurons and the preovulatory surge in luteinizing hormone in diurnal and nocturnal rodents Introduction Reproductive events such as copulatory behavior, parturition, ovarian cyclicity, and the preovulatory surge in luteinizing hormone (LH) can occur at very different times of day in nocturnal and diurnal animals. Nocturnal female rats ("lab rats", Rattus norvegicus), mice (Mus musculus) and Syrian hamsters (Mesocricetus auratus) mate and have the LH surge in the late afternoon or early evening, around lights-off (Blake, 1976; Bronson and Vom Saal, 1979; Legan and Karsch, 1975; Seegal and Goldman, 1975; Sodersten, 1988; Stetson and Gibson, 1977; Wu et al., 1992), whereas in the diurnal rodent, Arvicanthis niloticus ("grass rat"), sexual behavior and the LH surge occur very early in the morning, before lights-on (McElhinny et al., 1999; McElhinny et al., 1997). In addition grass rats and lab rats exhibit a rise in activity in gonadotropin releasing hormone (GnRH) cells at opposite times of day (Lee et al., 1992; McElhinny et al., 1999; Wang et al., 1995). The release of GnRH from neurons in the hypothalamus induces LH secretion from the anterior pituitary. Most chronobiology research focuses on nocturnal lab rodents and it is not known what causes these estrus-related events to occur at different times of day in diurnal and nocturnal species. Several lines of evidence have established that endogenous signals from the circadian system play a critical role in the coordination of reproductive rhythms in lab rats and hamsters. If the surge is suppressed by barbiturate treatment on the day of proestrous it does not occur immediately after recovery from the treatment, but is instead delayed a full circadian cycle (Everett and Sawyer, 1950; Stetson and Watson-Whitmyre, 1977). Furthermore, when these animals are housed in constant light they have free-running rhythms in the timing of the LH surge and in the onset of sexual receptivity that are "circa-quadridian". That is, these events occur at intervals that are four times as long as the period of the circadian rhythms (Alleva et al., 1971; Fitzgerald and Zucker, 1976; Takeo, 1984). In lab rats and hamsters kept in a lightzdark cycle both the LH surge and the onset of lordosis behavior are tightly coupled to the onset of activity (Alleva et al., 1971; Moline et al., 1981; Stetson and Gibson, 1977). When activity rhythms are phase shifted by pharmacological treatment, or when “splitting” of activity bouts occurs, the timing of the LH surge also changes such that the same temporal relationship is maintained between these functions (Fitzgerald and Zucker, 1976; Swann and Turek, 1982; Swann and Turek, 1985). A daily signal for this surge is also evident in ovariectomized lab rats exposed to constant levels of estradiol in which a preovulatory-like surge occurs at the same time each day (Legan et al., 1975; Legan and Karsch, 1975). The circadian signal that gates the timing of estrous-related events originates in the suprachiasmatic nucleus (SCN), the site of the primary circadian clock in mammals. Destruction of the SCN abolishes the preovulatory LH surge, 10 a consistently functional estrous cycle, and a behavioral rhythm in responsiveness to steroid hormones (Gray et al., 1978; Kawakami et al., 1980; Meyer-Bernstein et al., 1999; Weigand and Terasawa, 1982). There is a direct pathway from the SCN to GnRH neurons in lab rats and hamsters, and in lab rats these contacts are clearly synaptic (de la lglesia et al., 1995; van der Beek et al., 1997). It is not clear whether the SCN and circadian system are involved in the timing of estrous-related events in diurnal rodents as, until recently, there has not been a suitable model with which to investigate this question. The unstriped Nile grass rat, Arvicanthis niloticus, has proven to be ideal for the elucidation of questions regarding the neural control of biological rhythms in diurnal mammals. This murid rodent exhibits a diurnal pattern of mating activity, body temperature, and above ground activity in the field (Blanchong and Smale, 2000; McElhinny et al., 1997). These animals also have an LH surge during the post-partum estrus period that occurs nearly 12 hours out of phase with that of lab rats (McElhinny et al., 1999). Previous studies of grass rats have focused on the postpartum period because estrus at this time can be predicted with a high degree of certainty, which is not the case at other times (e.g. vaginal smears do not predict estrus). The goals of this research were to investigate the neuroendocrine events associated with the LH surge in a diurnal rodent, and the timing of these events in nocturnal and diurnal rodents. More specifically I had three objectives: 1) to determine whether a surge could be induced in ovariectomized females primed with steroid hormones, 2) to evaluate the hypothesis that lab rats and grass rats 11 exhibit reversed temporal patterns of GnRH cell activity when given identical steroid hormone treatment and 3) to evaluate the hypothesis that the rhythms in GnRH cell activity are endogenous in these species. For the purposes of this paper, GnRH cell activity is used to refer to levels of Fos expression in GnRH neurons. The recruitment of GnRH neurons during the preovulatory LH surge of lab rats, guinea pigs, and hamsters has been clearly associated with the detection of the immediate early gene product Fos within these neuroendocrine cells (Doan and Urbanski, 1994; Hoffman et al., 1990; King et al., 1998; Lee et al., 1992; Lee etal., 1990a; Wang etal., 1995; Wu etal., 1992). Materials and methods: Animals Adult female grass rats (>60 days) bred from laboratory stock and Sprague Dawley rats (Charles River) were housed in a 12:12 lightzdark cycle and provided food (Teklad rodent chow 8640, Harlan Industries) and water ad libitum. A red light (<5 qu) was left on continuously in the animal rooms. Females were anesthetized with sodium pentobarbital (Nembutal, Abbott Laboratories, <50 mg/kg sodium pentobarbital) and methoxyflurane (Metofane, Mallinckrodt Veterinary) and bilaterally ovariectomized. lncisions were closed with sutures (grass rats) or wound clips (lab rats) and treated with topical antibiotic (Nolvasan, Fort Dodge Animal Health). Following ovariectomy animals were given 1 cc 0.9% saline subcutaneously (so) and 0.03 mg buprenorphine hydrochloride (intramuscularly; im, Buprenex, Reckitt & Coleman). I waited 7-14 days after ovariectomy before beginning each experiment. All experiments were performed 12 in compliance with Michigan State University All-University Committee on Animal Use and in accordance with the standard in the National Research Council Guide for Care and Use of Laboratory Animals. All efforts were made to minimize the suffering and the number of animals used in these experiments. lmmunocytochemistry All animals were deeply anaesthetized with sodium pentobarbital and perfused transcardially with 0.01 M PBS (pH 7.2, 150-200 ml/animal) followed by 4% paraformaldehyde (Sigma) in 0.1 M phosphate buffer. Brains were post-fixed in paraformaldehyde for 4 hours, transferred to 20% sucrose in 0.1 M phosphate buffer overnight, then sectioned using a freezing microtome at 30 um into 3 series, from the medial septum to the bed nucleus of the stria terminalis. Brains from animals in the first study, however, were cut into 2 series. One series of brain sections from each animal was processed for the dual detection of GnRH and Fos immunoreactivity (+). Free floating tissue sections were incubated in (1) 5% normal goat serum (NGS, Vector Laboratories; in PBS with 0.3% triton-X; TX) for one hour at room temperature, then (2) rabbit anti-Fos primary antibody for 24 hours at 4°C (Santa Cruz Biotechnologies) in PBS with 0.3% TX and 3% NGS), followed by (3) biotinylated secondary antibody for one hour at room temperature (1:200, goat anti-rabbit in PBS with 0.3% TX and 3% NGS, Vector Laboratories) then (4) Avidin-biotin complex (ABC, Vectastain Elite Kit, Vector Laboratories) for one hour at room temperature. Fos was then visualized by incubating tissue in 0.175 M sodium acetate buffer containing diaminobenzidine (DAB, 0.5 mg/ml), 3% hydrogen peroxide (0.825 ul/ml buffer) 13 and 2.5% nickel sulfate. In between each step, tissue was rinsed three times for 10 minutes in PBS. After Fos labeling, sections were processed for the detection of GnRH. The same immunocytochemistry procedure was used with the following exceptions: I used rabbit anti-GnRH primary antibody (1 :5000, Chemicon lntemational), normal donkey serum, and donkey anti-rabbit F(ab)2 secondary antibody (Jackson lmmunoResearch Laboratories). Tissue was reacted in DAB (0.5 mg/ml, in Trizma buffer, pH 7.2) with 30% hydrogen peroxide (0.35 ul/ml buffer). Controls were done by repeating this dual label procedure but either Fos, or GnRH, or both antibodies were omitted. Following the immunocytochemical reactions, tissue was mounted, dehydrated, coverslipped and examined under a light microscope (Labortux S, Leitz Wetzlar GBH). For each study, the numbers of GnRH+ cells with and without Fos+ were quantified. Patterns of change in LH and in GnRH+ cells in grass rats In this first study ovariectomized grass rats were primed with steroid hormones in order to induce a surge of LH. For two days, animals were injected (sc) with 10 pg 17-(3 estradiol benzoate suspended in sesame oil (EB) at zeitgeber time 19 (ZT; ZT 0 = lights on). On the third day, at the same ZT, females received 125 pg progesterone (P; sc). Following the P injection females were perfused at either ZT 20.5, 22, or 23.5 (n=5/timepoint). Another group of females was perfused at ZT 18.5, without receiving a P injection (n=5). At the time of perfusion, brains were collected, cardiac blood samples were taken and centrifuged, and the plasma was stored at -80°F. Brains were processed and the 14 numbers of GnRH+ cells with and without Fos+ were counted from 14 sections taken from each animal. Six of these contained the medial septum and diagonal and horizontal bands of Broca (MS/DBB), two contained the organum vasculosum of the lamina terminalis (OVLT) and six contained the anteroventral portion of the periventricular nucleus and preoptic area (AVPV/POA). Plasma LH concentrations were measured by a double-antibody radioimmunoassay, which had previously been validated for use with grass rat plasma (McElhinny et al., 1999). The primary antibody, mouse monoclonal anti- bovine LH (518B7, lot 4), was provided by Dr. Jan Roser (University of California, Davis). Dr. L. Reichert (Albany Medical College, NY) provided iodinated ovine LH LER 1056 C2 which was used as trace. Tubes for the standard curve were prepared with 826 ovine LH reference preparation obtained from NIDDK. LH values that fell below the lowest limit of detectability (0.5 ng) were rounded up to this value for statistical analysis. GnRH+ and Fos+ in grass rats and lab rats in a Iight:dark cycle In the second study, ovariectomized grass rats and lab rats received an injection of EB (so; 50 ug/kg) at either ZT 7 or ZT 19 for two days. On the third day, at the same ZT, females were injected with P (sc; 2.5 mg/kg). Three hours following P injection animals were perfused at either ZT 10 or ZT 22 (n=7/group