Chronic predator stress in female mice reduces primordial follicle numbers: implications for the role of ghrelin

in Journal of Endocrinology
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  • 1 School of Health and Biomedical Sciences, RMIT University, Melbourne, Victoria, Australia

Correspondence should be addressed to L Sominsky: Luba.Sominsky@rmit.edu.au

Chronic stress is a known suppressor of female reproductive function. However, attempts to isolate single causal links between stress and reproductive dysfunction have not yet been successful due to their multi-faceted aetiologies. The gut-derived hormone ghrelin regulates stress and reproductive function and may therefore be pivotal in the neuroendocrine integration of the hypothalamic–pituitary–adrenal (HPA) and –gonadal (HPG) axes. Here, we hypothesised that chronic stress disrupts ovarian follicle maturation and that this effect is mediated by a stress-induced increase in acyl ghrelin and activation of the growth hormone secretatogue receptor (GHSR). We gave C57BL/6J female mice 30 min daily chronic predator stress for 4 weeks, or no stress, and gave them daily GHSR antagonist (d-Lys3-GHRP-6) or saline. Exposure to chronic predator stress reduced circulating corticosterone, elevated acyl ghrelin levels and led to significantly depleted primordial follicle numbers. GHSR antagonism stress-dependently altered the expression of genes regulating ovarian responsiveness to gonadotropins and was able to attenuate the stress-induced depletion of primordial follicles. These findings suggest that chronic stress-induced elevations of acyl ghrelin may be detrimental for ovarian follicle maturation.

Abstract

Chronic stress is a known suppressor of female reproductive function. However, attempts to isolate single causal links between stress and reproductive dysfunction have not yet been successful due to their multi-faceted aetiologies. The gut-derived hormone ghrelin regulates stress and reproductive function and may therefore be pivotal in the neuroendocrine integration of the hypothalamic–pituitary–adrenal (HPA) and –gonadal (HPG) axes. Here, we hypothesised that chronic stress disrupts ovarian follicle maturation and that this effect is mediated by a stress-induced increase in acyl ghrelin and activation of the growth hormone secretatogue receptor (GHSR). We gave C57BL/6J female mice 30 min daily chronic predator stress for 4 weeks, or no stress, and gave them daily GHSR antagonist (d-Lys3-GHRP-6) or saline. Exposure to chronic predator stress reduced circulating corticosterone, elevated acyl ghrelin levels and led to significantly depleted primordial follicle numbers. GHSR antagonism stress-dependently altered the expression of genes regulating ovarian responsiveness to gonadotropins and was able to attenuate the stress-induced depletion of primordial follicles. These findings suggest that chronic stress-induced elevations of acyl ghrelin may be detrimental for ovarian follicle maturation.

Introduction

Psychological stress has well-known inhibitory effects on reproductive function (Rivier & Rivest 1991, Tilbrook et al. 2002, Young et al. 2006, Lynch et al. 2014), suppressing hypothalamic–pituitary–gonadal (HPG) axis activity at every level. At the hypothalamus, stress-induced release of corticotropin-releasing hormone (CRH) and activation of γ-aminobutyric acid (GABA)-ergic signalling mediate the suppression of gonadotropin-releasing hormone (GnRH) pulsatility (Williams et al. 1990, Rivier & Rivest 1991, Han et al. 2002, Lin et al. 2012, Michopoulos et al. 2013), leading to the inhibition of luteinising hormone (LH) release from the pituitary (Tilbrook et al. 1999, Li et al. 2005, 2015) and disruption of oestrous cyclicity (Mourlon et al. 2011, Divyashree & Yajurvedi 2016). This decrease in pituitary gonadotropins reduces ovulatory capacity and gonadal steroid production (Macfarlane et al. 2000, Whirledge & Cidlowski 2010). Likewise, stress-induced catecholaminergic signalling directly affects the ovary, contributing to the development of ovarian cysts and disrupting ovarian follicle maturation (Paredes et al. 1998, Dorfman et al. 2003, Bernuci et al. 2008).

We, and others, have recently identified a key player in mediating these neuroendocrine effects of stress on reproductive function: ghrelin (reviewed in Sominsky et al. 2017). Ghrelin is a 28 amino acid peptide that exists in circulation in its acylated and des-acylated forms (Kojima et al. 1999). While des-acyl ghrelin represents approximately 90% of total ghrelin in circulation, and has independent physiological functionality, its receptor is currently unidentified (Kojima & Kangawa 2005). Acyl ghrelin, on the other hand, is known to act via the growth hormone secretagogue receptor (GHSR) and has a variety of functions, including modulation of the neuroendocrine response to stress, which at least in part occurs via activation of GHSR-expressing cells in the anterior pituitary (Spencer et al. 2012).