except grass rats at ZT 10; n=13). One series of tissue was processed for the immunocytochemical detection of GnRH+ and Fos+ as described above. In the first study I found that the highest percentage of GnRH+ cells containing Fos was in the OVLT and that GnRH+ cell activity in this region was representative of that 15 of the total GnRH+ cell population. Therefore, in this study I counted cells in two sections containing the OVLT. GnRH-I- and Fos in grass rats and lab rats in constant darkness In the third study ovariectomized lab rats and grass rats were released into constant darkness (DD). On the third and fourth day of DD animals received injections of EB (sc, grass rats=10 jig/animal, lab rats =50 pg/kg) between either ZT 7 to 8 or ZT 19 and 20. On the fifth day in DD animals were injected with P (so, grass rats=125 jig/animal, lab rats=2.5 mg/kg) then perfused three hours later between ZT 10 and 11 or ZT 22 and 23 (grass rats n=4/timepoint, lab rats n= 5/timepoint). One tissue series from each animal was processed for GnRH+ and Fos+, and two sections containing the OVLT were analyzed as described above. Statistical analysis Percentage data, which is nonparametric, were transformed for statistical analyses. In the first study I wanted to determine if hormone treatment was able to induce a rise in LH levels and/or percent of GnRH cells that contained Fos (log transformed). Data were plotted and examined to identify the timepoint when the “surge” occurred. Data from animals at this timepoint were then compared those of animals killed at ZT 18.5 using an unpaired t-test. These data were also examined with a two-sample t-test using P treatment as the independent variable. That is, animals perfused at 2T 18.5 were compared to the rest of the animals combined. I also used a chi-square analysis to compare plasma LH values in P treated (ZT 20.5, 22, 23.5) and untreated animals (ZT 18.5). I 16 predicted that both LH and GnRH+ cell activity would be higher at timepoints following P treatment, thus I used one-tailed tests in these analyses. To determine whether sub-populations of GnRH+ neurons were different with respect to temporal patterns of activity, I used a two-way ANOVA. Time and region (MS/DBB, OVLT, and AVPV/POA) were the independent variables, and percent of GnRH+ cells that were active (sine transformed) was the dependent variable. The percent of GnRH+ cells that contained Fos (not transformed) was correlated with the plasma LH concentration (ng/ml). Data from the second and third experiments were analyzed using a two-way ANOVA with species and time as independent factors and the log-corrected percent of GnRH+ cells that were active as the dependent variable. All analyses were done using Statview 5.0 and differences were considered significant when p < 0.05. Results Patterns of change in LH and in GnRH-I- cells in grass rats The objective of this first study was to create a model of the induction of the LH surge and associated neuroendocrine events in a diurnal rodent. Both LH levels and GnRH+ cell activity were highest at ZT 22 whereas animals at ZT 18.5 had virtually undetectable plasma LH or active GnRH+ cells (Figure 2.1, Figure 2.2). I therefore used data from ZT 22 to represent the surge in statistical analyses. In steroid-primed grass rats, the percent of GnRH+ cells that were active had a significant increase from ZT 18.5 to ZT 22 (t=-7.12, df=8, p<0.0001, Figure 2.1). Animals perfused at ZT 18.5 (n=5), before P treatment, had 17 Percent of GnR_H cells containing Fos (X:SEM) is __l_. O N - (.3 o 10‘ J O 0 "cm—— 1 - E 18.5 20.5 22 23.5 Zeitgeber time of perfusion Figure 2.1. The average percent (:SEM) of GnRH cells that contained Fos in grass rats. Hormone injections occurred at ZT 19 (arrow). Dots represent the value for each individual (n=5/time point). Zeitgeber time 0 = lights-on. 18 l . o l A o 2 30. w 1 (D 4 +I l5 * _ 20 j E U) C P I . _J 10 . . l . . ,1 . 1 O Owl ..11 I .' ; 18.5 20.5 22 23.5 Zeitgeber time of perfusion Figure 2.2. The average LH value (i SEM) In grass rats. See legend of Figure 2.1 for detail. 19 significantly lower GnRH+ cell activity than did animals killed at the three remaining timepoints (n=15, t = -3.44, df=18, p=0.001). The pattern of change in plasma LH over time resembled that of GnRH+ cell activity. That is, LH levels were significantly higher at ZT 22 compared to levels at ZT 18.5 (t=-1.96, df=8, p=0.04, Figure 2.2). No significant differences in plasma LH levels were detected between P treated animals and untreated animals. There were, however, more LH values over 1 ng/ml among the P treated animals (ZT 20.5, 22, 23.5) than among the untreated animals (ZT 18.5, X2=6.67, df=1, p=0.009). LH values were significantly correlated with the percent of GnRH+ neurons that were active (r = 0.635, 2 value=3.0, p= 0.002). I detected a difference between sub-populations of GnRH+ cells with respect to temporal patterns of activity (time x sub-population, F=2.26, df=6, p=0.05). However, the GnRH+ cell population in each of the regions (MS/DBB, OVLT, and AVPV/POA) had peaks in activity at ZT 22 and this temporal pattern resembled the activity of GnRH+ cells from all regions combined (Figure 2.1). GnRH+ and Fos+ in grass rats and lab rats in a Iight:dark cycle In the second study, steroid primed lab rats and grass rats were perfused just before lights-on (ZT 22) or lights-off (ZT 10). Although hormone treatments were identical in the two groups, GnRH+ cell activity was higher in lab rats perfused at ZT 10 than at ZT 22, whereas grass rats had the inverse pattern (Figure 2.3). This interaction between species and time of day was significant (F=11.37. df=1, p<0.002). 20 El Grass rat 4o - I Lab rat 00 0 containing Fos (X:SEM) N O Percent of GnRH cells A O A l to 2'2 Zeitgeber time of perfusion Figure 2.3. GnRH cell activity in steroid primed grass rats and lab rats housed in a 12:12 Iight:dark cycle. Zeitgeber time 0 = lights-on. 21 GnRH-I- and Fos+ in grass rats and lab rats in constant darkness Lab rats and grass rats kept in DD for 5 days had patterns of change in GnRH+ cell activity that were similar to those of animals kept in Iight:dark cycle (compare Figure 2.3 to 2.4). Specifically, the percent of GnRH+ cells that were active was higher in grass rats perfused at ZT 22-23 than at ZT 10-11 (Figure 2.4). Lab rats showed the reverse pattern, with a higher percentage of GnRH+ cells that were active at ZT 10-11 than at ZT 22-23. There was a significant interaction between species and time (F=152.5, df=1, p<0.001). Discussion In the first study, I established an effective diurnal animal model of neuroendocrine events associated with the estrous cycle. Grass rats treated with EB and P had a peak in both LH and GnRH+ cell activity two hours before lights- on (Figure 2.1, Figure 2.2). The second experiment established that both grass rats and lab rats have rhythms in sensitivity to steroid hormones with respect to GnRH+ neuron activity, and that these rhythms are temporally reversed in these two species (Figure 2.3). Data from the third experiment indicate that in these two species the temporal organization of responsiveness to steroid hormones is under circadian control (Figure 2.4). In grass rats in the current study, the steroid induced LH surge and the activation of GnRH+ cells mirrored the patterns that occur in intact grass rats during the post partum estrous period, confirming the physiological relevance of this model (McElhinny et al., 1999). The peaks in LH and GnRH+ cell activity also resembled those occurring in other proestrous or steroid primed laboratory 22 [3 Grass rat I Labrat 90: .223 1‘ 8%70‘ 1 I - Eéwi (I) 0 Eu. . O o) d 4—0 c I 82 30‘ 29 ‘ a) . o.§ . 10— l h 10-11 22-23 Zeitgeber time of perfusion Figure 2.4. Percent of GnRH cells that were active in steroid primed grass rats and lab rats housed in constant darkness for 5 days before perfusion. 23 rodents (e.g. Bronson and Vom Saal, 1979; Doan and Urbanski, 1994; Finn et al., 1998; Lee et al., 1992; Rajendren, 2001; Wu et al., 1992). Average LH values, even at ZT 22 (Figure 2.2), were somewhat lower than those typical of a surge produced by females during the postpartum estrus (McElhinny et al., 1999) and the variability was high. One explanation for this could be that the hormone treatments I utilized were only partially effective for the induction of the surge. Alternatively, the variability might be due to inter-individual differences with respect to the exact time of the surge. My sampling times may not have coincided with the peak in serum LH levels which may vary among individuals and may be brief, as is the case in lab rats and mice (Bronson and Vom Saal, 1979; Levine and Ramirez, 1982). However, at ZT 18.5 no individuals had an LH titer over 1 ng/ml whereas all five animals at ZT 22 and 4 of 5 animals at ZT 23.5 had values that were above 1 ng/ml, suggesting that the steroid treatment did cause LH levels to rise. There was a tight correlation between the level of LH and the GnRH+ cell activity in the first study. This relationship has also been seen in other laboratory animals such as mice, lab rats and guinea pigs (King et al., 1998; Lee et al., 1992; van der Beek et al., 1994; Wu et al., 1992). In intact and steroid treated lab rats, the percent of GnRH cells that are active is highest during the ascending phase of the LH surge (Lee et al., 1992; van der Beek et al., 1994). In hamsters, the rise in GnRH cell activity follows the peak in plasma LH values and may be involved in the termination of the gonadotropin surge in this species (Doan and Urbanski, 1994). This is unlikely to be the case in hormone-primed grass rats 24 because both the LH values and GnRH+ cell activity were highest at ZT 22 and were quite low at ZT 20.5. If one event preceded the other it would have to occur within the same hour, a temporal pattern unlike that seen in hamsters. Results from the current studies indicate that grass rats and lab rats have a daily rhythm in the responsiveness of GnRH+ neurons to the positive feedback effects of steroid hormones (Figure 2.3, Figure 2.4). Ovarian steroids induce a rise in the proportion of GnRH-I- cells that contain Fos+ in ovariectomized lab rats and mice (Hoffman et al., 1990; Lee et al., 1990b; Wu et al., 1992). These studies did not reveal a rhythm in responsiveness, as they did not look outside the time of day at which the proestrous LH surge normally occurs. Earlier evidence of an endogenous rhythm has come from organotypic cultures of the POA. In these cultures, GnRH is secreted in a circadian pattern but only when the tissue is incubated with estradiol (Funabashi et al., 2000). To the best of my knowledge, the current results represent the first demonstration of a daily rhythm in the expression of Fos+ in GnRH+ neurons in any species. Endogenous timing mechanisms appear to drive this rhythm in GnRH+ cell activity in both lab rats and grass rats (Figure 2.3, Figure 2.4). For both species, animals appeared to retain the rhythm after five days in constant darkness. That is, grass rats sacrificed before their subjective day had a dramatically higher percentage of GnRH+ cells that were active than those sacrificed 12 hours later; in lab rats this pattern was reversed (Figure 2.4). It remains possible that the rhythm in either species would have dampened out if animals had been left in constant darkness for a longer period of time. This is 25 unlikely however, because there was no hint that the rhythm was diminished after animals had been kept in DD for five days. Furthermore, in lab rats, the timing of the LH surge is driven by an endogenous circadian system and LH release is driven by the GnRH neurons (Alleva et al., 1971; Fitzgerald and Zucker, 1976; Takeo, 1984). These data support the idea that the rhythms I observed in GnRH neurons are indeed endogenous. The neural mechanisms that regulate the rhythm in activity of GnRH neurons have not been established in either species but the SCN is likely to play a role though its projections to these neurons. The SCN may communicate temporal information directly to cells responsible for coordination of estrous—related events. In lab rats and hamsters, tract—tracing studies suggest that the SCN projects to both estrogen receptor and GnRH containing cells, and lesions of the SCN eliminate the majority of contacts between the SCN and GnRH neurons (de la lglesia et al., 1995; van der Beek et al., 1998; van der Beek et al., 1993; Watson et al., 1995). The temporal pattern of coupling observed here between GnRH cell activation and the LH surge was similar in lab rats and grass rats. The SCN may regulate the timing of these events, at least in part, through similar pathways in these two species. Preliminary evidence from my lab indicates that, in the grass rat, GnRH+ cells do in fact receive input from the SCN (Mahoney and Smale, 2000; chapter 3). The current data raise interesting questions regarding the specific mechanisms responsible for the species differences in the timing of GnRH cell recruitment. Whereas proestrous and steroid-primed grass rats and lab rats are similar with respect to the temporal relationship between estrous-related events, 26 the timing of these events relative to the Iight:dark cycle is dramatically different. This difference is related to the animal’s chronotype as both lab rats and grass rats have peaks in GnRH+ cell activity that occur around the onset of their activity periods. These differences in the timing of neuroendocrine events might be due to differences in the timing of SCN signals, differences in which neurochemical signals are emitted by the SCN, or differences in the responsiveness of GnRH and ER-containing cells to such signals. These closely related species provide a unique opportunity to evaluate these hypotheses and to determine how the neural mechanisms underlying circadian rhythms differ in diurnal and nocturnal species. 27 Chapter 3 Projections from the suprachiasmatic nucleus of grass rats to gonadotropin releasing hormone and estrogen receptor containing cells Introduction The mammalian suprachiasmatic nucleus (SCN) is the primary site for the generation and synchronization of circadian rhythms. Cells of the SCN have circadian rhythms in electrical activity, neurotransmitter secretion, and glucose uptake both in vivo and in vitro (lnouye and Kawamura, 1979; Murakami et al., 1991; Schwartz et al., 1983; Shinohara et al., 1998; Sodersten et al., 1985; Watanabe et al., 1993; Welsh et al., 1995; Yamazaki etal., 1998). Destruction of this nucleus eliminates behavioral and hormonal rhythms and transplants of fetal SCN tissue into animals with bilateral lesions restore a number of these rhythms (LeSauter and Silver, 1999; Mahoney et al., 2001; Moore and Eichler, 1972; Ralph et al., 1990; Stephan and Zucker, 1972). Several lines of evidence indicate that the circadian clock is critical for the timing of estrous-related events in some nocturnal rodents. In female lab rats and hamsters, rhythms in the timing of both the preovulatory surge of luteinizing hormone (LH) and the onset of sexual receptivity persist in the absence of environmental cues (Alleva et al., 1971; Lucas etal., 1999; Takeo, 1984). When these rhythms are phase shifted by changes in the Iight:dark cycle or by pharmacological treatments, the onset of behavioral estrus and the LH surge maintain a precise temporal relationship to other circadian rhythms such as the 28 onset of wheel-running activity (Fitzgerald and Zucker, 1976; Moline et al., 1981; Swann and Turek, 1982). Lab rats and hamsters with bilateral SCN lesions fail to exhibit an LH surge, a consistently functional estrous cycle, or behavioral rhythms in responsiveness to steroid hormones (Gray et al., 1978; Hansen et al., 1979; Kawakami et al., 1980; Meyer-Bernstein et al., 1999; Weigand and Terasawa, 1982). Interestingly, SCN transplants that restore behavioral rhythms in lesioned animals do not restore estrous cyclicity, suggesting that synaptic contacts, rather than humoral signals, communicate temporal information from the SCN to cells that regulate reproductive functions (Meyer-Bernstein et al., 1999) Two cell types that are critical for mammalian reproductive function are gonadotropin releasing hormone (GnRH) and estrogen receptor-o containing (ER) neurons. ER knock-out mice do not exhibit the full suite of female sexual behaviors (Ogawa et al., 1998; Rissman et al., 1997) and hypogonadal mice that lack a functional gene for GnRH are infertile and have relatively low levels of gonadotropins (Gibson et al., 1997). Tract-tracing studies of lab rats and hamsters have revealed that the SCN projects directly to both GnRH- and ER- containing cells. The majority of GnRH cells contacted by SCN efferents in these species are located in the organum vasculosum of the lamina terminalis (OVLT) and the preoptic area (POA) of lab rats, and in the diagonal band of Broca (DB8) and POA of hamsters (de la lglesia et al., 1995; van der Beek et al., 1997). The SCN is also known to project to ER containing cells throughout many regions of the hypothalamus of hamsters (de la lglesia et al., 1995). Although a similar 29 systematic analysis has not been done in lab rats, SCN efferents are known to synapse on the some of ER cells in the anteroventral portion of the periventricular nucleus (AVPV) in this species (Watson et al., 1995). The AVPV is essential for the LH surge; animals with lesions of this nucleus fail to exhibit a rise in LH in response to steroid hormone treatment (Simerly, 1998; Weigand and Terasawa, 1982). These data suggest that one mechanism by which the circadian system mediates rhythms in the timing of reproductive events is via a direct pathway from the SCN to GnRH- and steroid-sensitive cells. The region immediately dorsal to the SCN, the lower sub-paraventricular zone (LSPV), may also play an important role in the temporal organization of reproductive events. The LSPV receives a massive input from the SCN (Kalsbeek et al., 1993; Leak et al., 1999; Morin et al., 1994; Watts et al., 1987) and it appears to project to many of the same targets as the SCN, including GnRH- and ER-containing neurons (de la lglesia et al., 1995; van der Beek et al., 1997; Watson et al., 1995). Interestingly, these two kinds of cells also send efferents back to the LSPV and SCN (de la lglesia et al., 1999; van der Beek, Wiegant et al., 1997). In ovariectomized lab rats, lesions of the LSPV result in an increase in tonic levels of LH (Docke et al., 1982). Furthermore, knife cuts of the projection from the SCN to the LSPV result in an attenuation of the steroid- induced LH surge (Watts et al., 1989). These data raise the possibility that SCN projections to the LSPV might also play a role in transmitting circadian signals that regulate estrous-related events. 30 Several lines of evidence indicate that the neuropeptides vasoactive intestinal polypeptide (VIP) and arginine vasopressin (AVP) may mediate the effects of the SCN on the timing of the LH surge and GnRH cell activation. In lab rats and hamsters, VIP fibers appear to terminate on GnRH cells and in lab rats SCN lesions eliminate about 80% of these contacts (Horvath et al., 1998; Krajnak et al., 2001; Kriegsfeld et al., 2002; van der Beek et al., 1998; van der Beek et al., 1993). Receptors for VIP are expressed on GnRH neurons in vivo and in vitro (Olcese et al., 1997; Smith et al., 2000), and female lab rats have more contacts between VIP boutons and GnRH cells, and more hypothalamic VIP protein than do males (Horvath et al., 1998; Riskind et al., 1989). Furthermore, the rhythm in VIP mRNA expression in female lab rats is 12 hours out of phase from that of males (Krajnak et al., 1998). These differences between males and females may be causally related to sex differences in the ability to produce the preovulatory LH surge. Although VIP treatment affects LH secretion in lab rats, conflicting evidence exists as to whether VIP provides an excitatory or stimulatory signal. One interpretation of the literature is that acute VIP treatments have an excitatory effect on LH release whereas chronic exposure to VIP is inhibitory. Short pulses (60 seconds) of VIP infused into the ventricles (icv) lead to an increase in plasma LH (Vijayan et al., 1979). An icv microinjection of either antiserum or antisense oligonucleotides to VIP prior to the surge delays and inhibits the preovulatory or estradiol-induced peak of this hormone (Harney et al., 1996; van der Beek et al., 1999). On the other hand, lab rats administered VIP icv for 1.5 hours or more 31 have relatively low LH levels (Alexander et al., 1985; Stobie and Weick, 1989; Weick and Stobie, 1992; Weick and Stobie, 1995) and this inhibitory effect can be blocked by treatment with a VIP receptor antagonist (Weick and Stobie, 1995) AVP also appears to mediate the timing of estrous-related events such as LH secretion and sexual receptivity (Sodersten et al., 1985; Sodersten et al., 1983). However, as with VIP, the data are conflicting with respect to whether AVP inhibits or stimulates LH secretion. AVP administered icv to steroid-primed, ovariectomized female lab rats inhibits the LH surge (Salisbury et al., 1980) whereas infusions of AVP into the medial preoptic region have a stimulatory effect (Palm et al., 1999; Palm et al., 2001). In intact lab rats, the proestrous LH surge can be blocked by an icv injection of an AVP receptor antagonist (Funabashi et al., 1999). AVP may regulate GnRH neurons via direct input to them. AVP fibers contact GnRH neurons in the medial preoptic area of female lab rats and hamsters and in the supraoptic nucleus of female cynomolgus monkeys (Huhman and van Der Beek, 1998; Thind et al., 1991; van der Beek et al., 1998). SCN lesions in lab rats reduce the number of these contacts ' suggesting that at least some of these AVP fibers come from cells within the SCN (van der Beek et al., 1998). Interestingly, in co-cultures of tissue containing the SCN and POA, circadian rhythms in AVP and GnRH release have identical periods and these differ from those of rhythms in VIP secretion. AVP administration to these cultures induces GnRH release, but VIP has no effect (FunabashietaL,2000) 32 Rhythms in the timing of estrous-related events such as mating, the preovulatory surge in LH, and GnRH cell activity are inverted in diurnal and nocturnal species (Bronson and Vom Saal, 1979; Dobson and Michener, 1995; Legan and Karsch, 1975; McElhinny et al., 