Within the hypothalamus, acyl ghrelin indirectly suppresses GnRH pulsatility in the medial preoptic area and directly modulates gonadotropin release from the anterior pituitary (Fernandez-Fernandez et al. 2005, 2007, Smith et al. 2013). Ghrelin and GHSR are also expressed in the ovary and endometrium (Gualillo et al. 2001, Caminos et al. 2003, Gaytan et al. 2003, 2005, Miller et al. 2005). Acyl ghrelin’s stimulation of human granulosa-lutein cells in vitro, dose-dependently suppresses steroidogenesis, and these effects are reversed by the specific antagonism of the 1a, but not 1b isoform of GHSR (Viani et al. 2008). Ghrelin-mediated suppression of steroidogenesis has also been demonstrated in vitro in porcine ovarian follicles obtained from mature pigs (Rak-Mardyla et al. 2015). In contrast, in prepubertal porcine follicles, in vitro supplementation with ghrelin is able to stimulate steroidogenesis and proliferation, while inhibiting apoptosis (Rak & Gregoraszczuk 2008, Sirotkin & Meszarosova 2010). Interestingly, in vitro stimulation of prepubertal porcine granulosa cells with a GHSR antagonist, d-Lys3, similarly promotes steroidogenesis and proliferation and suppresses the expression of apoptotic markers (Sirotkin et al. 2011). In peripubertal rats, however, chronic in vivo treatment with acyl or des-acyl ghrelin or their combination has been shown to disrupt ovarian follicle development (Kheradmand et al. 2009, Roa et al. 2010). These data suggest a combination of GHSR and non-GHSR-dependent pathways may mediate the effects of ghrelin on the maturation of ovarian follicles, either directly at the level of the ovary or indirectly via the other components of the HPG axis, with a potential age and species specificity.

Acyl ghrelin levels rise in response to stress and GHSR antagonism is showing promising outcomes in attenuating the metabolic and behavioural consequences of chronic stress (Ochi et al. 2008, Kodomari et al. 2009, Patterson et al. 2013a, Meyer et al. 2014, Spencer et al. 2015, Sominsky et al. 2017, Yousufzai et al. 2018). Given the regulatory role of acyl ghrelin in stress responsivity and in reproductive function, here we hypothesised that by blocking the activity of GHSR-mediated pathways during chronic stress we would be able to normalise indices of female reproductive function. While the effects of stress on the hypothalamic–pituitary components of the HPG axis, and the role of acyl ghrelin in the regulation of these areas are well researched, evidence regarding the effects of stress and ghrelin on the ovary, the primary functional unit of female fertility, is still scarce. We therefore particularly focused on examining the effects of chronic stress and GHSR antagonism on the ovary.

Materials and methods

Animals

C57BL/6J female mice (8 weeks old) and Wistar male rats (10–12 weeks old) were obtained from the Animal Resources Centre, WA, Australia. Upon arrival at the RMIT Animal Facility, we maintained all the animals at 22°C on a 12 h light/darkness cycle (07:00–19:00 h) with pelleted chow (4.8% fat, 3.34 kcal/g) and water available ad libitum. All procedures were conducted in accordance with the National Health and Medical Research Council Australia Code of Practice for the Care of Experimental Animals and were approved by the RMIT Animal Ethics Committee.

Ghrelin antagonist

d-Lys3-GHRP-6 (d-Lys3, Sigma-Aldrich) was prepared and administered as previously described (Meyer et al. 2014). Briefly, d-Lys3 was diluted to 2.74 μg/mL in 0.9% saline and administered at 1 mL/kg intraperitoneally (i.p., 0.00274 mg/kg BW) 30 min before the start of the stress protocol. d-Lys3 is a potent GHSR1a antagonist that also weakly binds to melanocortin receptors (Ki = 26–120 μM). However, at this dilute dose, it is not expected to interact with these receptors (Meyer et al. 2014).