1999; Michener, 1980; Rood, 1970; Seegal and Goldman, 1975; Sodersten, 1988; Stetson and Gibson, 1977; Yeoman et al., 1991; Chapters 2 and 4). These differences might be due to differences in the functional and/or anatomical relationships between SCN efferents and GnRH neurons. Specifically, nocturnal and diurnal species might differ with respect to the temporal pattern of transmitter release from SCN cells, which neurochemical signals are released from these cells (e.g. VIP or AVP), or - the sensitivity of target cells to SCN signals. Nocturnal and diurnal species may also differ with respect to patterns of connectivity between the SCN and GnRH- and/or ER-containing neurons, or some combination of these various factors. The murid rodent Arvicanthis niloticus (grass rat) has proven to be an ideal model with which to investigate how the circadian system differs between diurnal and nocturnal species. This species exhibits a host of rhythms that are reversed relative to those of lab rats. These include rhythms in the timing of the LH surge, associated GnRH cell activation, and copulatory behavior (Blanchong et al., 1999; McElhinny etal., 1999; McElhinny et al., 1997). Here, I used the grass rat to address the issue of whether the SCN might be anatomically positioned to regulate rhythms in estrous-related events in a diurnal species. My aims were: 1) to determine whether the SCN innervates GnRH- and/or ER- containing cells and 2) to determine whether AVP and VIP immunoreactive fibers 33 contact GnRH cells in grass rats. I also mapped the distribution of SCN efferents, AVP- and VIP-containing fibers, and ER-containing cells in grass rats. The hypothalamic distribution of GnRH cells has already been described for this species (McElhinny et al., 1999). Materials and methods Animals Adult female grass rats (>60 days) bred from laboratory stock and Sprague Dawley rats (Charles River) were singly housed in a 12:12 Iight:dark cycle and provided food (Teklad rodent chow 8640, Harlan Industries) and water ad libitum. A red light (< 5 lux) was left on continuously in each animal room. All experiments were performed in compliance with Michigan State University All- University Committee on Animal Use and in accordance with the standard in the National Research Council Guide for the Care and Use of Laboratory Animals. All efforts were made to minimize the suffering and the number of animals used in these experiments. Surgeries 1. Tract-tracing Animals (n=13) were anesthetized using sodium pentobarbital (Nembutal, <50 mg/kg, Abbott Laboratories) and supplemented with methoxyflurane (Metofane, Mallinckrodt Veterinary). The tops of their heads were shaved, cleaned with topical antiseptic (Betadine, Novaplus), and 2% lidocaine was subcutaneously (sc; 0.03 ml, Abbott Labs). Animals were then placed in a stereotaxic apparatus with the tooth bar set at —0.6 mm, the skull was exposed 34 and a hole was drilled (0.14 anterior and +0.13 mm lateral to bregma). A glass pipette (inner tip diameter = 10-15 pm) was set at a 10° angle (relative to the dorsal-ventral plane), filled with tracer, and then lowered to -0.61 mm ventral to dura. Animals received a unilateral injection of the anterograde tracer biotinylated dextran amine (BDA; 10,000 MW Sigma, 10% solution in H20 (Coolen and Wood, 1998; Sequeira et al., 2000)). BDA was delivered via iontophoresis over a ten-minute period with an alternating 5-microamp positive current (7 sec on/7 sec off). After surgery, animals received sterile saline (1 cc 0.9% sc) and the analgesic buprenorphine hydrochloride (0.03 mg, intramuscularly; im, Buprenex, Reckitt & Coleman). The incision was closed with wound clips and treated with topical antibiotic (Nolvasan, Fort Dodge Animal Health). Animals were perfused after a seven-day recovery period (Richardson, 2002; Veenman et al., 1992). 2. Ovariectomy In some studies grass rats were bilaterally ovariectomized while under Nembutal anesthesia (<50 mg/kg) supplemented with Metofane. lncisions were closed with sutures and treated with Nolvasan. Following ovariectomy animals were given saline (1 cc 0.9%, so) and Buprenex (0.03 mg, im). Animals recovered for at least seven days prior to steroid hormone injections. Tissue processing and analysis 1. General immunocytochemical procedure Animals were deeply anaesthetized with Nembutal and perfused transcardially with 0.01 M phosphate buffered saline (PBS; pH 7.4, 150—200 35 mI/animal) followed by 4% paraformaldehyde (150-200 ml/animal, Sigma) in 0.1 M phosphate buffer. Brains were post-fixed in paraformaldehyde for four hours, transferred to 20% sucrose in 0.1 M phosphate buffer overnight, and then sectioned with a freezing microtome. Brains were cut into three series at 30 pm with the exception of tissue from steroid treated females labeled for VIP and GnRH, which was cut into two series at 40 um. Brains from animals with BDA injections were sectioned through to the intergeniculate leaflet, and brains from all other animals were cut from the medial septum to the supraoptic nucleus. All immunocytochemical labeling began with the following steps: free floating tissue sections were incubated in (1) 5% normal serum (in PBS with 0.3% triton-X; TX) for one hour at room temperature, followed by (2) primary antibody for 24 hours at 4°C (in PBS with 0.3% TX and 3% normal serum), followed by (3) biotinylated secondary antibody for one hour at room temperature (in PBS with 0.3% TX and 3% normal serum), followed by (4) avidin-biotin complex (Vectastain Elite Kit, Vector Laboratories, 0.9% of avidin and biotin) for one hour at room temperature. For nickel-enhanced labeling (blue chromagen) the tissue was then rinsed in sodium acetate buffer (0.175 M, Sigma) three times for ten minutes and then reacted in a solution of diaminobenzidine (DAB, 0.25 mg/ml, Sigma), 3% hydrogen peroxide (0.825 pl/ml buffer) and 2.5% nickel sulfate in sodium acetate buffer. Tissue labeled with standard DAB chromagen went from step (4) into DAB (0.5 mg/ml, in Trizma buffer, pH 7.2) with 30% hydrogen peroxide (0.35 ul/ml buffer). Unless otherwise noted, tissue was rinsed three times for ten 36 minutes in PBS between each incubation step. Reagents used for immunocytochemistry are listed in Table 3.1. Following the labeling, tissue was mounted, dehydrated, coverslipped and examined under a light microscope (Laborlux S, Leitz Wetzlar GBH). In all cases, negative controls were done by performing the above procedures but with the omission of the primary antibody(s). No inappropriate staining was detected in control tissue. 2. ER+ cell distribution Here I characterized the distribution of ER immunoreactive (+) cells in female grass rats. Ovariectomized animals (n=4) received an injection of estradiol benzoate (EB, in sesame oil, 10 ug/animal, so) for two days. Twenty- four hours after the second EB injection, animals were perfused. One series of tissue was processed for the detection of ER using standard DAB as the chromagen (Table 3.1 ). A camera lucida was used to draw the distribution of ER+ cells in regions known to receive SCN input in the lab rat (Watts et al., 1987). These images were then scanned into a drawing program (Photoshop 7.0) and retraced to produce pictures of ER+ cell distribution. Table 3.2 contains abbreviations for structures indicated in the drawings. 3. BDA labeled fibers 3a) Distribution of BDA labeled fibers One series of tissue from BDA treated animals was processed to determine the site of injection. The tracer was already biotinylated, thus tissue was processed for nickel enhanced labeling by beginning with the AB-complex 37 5532 2833.. “mum“. ESE .mmmfimu 83 .mwwwfifim 85.983 :82 88; mmmwcmmzswm mofimfiwwmg .moo mcfiwficowmmu com; «Marmfiupwwo motoficonmu m_:m:_con_ .wwwwmfi d><-=cm 9a 3530 wefiwfiufi aoo mefiwwfifi 8N; wwwrmfiunawmo 3.2983 «.358; 88; usage 9a 853 Momma”... >mxcoo ”flaw“... com; HMMMWHLM mmwmn .chszQE cooEoco coon; Imcwécm zoom”. 59.8.. 59.02. . . Louco> “HM” ..oucg cozazu “Wan—“Wm.“ ..ouco> .3225 325.5 9:: can ucaflbfiofi >23...” mm; medial POA > lateral POA. In the BNST, ER+ cells were observed both dorsal and ventral to the anterior commisure, and this cell population increased in more caudal regions of the nucleus (Figure 3.5). At the level of the SCN and RCH, numerous ER+ cells were diffusely distributed through the anterior hypothalamus and BNST (Figure 3.5). ER+ cells were not detected in the SCN. BDA fibers clearly projected to regions containing ER+ positive cells (Figure 3.4B, Figure 3.6). These labeled fibers wound around and between ER+ nuclei, and in many cases, appeared to contact the nuclei themselves (Figure 3.6C,D, E). ER+ cells were always detected in regions that contained BDA 48 Figure 3.4. Photomicrographs depicting A) ER+ cells in the AVPV of an estradiol primed grass rat. B) Low power view of patterns of overlap between BDA labeled fibers (blue) and ER+ cells (brown) in the PeVN (left side of image). Note that as ER+ staining decreases so does the density of BDA staining. Abbreviations are found in Table 3.2. 49 Figure 3.5. A series of line drawings at 5 levels from rostral (1) to caudal (5) depicting the distribution of ER+ and GnRH+ cells and AVP+ and VIP+ fibers in the hypothalamus of grass rats. In figures of ER+ and GnRH+ each black dot represents 1 cell. Pictures of GnRH+ cell distribution are modified from McElhinny et al. (1999). See Table 3.2 for abbreviations. 50 section ER-or VIP AVP GnRH .i V“ Figure 3.5 m f 51 \HP AVP GnRH Figure 3.5 cont. . ...-.4 1‘ Figure 3.6. Photomicrographs depicting BDA fibers (blue staining) overlapping with ER+ cells (brown staining). Arrows indicate places where BDA fibers are in close apposition with ER+ cells. 