Stress protocol

Mice were randomly assigned to one of four treatment groups: n = 8 per group: No stress, saline; no stress, d-Lys3; stress, saline; stress, d-Lys3. Sixteen mice allocated to the stress groups were singly-housed and 16 non-stressed mice were housed in groups of four. All rats used to impose the predator stress were singly-housed in a separate holding room. At 30 min before the beginning of the stress protocol we gave all mice d-Lys3 or saline, as described above. Non-stressed mice were returned to their home cages and left otherwise undisturbed. Chronic predator stress was conducted as in (Barnum et al. 2012, Burgado et al. 2014) with slight modifications. Each mouse was individually locked inside a clear plastic rodent exercise ball (Critter Roller Small 14 cm diameter, Pet One, Ingleburn, NSW, Australia) and the ball placed in the centre of the rat home cage for 30 min. This procedure was repeated daily for 4 weeks (Fig. 1A), at a different time of each day during the light phase of the diurnal cycle (between 09:00 and 15:00 h) to minimise predictability. During the stress session, mice were exposed to the sight, sound and smell of the rat through the openings in the exercise ball, but the rat could not make direct physical contact with the mouse. To avoid familiarity and possible habituation, chronically stressed mice were paired with a different rat for each stress session, rotated each day for 8 days and then beginning again. All mice were weekly weighed.

Figure 1
Figure 1

Effects of chronic stress and GHSR antagonism on oestrous cycle, body weight, circulating corticosterone and acyl ghrelin. (A) Study design. Female mice were treated with d-Lys3 (2.74 µg/kg, i.p.) or saline, and 30 min later were subjected to 30 min chronic predator stress or left undisturbed in their home cage. This procedure was repeated daily for 4 weeks. (B) Chronic stress and GHSR antagonism did not affect the regularity of oestrous cyclicity. (C) Exposure to chronic stress caused a significant reduction in body weight by two weeks after the commencement of stress, while d-Lys3 treatment significantly increased weight gain at the beginning of the second and third experimental weeks. (D) Chronic predator stress significantly reduced concentrations of circulating corticosterone, and (E) significantly increased the levels of acyl ghrelin. (F) Chronic stress increases the release of growth hormone (GH) in saline-treated but not d-Lys3-treated mice. Data are mean ± s.e.m. Parentheses and $ represent main effect of stress; Parentheses and # represent an effect of drug. Connecting lines with * represent post hoc differences. $,#,*P < 0.05.

Citation: Journal of Endocrinology 241, 3; 10.1530/JOE-19-0109

Oestrous cyclicity assessment

During the last week of the experiment we assessed oestrous cyclicity in all mice. We determined the stage of the oestrous cycle (proestrus, oestrus, metestrus and diestrus) by visual observation of the vaginal opening that changes in its appearance at different stages of the cycle and by the examination of vaginal cytology (Byers et al. 2012). For the assessment of vaginal cytology, a few drops of 0.9% NaCl were inserted into the vaginal cavity using a Pasteur pipette, withdrawn and smeared on a glass slide. Proestrous was indicated by the increased presence of nucleated epithelial cells; oestrous was determined by the presence of cornified epithelial cells; metestrous was indicated by a mix of cell types, including leukocytes, nucleated epithelial and/or cornified epithelial cells; diestrus was determined when the smear consisted of predominantly leukocytes (Felicio et al. 1984). If no oestrous was detected or if a mouse spent more than 50% of the time in diestrous or oestrous, we considered this as irregular (Li et al. 2017). It is important to note that due to individual variability in cycle length, a minimum of 2–3 weeks of daily vaginal smears is typically required to determine cycle regularity (Caligioni 2009). However, while minimally invasive, vaginal smearing imposes an additional minor stress on the animal. Although this additional stress may not be an issue for chronically stressed mice, we tried to avoid any possible interference with the stress responses of our mice in the non-stressed groups. We therefore used these observations primarily to allow us to terminate the experiment at the same stage of the oestrous cycle (non-ovulatory stage of the cycle, diestrous or metestrous). Mice were killed by an overdose of sodium pentobarbital (Lethabarb; ~100 mg/kg i.p.) between 10:00 to 12:00 h. Prior to killing, all mice were fasted for 2 h to standardise satiety levels without inducing negative energy balance.

Assessment of circulating corticosterone and ghrelin

Upon killing, we collected cardiac blood from all mice. Blood was allowed to clot for 30 min then centrifuged at 1000 × centrifugal force (g) for 15 min at 4 ± 2°C. Serum (top layer) was collected and kept at −20°C until processed.

We measured serum corticosterone concentrations using a standard rat corticosterone ELISA (Abnova Corp., Taipei, Taiwan). The intra-assay variability for this assay was 4.8% coefficient of variation (CV) and lower limit of detection, 40 pg/mL. We assayed samples from all treatment groups together in duplicate.

For acyl ghrelin assessment, we processed the blood samples according to the ELISA manufacturer’s instructions (Merck Millipore). Thus, 250 µL of whole blood was treated with Pefabloc (1 mg/mL final concentration) in a tube that contained no anticoagulant. Blood was allowed to clot for 30 min then centrifuged at 2500 g for 15 min at 4 ± 2°C. 0.5 M HCl (final concentration 0.05 M HCl) was added to the serum and samples stored at −20°C until processed. Intra-assay variability was 0.3–7% CV and lower limit of detection, 0.8 pg/mL. We assayed samples from all treatment groups together in duplicate.

Pituitary hormone assessment

We used a Milliplex Mouse Pituitary Magnetic Bead Panel (Millipore) and a Bio-Plex MAGPIX instrument (Bio-Rad) to measure the levels of circulating LH, follicle-stimulating hormone (FSH) and growth hormone, according to the manufacturer’s instructions. The intra-assay variability for this assay was <15% CV, and lower limit of detection, 1.9 pg/mL, 9.5 pg/mL and 1.7 pg/mL, for LH, FSH and growth hormone, respectively.

Oestrous cycle stage, time of day and satiety status were all controlled for to standardise the sample collection procedure.

Ovarian tissue collection

We collected ovaries from all mice for morphometric and molecular analysis. One ovary from each animal was snap frozen in liquid nitrogen and stored at −80°C for gene expression analysis, and one ovary was fixed in Bouin’s solution (Sigma-Aldrich) overnight, rinsed four times in 70% ethanol and stored in ethanol until processing.

RNA isolation, reverse transcription and quantitative real-time PCR (qRT-PCR)

We isolated total RNA using QIAzol reagent and RNeasy Mini Kits (Qiagen). RNA concentrations were determined using spectrophotometer, NanoDrop 2000/2000c (Thermo Fisher Scientific). We reverse-transcribed 1 µg of RNA to cDNA using a QuantiTect Reverse Transcription kit (Qiagen) with the elimination of genomic DNA, according to the manufacturer’s instructions. To assess the presence of genomic DNA contamination, non-template controls were generated using 500 ng of RNA by omitting the reverse transcriptase from the cDNA reaction.

qRT-PCR was completed for the assessment of Casp3, Bax, Bcl2, Bmp15, Gdf9, Foxo3 and Amh mRNA expression, using TaqMan Gene Expression Assays (Thermo Fisher Scientific) and amplified on the QuantStudio 7 Flex Real-Time PCR System (Applied Biosystems). Specific primer details are shown in Table 1. A relative quantitative measure of the target gene expression, compared with an endogenous control Actb mRNA expression, was completed using the equation 2−ΔΔC (t) (Livak & Schmittgen 2001), where C(t) is the threshold cycle at which fluorescence is first detected significantly above background. Data are presented as a fold change relative to non-stressed saline-treated mice.

Table 1

TaqMan probe details for qRT-PCR.

Target geneNCBI reference sequenceTaqMan assay IDProduct size
ActbNM_007393Mm00607939_s191
Casp3NM_001284409.1Mm01195085_m170
BaxNM_007527.3Mm00432051_m184
Bcl2NM_009741.5Mm00477631_m185
Bmp15NM_009757.5Mm00437797_m164
Gdf9NM_008110.2Mm00433565_m160
Foxo3NM_019740.2Mm01185722_m192
AmhNM_007445.2Mm00431795_g160

Real-time quantitative PCR array

We also used a Custom RT2 PCR array (Qiagen) designed to examine the gene expression of Crhr1, Crhr2, Nr3c1, Nr3c2, Ghsr, Mboat4, Ghrl, Lhcgr, Fshr, Inhba, Inhbb, Nppc (Table 2 for primer details). For this array, we transcribed 400 ng of total RNA, extracted as detailed above, to cDNA using the RT2 First Strand Kit (Qiagen), according to the manufacturer’s instructions. Samples were then diluted in RT2 SYBR Green Mastermix, loaded onto 384-well PCR array plates and amplified on the QuantStudio 7 Flex Real-Time PCR System (Applied Biosystems), including an initial activation step at 95°C for 10 min followed by 40 cycles at 95°C for 15 s and 60°C for 1 min. Actb and Gapdh were used as endogenous controls. The C(t) values for these genes were averaged and used for the comparative threshold cycle (ΔΔ C(t)) calculations, where C(t) is ≤40. Fold changes were then calculated using the 2−ΔΔC (t) equation (Livak & Schmittgen 2001).