53 ‘1' I l-‘ .m- 54 labeled fibers (compare Figure 3.2 with Figure 3.5). In comparison, BDA fibers were also detected in regions devoid of ER+ staining, however, fibers were less dense in these areas compared to regions of overlap between BDA and ER+ cells (Figure 3.48). BDA fibers and GnRH+ cells Bipolar GnRH+ cells were visible as large, brown, oval-shaped soma that had long processes with relatively evenly spaced varicosities extending from them. As reported previously (McElhinny et al., 1999), GnRH+ cells and fibers were observed in the MS, the horizontal and vertical limbs of the DBB, OVLT, subdivisions within the POA, and in a region dorsal and lateral to the SON (Figure 3.5). GnRH+ cells were not found within the SCN, however, GnRH+ cells and fibers were detected within the LSPV and along the optic chiasm, just lateral to the nucleus. BDA fibers were seen in apposition to GnRH+ cells in all regions examined (Figure 3.7C-G, Table 3.3). In particular, a dense plexus of BDA fibers surrounded GnRH+ cells that were located in the OVLT and AVPV. Interestingly, BDA and GnRH+ fibers also overlapped, particularly in these two regions (Figure 3.7A, B). AVP+ and VIP+ fiber distribution The distributions of AVP+ and VIP+ fibers in female grass rats are depicted in Figure 3.5. Both AVP+ and VIP+ fibers were relatively thin and fine, with irregular, small varicosities. In rostral sections, AVP+ fibers were present in the ventral and central portion of the LS, with scattered fibers found throughout 55 Figure 3.7. A, B) Photomicrographs depicting the pattern of overlap between BDA labeled fibers (blue) and GnRH+ fibers (brown) in female grass rats. C-G) BDA labeled fibers contacting GnRH+ cells. Arrows indicate putative contacts. Table 3.3. Percentage of GnRH+ cells contacted by BDA labeled fibers. GnRH+ cells were analyzed in tissue from 3 individuals with iontophoretic BDA injections in the SCN. Abbreviations can be found in Table 3.2 GnRH-I- cell location % GnRH+ cells contacted by BDA fiber (18EM) MS/DBB 66 1 0.08 OVLT 72 1 0.09 AVPV 90 1 0.10 L POA/LSPV 67 1 0.12 57 the MS. Fibers were also detected in the BNST, but were relatively sparse in the OVLT. Darkly stained AVP+ cells were concentrated in the dorsal portion of the SCN. Extensive AVP+ fibers extended from the SCN into the LSPV, and a moderate number of fibers continued into the PVN. Large magnocellular AVP+ cells were located in the SON and PVN. Labeled fibers originating in these nuclei were thicker than those found elsewhere. The distribution of VIP+ fibers overlapped with that of AVP+ fibers in some regions (Figure 3.5). In the most rostral sections examined, VIP+ fibers were present in the ventral portion of the LS and within the BNST surrounding the anterior commisure. In sections containing the OVLT, a moderate number of short VIP+ fibers were scattered around the 3rd ventricle and optic chiasm. At this level, abundant VIP+ fibers were detected within the BNST. VIP+ cells were seen in the ventral portion of the SCN, along the dorsal edge of the optic chiasm, and VIP+ fibers were abundant throughout the rostrocaudal extent of the SCN. Numerous VIP+ fibers extended dorsally from the SCN into the LSPV and the PeVN. AVP+ and VIP+ contacts on GnRH-I- neurons At the Iight microscope level, VIP+ and AVP+ contacts on GnRH+ cells were easily detected and were observed in all regions (Figure 3.8, Figure 3.9, Table 3.4). Typically, contacts were present as a dark blue bouton-like structure at the edge of the GnRH+ cell soma, although frequently a somewhat longer fiber was observed approaching and wrapping around the cell body and dendrite (Figure 3.80). 58 Figure 3.8. Photomicrographs depicting AVP+ fibers (blue) contacting GnRH+ cells (brown) in the hypothalamus of grass rats. Pictures in C and D are me same image taken at different planes of focus. Arrows indicate putative contacts. 59 Figure 3.9. Photomicrographs of VIP+ fibers (blue) contacting GnRH+ cells (brown) in a steroid primed female grass rat. Arrows indicate putative contacts. 60 Amm_mE9 3.0“ RN 3%.. 9.: two“ ....N 3m 0.: no“ 9N 3H 9 9% m: Eu 3 wwofim Bus :5 I I I I I I I I $28.9 8.? k3 3+ «.9. mm? a: N. F I F. mud + N3 5: man 8.? x: 9. mm 895 mus :5 9.0“ 84 in 3a to“ F: 03H n.3, SSH 9.0 9H cm 9.0“ 36 EH 3: mmuwflw __ 8 ...Imcw n909:8 =8 889:8 =8 +Imco 889:8 =8 +Imco 889:8 3:938 u 2.8 +Imco m=8 3:928 # m__8 8:938 3.. m=8 ..Imco as 89:8 a face as +195 as face as .28 >n_m.=:>< 5>o mew: :28: :2me 5.25 99 896 he maEflafioEE 05 :_ 28: 12> new +d>< >3 889:8 mcoSc: ...Imcw Co “:85.“ .vd 28... 61 AVP+ fibers contacted about 24% of GnRH+ cells (Table 3.4). The majority of these contacted cells were located in the AVPV and along the optic chiasm, lateral to the SCN. In intact female grass rats, VIP+ fibers contacted about 35% of the total population of GnRH+ neurons (Table 3.4). These cells were located predominantly in the MS and DBB; slightly more than half of the GnRH+ neurons present in these regions received VIP input. Generally, individual GnRH+ neurons had more contacts of VIP+ fibers than AVP+ (1.67 compared to 1.08 boutons/cell respectively). In the LSPV, VIP+ and AVP+ fibers overlapped with GnRH+ fibers. In steroid-primed grass rats, as in intact female grass rats, VIP+ fibers contacted GnRH+ neurons. Whereas females killed at ZT 20.5 had relatively low LH titers (1.68 11.2 ng/ml 1SEM) when compared to those of animals killed at ZT 22 (15.15 1 7 ng/ml 1SEM), I did not detect a significant difference in the percent of GnRH+ cells contacted by VIP+ fibers between the two groups (Mann-Whitney U, p=0.17). I thus combined the data; VIP+ fibers contacted about 14% of the total GnRH+ cell population (Table 3.4). Although there were typically more GnRH+ neurons within the OVLT (data not shown), all regions contained about the same percentage of GnRH+ neurons contacted by VIP fibers (range 14- 18%). The average percent of GnRH neurons contacted by VIP+ fibers was somewhat less in steroid-primed females than in intact females. Table 3.5 summarizes the data from all these studies. 62 Table 3.5. Summary of data indicating the location and density of BDA labeled fibers, VIP+ and AVP+ fibers, and ER+ and GnRH+ neurons in grass rats. Scale of density ranges from least dense (+) to most dense (++++). A dashed line indicates that labeling was not detected. Fiber and cell densities reflect the range for a given type of labeling and are not directly comparable across substances. See Table 3.2 for abbreviations. BDA VIP+ fibers AVP+ fibers ER+ cells GnRH-I- cells LS +++ + ++ + - MS + - ++ - + BNST ++ +++ ++ ++ - PeVN +++ ++ + ++++ + OVLT ++ ++ + + ++++ AVPV +++ + + ++++ ++ POA ++ + + ++ ++ SCN ++++ ++ ++ - - LSPV ++++ +++ +++ - + SON + - + - ++ 63 Discussion Results from this study indicate that grass rats are similar to nocturnal rodents with respect to projections of the SCN to the anterior hypothalamus. In grass rats, BDA injections that hit the SCN labeled fibers in nuclei known to receive SCN input in lab rats and hamsters (Table 3.5, Buijs et al., 1993; Kalsbeek et al., 1993; Morin et al., 1994; Watts and Swanson, 1987; Watts et al., 1987). The distributions of VIP+ and AVP+ fibers in grass rats also resembled those of other rodents (Figure 3.5; Abrahamson and Moore, 2001; Kalsbeek et al., 1993; Morin et al., 1994; Watts and Swanson, 1987). The distribution of ER+ cells in the hypothalamus of grass rats also resembled that seen in a host of other mammals, including nocturnal lab rats and hamsters (Table 3.5; Figure 3.5; Blaustein, 1992; Cintra et al., 1982; Don Carlos etal., 1991; Fox etal., 1991; Hnatczuk etal., 1994; Li et al., 1993; Tobet etal., 1993). ER+ cells were all located in regions that received input from the SCN (Table 3.5). ER is typically expressed in the cell nucleus; thus, I was unable to establish definitely whether BDA-labeled fibers contacted ER+ cells. However, frequently ER+ cells were seen surrounded by a dense plexus of BDA fibers suggesting that these steroid sensitive cells receive information directly from the SCN in grass rats, as is the case in lab rats and hamsters (Figure 3.4, Figure 3.6; de la lglesia et al., 1995; Watson et al., 1995). As in McElhinny et al., 1999, I found that the distribution of GnRH+ neurons in grass rats parallels that seen in other mammalian species. Additionally, I detected SCN input on GnRH+ cells in all regions that l analyzed 64 (Table 3.3, Table 3.5). However, I found that 60-90% of these neuroendocrine cells were contacted by BDA labeled fibers, an estimate that is much higher than that for lab rats (~2-20%; van der Beek et al., 1997) or hamsters (11-13%; de la Iglesia et al., 1995). These differences might be related to the tracers used (BDA vs. Phaseolus vulgaris leucoagglutinin), the locations and sizes of the injections, numbers of filled cells, efficacy of the transport, or the presence of retrogradely labeled fibers. BDA is predominantly transported in the anterograde direction, but some BDA contacts on GnRH+ perikarya might come from retrogradely labeled neurons (Rajakumar et al., 1993; Veenman et al., 1992). I found that in grass rats, 24% of GnRH+ neurons were contacted by AVP+ fibers, a proportion similar to that reported for male lab rats (19%) but higher than that seen in females (10%; van der Beck et al., 1998). It is quite possible that some AVP+ fibers contacting GnRH+ cells were not SCN efferents but rather originated from cells in the SCN, PVN, or elsewhere. The largest proportion of GnRH+ cells contacted by AVP+ fibers was found in the AVPV/POA/LSPV. These regions are relatively close to the SCN so it is likely that at least these GnRH+ neurons were receiving input from AVP+ cells in the SCN. SCN lesions in female lab rats significantly reduced the percent of GnRH+ neurons contacted by AVP fibers in the POA (van der Beek et al., 1998). In intact female grass rats, about 35% of the GnRH+ cells were contacted by VIP+ fibers. These contacts probably came from cells in the SCN, where most of this peptide is produced. This is supported by studies of lab rats where unilateral SCN lesions decreased VIP innervation of GnRH cells on the side 65 ipsilateral to the lesion, compared to the contralateral side (van der Beek et al., 1993). Previous studies on lab rats provide widely varying estimates of the percentage of GnRH cells that receive input from VIP cells. Some report that about 5% of GnRH neurons are contacted by VIP fibers (Kriegsfeld et al., 2002), whereas others report figures of 34% (van der Beek et al., 1994; Horvath et al., 1998) or 45% (van der Beek et al., 1993). In the current study I found that in intact grass rats GnRH neurons were more likely to be innervated by VIP+ fibers (35%) than were steroid-treated females (14%). l was unable to determine the endocrine state of intact grass rats, but undoubtedly they had lower estradiol levels than did steroid-primed females. These varied results from lab rats and grass rats may be due in part to: 1) varying levels of circulating steroid hormones, 2) differences in microscopy techniques (i.e. fluorescent labeling combined with confocal vs. DAB labeling combined with light), or 3) differences regarding whether contacts were counted on dendrites and soma or just on the soma. By combining the data on anterograde tracing of SCN efferents, and immunocytochemistry for two known SCN peptides, I am able to make some inferences about which subsets of SCN cells may project to GnRH neurons in the grass rat (Table 3.5). The distribution of AVP+ fibers matched that of BDA labeled fibers in the SON and MS, regions where VIP+ fibers were not detected. On the other hand, the distribution of AVP+ and VIP+ fibers overlapped in some regions that received SCN input. Fibers containing these two peptides appeared to overlap to a similar extent with regions containing BDA labeled fibers in the 66 POA and LSPV. Within the lateral septum, VIP was contained in fibers in the ventral portion of the nucleus, whereas AVP+ fibers were present in the ventral and central portions. The SCN efferents in the BNST, PeVN, and OVLT overlapped predominantly with VIP+ fibers, but some AVP+ fibers were also detected in these regions. Interestingly, GnRH+ neurons were detected in the AVPV, a region that received extensive SCN input but contained relatively few VIP+ or AVP+ fibers. These data suggest that the neurotransmitters AVP and VIP communicate circadian information to the reproductive axis in this diurnal species, as occurs in nocturnal rodents. It is still possible, however, that the inverted rhythms of estrous-related events in grass rats relative to nocturnal rodents are due to differences in communication between VIP and/or AVP cells in the SCN and GnRH+ cells. For example, the contacts I, or others, have