Table 2

PCR array primer details.

Target geneNCBI reference sequenceTaqMan assay ID
ActbNM_007393PPM02945B
Crhr1NM_007762PPM04312A
Crhr2NM_009953PPM04313A
Nr3c1NM_008173PPM05371F
Nr3c2NM_001083906PPM06002A
GhsrNM_177330PPM05304A
Mboat4NM_001126314PPM40440B
GhrlNM_021488PPM31564C
LhcgrNM_013582PPM04900A
FshrNM_013523PPM29591A
InhbaNM_008380PPM02994B
InhbbNM_008381PPM02995A
NppcNM_010933PPM25709A

Ovarian morphometry

As previously described (Sominsky et al. 2012, 2016, Fuller et al. 2017), fixed ovaries were dehydrated, embedded in paraffin and sectioned at 4 μm. We used principles in accordance with design-based stereology as far as possible, with randomised serial sampling of ovarian sections, using the physical dissector method (Myers et al. 2004). Thus, five consecutive sections were mounted on each slide, and every fourth slide was H&E stained and counted throughout 400 µm of the ovary, from a random start, beginning at approximately the middle of the ovary. Total counts were carried out on every first, third and fifth section of each of five H&E slides, with the first and fifth sections acting similar to ‘lookup’ sections as described in (Myers et al. 2004). For all healthy follicle populations, only follicles with visible oocyte nuclei were counted, with each follicle counted only once on each H&E slide, even if appearing in more than one section. Follicles were classified as (a) ‘primordial’: an oocyte surrounded by a single layer of flattened pregranulosa cells; (b) ‘early primary’: an oocyte surrounded by a single layer of flattened pregranulosa cells with at least two cuboidal granulosa cells; (c) ‘primary’: an oocyte surrounded by cuboidal granulosa cells; (d) ‘preantral’: follicles with no antral cavity and two or more layers of cuboidal granulosa cells; (e) ‘antral’: an antral cavity is visible, with at least two layers of cuboidal granulosa cells; or (f) ‘preovulatory’: the largest follicles with a cumulus granulosa cell layer (Sobinoff et al. 2012, Camlin et al. 2016). Follicle counts were performed by an experimenter blinded to the experimental conditions. We also assessed the presence of atretic follicles, characterised by oocyte degeneration and/or granulosa cell degeneration, disorganisation and/or the appearance of pyknosis (Pangas et al. 2002, Tomic et al. 2004). Data are shown as an average number of follicles normalised per mm2 of ovary area, as in (Goldman et al. 2017), there being no differences between the groups in ovary area. Raw follicle counts are reported in Table 3.

Table 3

Raw follicle counts.

Follicle stageNs SalNs d-Lys3Stress SalStress d-Lys3Statistics
Primordial82.6 ± 16.762.4 ± 10.730.5 ± 5.567.8 ± 19.6$ x #
Early primary50.6 ± 7.136.9 ± 5.832.5 ± 2.839.7 ± 9.0ns
Primary15.6 ± 3.314.1 ± 1.610.5 ± 2.217.2 ± 3.3ns
Secondary21.5 ± 2.919.4 ± 1.517.2 ± 2.625.3 ± 3.7ns
Antral3.1 ± 0.53.1 ± 0.83.0 ± 1.34.8 ± 1.5ns
Ovulatory2.1 ± 0.41.1 ± 0.32.2 ± 0.72.0 ± 0.6ns
Atretic15.7 ± 2 14.6 ± 1.917.8 ± 2.516.5 ± 3.5ns

N = 6–8. Data are mean ± s.e.m. $ × # represents significant stress × drug interaction; P < 0.05.

ns, not significant.

Ovarian volume

We used the Cavalieri point-counting principle to estimate the volumes of the ovarian cortex, medulla and corpora lutea, as previously described (Karbalay-Doust & Noorafshan 2012, Guven et al. 2013). Briefly, we used ImageJ (National Institutes of Health) to randomly superimpose a grid of systematic uniform test points representing an area of 0.01 mm2 apart. The number of points hitting each area of interest were counted and the volumes of the structures of interest were estimated using the following equation: volume (V) = t × a(p) × (ΣP), where V is the volume of interest; t is section thickness, a(p) is the interpoint area and ΣP is the number points hitting the region of interest in that section. The proportion of each ovarian division of the total ovarian volume is presented in Table 4.