observed may not reflect numbers of actual synapses; ultrastructural studies will be needed to evaluate that possibility. Furthermore, the numbers of contacts are not necessarily indicative of the overall intensity or pattern of signals. It is also important to consider the possibility that other transmitters such as GABA or glutamate are released from VIP and/or AVP terminals in a manner that differs between species. The SCN may also influence GnRH+ neurons indirectly via intermediate cells that might modify the temporal pattern of the signal received by GnRH+ neurons. One such “relay station" might be the LSPV, which receives input from the SCN and projects to many of the same sites, including GnRH+ neurons (de la 67 lglesia et al., 1995; Morin et al., 1994; van der Beek et al., 1997; Watts and Swanson, 1987; Watts et al., 1987). In lab rats, disruption of the projection from the SCN to the LSPV attenuates the steroid-induced LH surge (Watts et al., 1989) and GnRH fibers synapse on structures in this region (van der Beek, Wiegant et al., 1997). In the current study, I detected AVP+ and VIP+ fibers extending from the SCN to the LSPV and some of these fibers contacted GnRH+ cells located there. Together, these data raise the possibility that the LSPV modulates SCN signals relayed to the GnRH system. Unfortunately, I was unable to determine which BDA positive fibers emanated from cells in the SCN and which came from the LSPV. The results of the current study suggest that the SCN may communicate temporal information directly to cells that regulate the timing of estrous-related events in a diurnal species, as they do in nocturnal ones. The similarity between diurnal and nocturnal rodents with respect to SCN targets suggests that SCN input to GnRH+, and perhaps ER+ neurons, might represent a common anatomical pathway for the regulation of reproductive events by the circadian system. This raises the possibility that the mechanisms underlying fundamental differences between nocturnal and diurnal species might reside within GnRH cells, SCN neurons projecting directly to them, or in intermediate structures such as the LSPV. Now that the anatomical groundwork has been laid, future studies may be directed at answering such functional questions. 68 Chapter 4: A daily rhythm in mating behavior and progestin receptor expression in a diurnal murid rodent Arvicanthis niloticus Introduction The timing of many reproductive behaviors and associated physiological events in mammals is related to the activity patterns of the species. For example, diurnal sciurid (e.g. ground squirrels) and hystricomorph rodents (e.g. degus and cavies) mate during daylight hours (Rood, 1970; Dobson and Michener, 1995; Michener, 1980; Rossi and Lee, personal communication), whereas nocturnal rodents such as laboratory rats (“lab rats”), hamsters, and mice typically display more mating behaviors at the beginning and throughout their active period, during the dark portion of the Iight:dark cycle. Female lab rats for example, have higher lordosis quotients (L0) and male rats have shorter latencies to first intromissions and ejaculations, and shorter intervals between these events, during the dark than the light phase of the dayznight cycle (Beach and Levinson; Hansen et al., 1979; Harlan et al., 1980; Spinka, 1990; Stefanick, 1983) In lab rats the daily rhythm in female sexual behaviors is due in part to rhythms in steroid hormone secretion but also to changes in responsiveness to these hormones (Hansen et al., 1979; Hansen et al., 1978; Sodersten et al., 1981 ). Ovariectomized lab rats implanted with estradiol capsules and tested for sexual behavior 3 days later have peak LQ values during the dark and the lowest 69 LQ values during the light portion of the 24-hour cycle (Hansen et al., 1979). Additionally, the response of ovariectomized lab rats to steroid hormone stimulation is phase-dependent. Specifically, when females are tested for sexual behavior 96 hours after receiving an estradiol capsule, they exhibit higher LQ values if the capsule was implanted 4 hours after lights-off than at other times of day (Hansen et al., 1978). Interestingly, not all studies of lab rats have found daily rhythms in sexual behavior (Campbell and Baum, 1979; Erskine et al., 1980; Harlan et al., 1980). Erskine and colleagues (1980) suggest that the discrepancies may be due to differences in the strains of rats that were used in these studies (Wistar, Long Evans, Sprague-Dawley). That is, these strains may exhibit different thresholds of responsiveness to hormone treatment. Rhythms in behavioral sensitivity to steroid hormones might theoretically be caused by rhythms in numbers of available hormone receptors. Progesting receptor-immunoreactive (PR+) cells that are critical for female sexual behavior are located in the ventromedial hypothalamus (VMH) of lab rats and hamsters, and in the ventrolateral hypothalamus (VLH) of guinea pigs (Blaustein and Brown, 1983; Blaustein et al., 1994; DeBold et al., 1982; Don Carlos et al., 1989; Rainbow et al., 1982; Sterner etal., 1992; Young, 1969). Ovariectomized guinea pigs, lab rats, and hamsters implanted with intracranial estradiol-containing cannulae become sexually receptive to systemic progesterone (P) when the cannulae are placed bilaterally in the VMHNLH region (Barfield et al., 1983; Davis et al., 1982; Delville and Blaustein, 1991; Rubin and Barfield, 1983). Furthermore, blocking the expression of PR in the VMH of lab rats with either 70 antisense oligonucleotides to PR mRNA, or by the administration of the P antagonist RU-486, inhibits female sexual behaviors (Etgen and Barfield, 1986; Mani et al., 1994). It is therefore possible that the rhythm in female sexual behavior is promoted in part by a rhythm in the number of PRs in the VMHNLH. To my knowledge, this issue has not been examined in any rodent species. The timing of mating is reversed in the diurnal rodent, Arvicanthis niloticus (“grass rat”) from that of the lab rat. These murid rodents are therefore ideal species for comparative studies that examine the mechanisms underlying the timing of reproductive events. Intact grass rats demonstrate diurnal patterns of rhythmicity with respect to the timing of copulation, the ovulatory surge in luteinizing hormone (LH), and activation of gonadotropin releasing hormone cells (Blanchong et al., 1999; McElhinny, 1996; McElhinny et al., 1999; McElhinny et al., 1997). The reversal in rhythms of mating behaviors in diurnal compared to nocturnal species might be due to inverted rhythms in responsiveness to hormones, to altered patterns of hormone secretion, or both. In this study I examined the first of these hypotheses by pairing male and hormone-primed female grass rats and testing them for mating behavior at four different times of day. I then characterized rhythms in numbers of PR+ cells to determine whether numbers of these cells might account for rhythms in sexual receptivity in female grass rats. Materials and methods: Adult grass rats (>60 days) were singly housed in a 12:12 Iight:dark cycle and provided food and water ad libitum. A red light (<5 lux) was left on 71 continuously for the purposes of animal care and videotaping of behavioral tests. All experiments were performed in compliance with Michigan State University All- University Committee on Animal Use and in accordance with the standard in the National Research Council Guide for Care and Use of Laboratory Animals. All efforts were made to minimize the suffering and the number of animals used in these experiments. Sexual behavior In this first study I determined whether female grass rats have a rhythm in behavioral responsiveness to steroid hormones. Females were anesthetized with sodium pentobarbital (<50 mg/kg Nembutal; Abbott Laboratories) supplemented with methoxyflurane (Metofane, Mallinckrodt Veterinary) then bilaterally ovariectomized. lncisions were closed with sutures and treated with topical antibiotic (Nolvasan, Fort Dodge Animal Health). Animals were then given a subcutaneous (sc) injection of 1 cc 0.9% saline, and 0.03 mg buprenorphine hydrochloride (intramuscularly; im, Buprenex, Reckitt & Colman Pharmaceuticals Inc.). Seven to fourteen days after ovariectomy, females (n=16) were divided into four groups that received steroid hormone injections (sc) at different times of day. For two days, at zeitgeber time (ZT) 1, 7, 13, or 19 (ZT 0= lights-on), females were injected (sc) with10 pg 17-8 estradiol benzoate in sesame oil (EB). On the third day, at the same ZT, females received injections (so) of 125 pg P. Four hours following the P injection (at either ZT 5, 11, 17, or 23), females were placed in a clean, glass aquarium containing cedar shavings, food, and a water bottle and allowed to acclimate for 5 minutes. Then a sexually 72 experienced male was introduced and the pair was videotaped for 30 minutes. Each pair of animals was tested together at all 4 time points, in counter-balanced order. At least one week passed between the conclusion of one behavioral test and the start of the next series of hormone injections. Control females (n=4) were ovariectomized, injected with sesame oil (so, 0.1 cc) and paired with the same male at each of two time points. These females received injections for 3 days at ZT 13 or 19 and were tested at ZT 17 or 23, respectively. The initial 15 minutes of each behavioral test were analyzed. The scored behaviors were mounts, lordosis, and copulation. A mount was counted when the male placed his forepaws on the hindquarters of the female, and a lordosis was scored when a female adopted a stereotyped posture with her head and tail elevated and her back in concave flexion. The L0 was calculated from these scores as the number of female lordosis postures/number of male mounting attempts. A copulation was scored each time the female displayed lordosis as the male mounted her. Finally, I evaluated the longest interval between consecutive copulations as an indicator of mating intensity. These behavioral variables were analyzed using repeated measure ANOVA and pain~ise comparisons were analyzed with Fisher’s PLSD post-hoc tests (Statview 5.0). Data were considered significant when p<0.05. One female was not receptive at any time point and all data from this animal were excluded from analysis. 73 Progesterone receptors Here I determined whether the rhythm of sexual receptivity in female grass rats is associated with a rhythm in the number of PR+ cells. One week following ovariectomy, females were injected with 10 pg EB (so) in sesame oil for two days at either ZT 13 or 19 (n=4/time point). On the third day at the same ZT, females were perfused. In this study, thus, animals received EB injections at times that steroid treatment led to peak and trough levels of female sex behaviors in the first study (ZT 19 and ZT 13, respectively), and the perfusions were done at the corresponding times at which P was injected in that study. Ovariectomized control females did not receive any injections and were perfused at either ZT 13 or ZT 17 (n=4/time point). Because the results of the first study of PRs were unexpected I conducted a second study to determine whether those findings might be replicated, examine PR+ cell numbers in animals perfused at additional time points, and evaluate the efficacy of a different steroid priming protocol. In this study I used intact females implanted with EB-filled capsules because the procedure is less disruptive to the animals than gonadectomy followed by hormone injection. Capsules were prepared by cutting 20 mm lengths of silastic tubing (ID 0.04 in., Dow Corning), which were filled with EB in sesame oil (180 pg/ml, Krajnak et al., 1998). The ends of the capsules were sealed with medical adhesive (silicone Type A, Dow Corning). Animals were anesthetized with metofane, capsules were implanted at the nape of neck (so), and the incision closed with a wound clip. Ten days after 74 animals received the EB-capsule, they were perfused at either ZT 1, 5, 13, 17, or 20 (n= 6/time point except ZT 5; n=5). All animals were deeply anaesthetized with sodium pentobarbital and perfused transcardially with 0.01 M PBS (pH 7.2, 150-200 mllanimal) followed by 4% paraformaldehyde (Sigma) in 0.1 M phosphate buffer. Brains were post-fixed in paraformaldehyde for four hours, transferred to 20% sucrose in 0.1 M phosphate buffer, and then sectioned at 30 pm using a freezing microtome. Every third section from the medial septum to the VLH of each animal was processed for the immunocytochemical detection of PR as follows: free floating tissue was incubated in (1) 5% normal horse serum (NHS, Vector Laboratories) for one hour at room temperature (in PBS with 0.3% triton-X; PBS-TX), followed by (2) mouse anti-human PR primary antibody for 44 hours at 4°C (1 :5000, Chemicon lntemational, in PBS-TX and 3% NHS), followed by (3) biotinylated horse anti-mouse secondary antibody for one hour at room temperature (1:200, Vector Laboratories, in PBS- TX and 3% NHS), followed by (4) avidin-biotin complex for one hour at room temperature (in PBS-TX, Vectastain Elite Kit, Vector Laboratories). PR was visualized by incubating tissue in diaminobenzidine (0.5 mg/ml, in Trizma buffer, pH 7.2, Sigma) with 30% hydrogen peroxide (0.35 pl/ml buffer). A negative control was done by repeating the above procedures but omitting the PR antibody. Tissue was mounted, dehydrated, coverslipped, and examined under a light microscope (Laborlux S, Leitz Wetzlar GBH). Sections containing the VLH were photographed using a black and white closed-circuit digital camera (Cohu) 75 and captured using Scion Image (v. 4.02, Scion Corp.). A box 200 x 400 pm was placed over the VLH region and the number of PR+ cells was counted in each of 6 hemi-sections (Figure 4.1 ). Data from EB-injected animals in the first study were analyzed using a two-way ANOVA with ZT and EB treatment as the independent variables and the total number of PR+ cells as the dependent variable. Pairwise comparisons were computed with Tukey Least Significant Difference post hoc test (Keppel, 1982). Data from animals with EB-capsules in the second study were log-transformed to correct for normality and two outlier samples were dropped, as they were more than 2 standard deviations away from the mean of their group. These data were analyzed using an ANOVA with the corrected PR value as the dependent variable and ZT as the independent variable. Post-hoc painrvise comparisons were done using Fisher’s PLSD. The numbers of PR+ cells were also determined for the anteroventral portion of the periventricular nucleus (AVPV) of EB-injected animals because this region has been implicated in the regulation of the LH surge (Simerly, 1998; Weigand and Terasawa, 1982). For each animal, one section containing the AVPV was selected and bilateral counts were performed. An ocular grid was placed over the densest staining, aligned with the edge of the 3rd ventricle, and the number of PR+ cells was counted in a region 260 pm wide X 360 pm long. These data were analyzed using a two-way ANOVA with ZT and EB treatment as the independent variables and the total number of PR+ cells as the dependent variable. In all statistical analyses differences were considered significant when p<0.05 (Statview 5.0). 76 Figure 4.1. An example of PR+ cells in the ventrolateral hypothalamus in an estradiol-treated female grass rat. The box (200 x 400 um) in the figure indicates the region in which PR+ cells were counted. ARC = arcuate nucleus. 77 Results Sexual behavior In a typical mating sequence in grass rats the male approached the female and investigated her anogenital region. The female almost always moved away from the male after this contact and the male generally chased her in a tight circle for 1—2 seconds before copulation. Each mount was very brief lasting 1-2 seconds, and after the pair separated the male typically groomed his anogenital region. The female also groomed herself following most copulations and on many occasions she ran several centimeters away from the male before doing so. Grooming typically lasted about 1-2 seconds before the sequence of mating behaviors was repeated. Occasionally, following a copulation the pair would remain separated for several minutes before the male reinitiated contact with the female. Oil-treated control females always exhibited aggressive behaviors when the male approached (i.e. lunging, biting, rearing up) and no mating occurred in these pairs. In male and female grass rats several indices of sexual behavior varied significantly as a function of time of day. Female sexual responsiveness, as indicated by the LC, was affected by time (F: 6.186, df=3, p=0.001, Figure 4.2) and was significantly greater at ZT 23 than at either ZT 11 or 17 (p<0.05). When the LQs of individual females were plotted across the four time points the animals fell into two distinct groups (Figure 4.3). Approximately half (7/15) had a rhythm in sexual receptivity with a peak at ZT 23 and a trough at ZT 17 while the second 78 70' 50« 30‘ Mean (1SEM) 50- 30- . A ._l_ Lordosis quotient b C Copulations 5 11 17 23 Zeitgeber time of testing 90: 70- B _l_ Mounts 0 lights on I lights off 700- 300- 1004 500: D 5 11 17 23 Longest interval (sec) b 5 11 17 23 Zeitgeber time of testing Figure 4.2. Average rates of sexual behaviors (1 SEM) In 15 pairs of grass rats tested at 4 different times of day. Bars with different letters over them are significantly different from one another, p<0.05. Zeitgeber time 0=lights-on. 79 Flgur mour Figure 4.3. Lordosis quotient (number of lordosis responses/number of mounting attempts) for 15 individual grass rats. Animals were grouped into two types of sexual responsiveness on the basis of their behavior at ZT 17. Animals in the “low” group exhibited an L0 below 50% at ZT 17 (grey dots or bars); remaining individuals were placed in the “high” group (black dots or bars). Zeitgeber time 0=lights-on. A) LQ as a function of time for each individual. B) Average lordosis quotient (1 SEM) for the two groups of responders. 80 h .nluo .nH Hlow gh w Ihi Ilo E8030 £820. 0 9 22mm“ 8 E98: 2895. Zeitgeber time of testing Figure 4.3 81 group (8/15) displayed no rhythm at all and had high levels of receptivity at all timepoints. Male sexual behavior, indicated by the rate of mounting attempts, also varied significantly as a function of time (F: 3.97, df=3, p=0.01, Figure 4.2). Unlike female grass rats however, males had high levels of sexual behavior at both lights-on (ZT 23) and lights-off (ZT 11). The total number of copulations varied significantly as a function of time (F=4.023, df=3, p=0.012, Figure 4.2). The longest interval between copulations also varied across the 4 time points (F=5.58, df=3, p=0.002, Figure 4.2) and the duration of this interval was shorter when animals were tested at ZT 23 than when they were tested at ZT 17 (p<0.05). Progesterone receptors All EB-injected females and all females implanted with EB-filled capsules had distinct and darkly stained PR+ cells that were most evident in the arcuate nucleus, AVPV, and the VLH. No PR staining was observed in tissue incubated without the primary antibody. In contrast to EB-treated animals, virtually no PR+ cells were observed in control females. In the first study the number of PR+ cells present in the VLH was significantly affected by both time of day (F=6.86, df=1, p=0.02) and EB treatment (F=15.0, df=1, p=0.002), and there was a significant interaction between these two variables (F=8.77, df=1, p=0.012, Figure 4.4). Specifically, EB-injected animals sacrificed at ZT 13 had significantly more PR+ cells than did 82 ‘ Doontrol IEB 250 . 5200. U) +I 25,150 2 8100 + 1 a E a 50 . a i l 0 13 1'9 Zeitgeber time of injection Figure 4.4. Average number (1 SEM) of PR+ cells in the ventrolateral hypothalamus in female grass rats. Estradiol benzoate treated females received injections on days 1 and 2 and were sacrificed on day 3, at the same time. Bars with different letters over them are significantly different from one another, p <0.05. ZT 0 = lights-on. 83 any other group of animals (p<0.05). The number of PR+ cells in the VLH was similar, and low, in EB-injected females sacrificed at ZT 19 and control females sacrificed at ZT 13 and 19. In EB-injected animals, no significant effects of time, hormone treatment, or an interaction of these variables was found on the number of PR+ cells present in the AVPV. In the second study, the number of PR+ cells in the VLH of grass rats implanted with an EB-capsule also varied significantly as a function of time (F=3.41, df=3, p=0.02, Figure 4.5). Animals killed at ZT 20 had significantly fewer PR+ cells then did animals killed at ZT 13 or ZT 17 (p< 0.05). The drop in the number of PR+ cells from ZT 13 to ZT 20 mirrored that seen between ZT 13 and 19 in ovariectomized females injected with EB (compare Figure 4.4 and 4.5). Discussion These data show that grass rats have a rhythm in sexual behavior and suggest that it is due in part to changes in responsiveness of females to steroid hormones. Hormone-primed female grass rats exhibited the highest rates of sexual behaviors just before the lights came on (ZT 23, Figure 4.2, Figure 4.3) which is when intact grass rats mate (McElhinny, 1996; McElhinny et al., 1997). Mating may occur around the beginning of the active time of day because heightened arousal at this time maximizes the chances of locating a mate, and the animals may be better able to avoid predators at this time. The rhythm in behavioral responsiveness to hormones in female grass rats is presumably one factor regulating the timing of sexual activities. However, 84 \I O L on 9 U1 0 .p C? co (3 0' N O PR+ cells (x _+_SEM) .3 O 1 5 13 17 20 Zeitgeber time of perfusion Figure 4.5. Average number (1 SEM) of PR+ cells in the ventrolateral hypothalamus in female grass rats. Intact females were implanted with EB-containing silastic capsules. Bars with different letters over them are significantly different from one another, p <0.05. ZT 0 = lights-on 85 it is probable that the timing of hormone secretion is also important. Intact grass rats mate almost exclusively around the time of lights-on (Blanchong et al., 1999; McElhinny et al., 1997) yet females are capable of exhibiting sexual behavior at other times of day when given appropriate hormonal treatment (Figure 4.2, Figure 4.3). This suggests that patterns in hormone secretion as well as the changes in responsiveness to these hormones, documented here, must regulate the expression of female sexual behavior in intact grass rats. I did not address the hypothesis that changes in hormone secretion are related to sexual behavior in this species because these animals do not undergo predictable estrous cycles and there is no clear outward indicator of when an intact female is in estrus. Interestingly, the pattern of change in male sexual behavior did not parallel that of the females. Male sexual interest, as indicated by rates of mounting behavior, was highest when peaks in general activity are seen in grass rats in the lab, around lights-on (ZT 23) and lights-off (ZT 11) (McElhinny et al., 1997). Rates of mounting behavior were lowest at ZT 17, when these animals are usually inactive (Blanchong et al., 1999; Blanchong and Smale, 2000; Novak et al., 1999). By contrast, the rhythm in lordosis, indicative of female receptivity, had a single peak at ZT 23 (Figure 4.2). This suggests that the rhythm in sexual behavior seen in intact grass rats reflects rhythms of the female rather than the male. The daily rhythm in sexual behavior of male grass rats may be mediated by a rhythm in general arousal whereas the rhythm in female sexual behavior may be more tightly coupled to changes in responsiveness to, and secretion of, hormones. It is also important to consider that lordosis is a reflexive action 86 whereas mounting behavior may be indicative of sexual motivation. Had I examined a similar reflex in males, such as ejaculation, I may have seen a single daily peak around lights-on. Conversely, a measure of female sexual motivation might have revealed a rhythm that more closely paralleled the rhythm in general activity. The current data reveal considerable inter-individual variability with respect to responsiveness to hormones. Female grass rats administered identical doses of EB differed dramatically with respect to sexual receptivity, particularly at ZT 17, which was not the case at the other three time points examined. This raises the possibility that the hormone doses utilized in this study were close to a threshold level for the induction of sexual receptivity. Specifically, individuals with higher thresholds may have been unresponsive to hormone treatment at ZT 17, but as their behavioral responsiveness to progesterone changed across the day, the same hormone doses became sufficient to induce sexual receptivity. The L0 rhythm may have been more dramatic if all animals were treated with a lower concentration of hormones. Alternatively, these data might indicate individual variability with respect to whether or not animals have a rhythm in responsiveness to hormones. Although this study was conducted in a 12:12 Iight:dark cycle, it is likely that a rhythm of sexual behavior in grass rats would also be seen in constant conditions, as it is in nocturnal rodents (Alleva et al., 1971; Richter, 1970). In female lab rats and hamsters, rhythms in sexual receptivity and the timing of the LH surge are phase locked to the rhythm in activity onset and this temporal 87 relationship is maintained in animals free-running in constant conditions (Alleva et al., 1971; Fitzgerald and Zucker, 1976; Moline et al., 1981; Richter, 1970; Stetson and Gibson, 1977; Swann and Turek, 1982). The primary circadian clock in mammals, located in the suprachiasmatic nucleus (SCN), regulates the timing of mating behavior and estrous cyclicity in hamsters and lab rats. In these nocturnal rodents, SCN lesions eliminate estrous cyclicity, and behavioral rhythms in sensitivity to hormone treatment (Gray et al., 1978; Hansen et al., 1979; Hansen et al., 1978; Meyer-Bernstein et al., 1999). The SCN of grass rats and lab rats are similar with respect to rhythms in Fos expression, peptide distribution, and the effect of SCN lesions on locomotor rhythms (Katona et al., 1998; Mahoney et al., 2001; Nunez et al., 1999; Rose et al., 1999; Smale and Boverhof, 1999). Thus it is likely that the SCN of grass rats also contains a circadian clock, which regulates the timing of the rhythm in sexual behavior. The inverse relationship between rhythms in sexual behavior and in the number of PR+ cells in the VLH was surprising. EB-injected females sacrificed at ZT 19 had fewer PR+ cells than did EB treated females sacrificed at ZT 13, despite the fact that females treated with P at ZT 19 had a significantly higher LQ at ZT 23 than at any other time of day. This pattern was confirmed in animals with EB-capsules, who also had significantly fewer PR+ cells at ZT 20 than at ZT 13. In other rodents estradiol treatment increases cytosolic PRs and increases behavioral sensitivity to P (Blaustein and Feder, 1979b; Don Carlos et al., 1989; Fraile et al., 1987; Parsons et al., 1980; Parsons, Rainbow et al., 1981). Furthermore, in lab rats, guinea pigs, and hamsters, sexual receptivity is 88 positively correlated with the number of PR+ cells (Blaustein and Feder, 1979b; Delville and Blaustein, 1991; Don Carlos et al., 1989; Fraile et al., 1987; Parsons et al., 1980; Parsons, Rainbow et al., 1981). Grass rats thus appear to be different from these other species with respect to the temporal relationship between mating behavior and PRs. One interpretation is that, although I did not see an increase in PR+ cell numbers from ZT 20 to ZT 1 in EB-capsule treated female grass rats, PR numbers might rise from ZT 19/20 to 23, and peak around the time when females have maximal sexual receptivity. Indeed, in guinea pigs, nuclear PR levels reach their peak two to four hours after a subcutaneous P injection, and in lab rats nuclear PR levels rise within 30 minutes of an intravenous P injection (Blaustein and Feder, 1980; McGinnis et al., 1981). An alternative hypothesis is that females with high levels of PR at ZT 13 are hypersensitive to refractory effects of P on sexual behavior, which might be responsible for the decrease in sexual behavior at ZT 17. Refractory effects of P have been observed in other rodents at the end of a period of sexual receptivity (Blaustein and Feder, 1979a; Morin, 1977; Parsons, McGinnis et al., 1981; Zucker, 1968). This hypothesis is supported by the fact that grass rats with EB- capsules had high levels of PR+ cells at both ZT 13 and 17, perhaps reflecting a window of time during which P cannot induce sexual receptivity. Lastly, it is possible that changes in PR expression are not causally related to changes in sexual receptivity in this species. Animals in this study had a daily rhythm in estrogen-induced PR expression (Figure 4.4, Figure 4.5). To the best of my knowledge, this is the first 89 report of which I am aware of that describes a daily rhythm in PRs in any species; although PRs have been found to change in other rhythmic contexts. PR protein and mRNA levels in the hypothalamus fluctuate over the course of the estrous cycle of lab rats (Guerra-Araiza et al., 2000; Numan et al., 1999; Siegel et al., 1989; Simerly et al., 1996), and female Syrian hamsters housed in short days (10L:14D) have significantly fewer estradiol-induced PRs than do females housed in long days (14L:1OD; Mangels et al., 1998). These patterns of change in PR expression are likely to be caused in part by changes in secretion of steroids from the ovary. Changes in circulating steroids are unlikely to account for this rhythm in PRs reported here because hormone treatments were standardized. It is not currently clear what causes the rhythm in PRs in grass rats, or whether time of day affects PR+ cell number in other rodents. In summary, these findings demonstrate that diurnal grass rats have a rhythm in sexual behavior that is reversed from that of nocturnal lab rats, and that time of day influences PR expression of hormone-treated females. It will be interesting to determine whether, and how, these rhythms are functionally related, whether they are endogenous, and how the mechanisms underlying them differ between grass rats and nocturnal rodents. 90 Chapter 5 Conclusion Chapter summary Diurnal and nocturnal rodents differ dramatically with respect to rhythms in the timing of estrous-related events, but the mechanisms that underlie these rhythms are not known. In the second chapter I examined the hypothesis that differences in the timing of neuroendocrine events associated with estrous are I» ‘. ..‘nflJ-FILF due, in part, to rhythms in responsiveness to steroid hormones. I found that ,_ estradiol and progesterone treatment induced a rise in activity of gonadotropin releasing hormone-containing (GnRH+) neurons in both lab rats and grass rats. However, steroids were only able to increase GnRH+ cell activity at one time of day; in lab rats this was before lights-off, and in grass rats it was around lights- on. This rhythm in responsiveness to steroids appeared to be endogenous as it persisted in both species when they were housed in constant darkness for five days. In the third chapter I examined the hypothesis that differences in the timing of reproductive events in diurnal and nocturnal rodents are due to differences in the pathways or connections from the suprachiasmatic nucleus (SCN) to GnRH+ and estrogen receptor (ER)+ neurons. Using anterograde tract-tracing in grass rats, I found that the SCN projects to both ER+ and GnRH+ neurons, as is the case in lab rats and hamsters (de la lglesia et al., 1995; van der Beek et al., 1997). Furthermore, vasoactive intestinal polypeptide+ fibers 91 contacted about 34% of the GnRH+ neurons in grass rats and arginine vasopressin+ fibers contacted about 24% of these neuroendocrine cells. In the fourth chapter I determined that grass rats had a daily rhythm in sensitivity of sexual behavior to steroid hormones. Ovariectomized female grass rats were primed with steroid hormones, paired with an intact male, and tested for mating activity at four different times of day. The lordosis quotient and rate of copulation rose from zeitgeber time (ZT) 17 to ZT 23, whereas rates of mounting behavior were relatively high at both ZT 23 and ZT 11. The work presented in this dissertation indicates that differences in the timing of reproductive events in diurnal and nocturnal rodents are due, in part, to a reversal in rhythms in responsiveness to steroid hormones. What do these data mean? Where do I go from here? Extensive evidence indicates that in lab rats, both the circadian system and ovarian hormones regulate estrous cyclicity (Herbison, 1998). Experiments described in this dissertation indicate that the same is true for grass rats; steroid hormones and time of day interact to regulate rhythms in mating behavior and GnRH+ cell activity (Chapters 2, 4). Furthermore, the SCN sends direct projections to GnRH+ and ER+ neurons in grass rats, as is the case in lab rats (Chapter 3; van der Beek et al., 1997; Watson et al., 1995). These anatomical data indicate a possible pathway, common to both lab rats and grass rats, by which the circadian system might mediate the timing of reproductive events. Diurnal and nocturnal murid rodents, and perhaps mammals in general, may be similar with respect to the neuroendocrine mechanisms that regulate 92 reproductive functions such as hormone surges, copulatory behaviors and parturition. 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