Table 4

Mean proportion of ovarian divisions of the total ovarian volume (%) in non-stressed and chronically stressed saline and d-Lys3-treated mice.

Ovarian divisionsNs SalNs d-Lys3Stress SalStress d-Lys3Statistics
Cortex68.6 ± 2.961.8 ± 2.966.3 ± 4.369.5 ± 1.7ns
Medulla19.9 ± 0.922.7 ± 1.120.1 ± 1.120.1 ± 2.1ns
Corpus luteum11.5 ± 2.715.5 ± 2.913.6 ± 4.010.4 ± 2.7ns

N = 6–8. Data are mean ± s.e.m.

ns, not significant.

Follicle and oocyte measurements

We determined the follicle and oocyte diameters by using ImageJ, as previously described (Griffin et al. 2006, Sominsky et al. 2016). Briefly, we obtained two measurements at the widest cross-section of each follicle/oocyte. Follicles were measured from the outer layer of the theca cells, if present, or from the outer layer of the granulosa cells. Oocyte measurements were obtained including the zona pellucida, when present.

Immunohistochemistry

Immunohistochemistry was used to visualise Ki-67, a marker of cell proliferation expressed at all stages of the cell cycle (Gerdes et al. 1984), and anti-Müllerian hormone (AMH), a hormone that reduces follicle sensitivity to FSH and inhibits primordial follicle recruitment into the growing pool (Durlinger et al. 1999, 2002). Antigen retrieval was carried out by microwaving sections in sodium citrate buffer at 800 W for 15 min (10 mM sodium citrate, pH = 6), and slides were cooled down to room temperature (RT). For Ki-67 detection, sections were blocked in 10% goat serum/1% Triton X-100/phosphate-buffered saline (PBS) for 1 h at RT. Slides were then incubated overnight at 4°C with Ki-67 antibody (1:200; rabbit polyclonal; Ab15580; Abcam), washed in PBS/0.1% Triton X-100 and incubated with Alexa-Fluor 488 fluorescent-conjugated secondary antibody (1:500; Thermo Fisher Scientific) for 1 h at RT. For AMH detection, slides were blocked in 3% BSA in PBS for 90 min at RT. Sections were then incubated overnight at 4°C with AMH (1:20; mouse monoclonal; MCA2246; Bio-Rad). Slides were then washed in PBS/0.1% Triton X-100 and incubated with Alexa-Fluor 594 fluorescent-conjugated secondary antibody (1:200, Thermo Fisher Scientific) for 1 h, RT. We counterstained the sections with 4’,6-diamidino-2-phenylindole (DAPI) using Fluoroshield with DAPI mounting medium (Sigma-Aldrich), coverslipped them and viewed them under an Olympus BX61 fluorescent microscope fitted with a Nikon DS-Ri2 camera.

The antibodies used in our study have been previously validated by the manufacturers and used by other researchers. Thus, the Ki-67 antibody is batch-tested for immunohistochemistry applications by the manufacturer and has been used by others as a marker of granulosa cell proliferation in the mouse ovary (Meinsohn et al. 2018, Sun et al. 2018). The AMH antibody has been validated for use in immunohistochemistry in paraffin-embedded tissues by the manufacturer and has been used by others for immunolocalisation in female mice (Kevenaar et al. 2006, Camlin et al. 2016). Negative controls omitting the primary antibody were included with each run.

Follicles were considered proliferating if they included a visible oocyte nucleus and at least one Ki-67-positive granulosa cell (Dolmans et al. 2007). Ki-67 expression was assessed in granulosa cells from primary, secondary and antral follicles. To analyse the expression of AMH, we assessed fluorescence intensity in AMH-positive secondary and antral follicles with a visible oocyte nucleus using ImageJ (National Institutes of Health). Mean fluorescence intensity was calculated using a corrected total cell fluorescence (CTCF) formula (CTCF = integrated density − (area of selected section × mean fluorescence of background readings)), as previously described (McCloy et al. 2014). Data are normalised to the mean fluorescence intensity of the control group and expressed in arbitrary units (AU), as we and others have previously described in 2–4 randomly selected non-consecutive sections per animal (Camlin et al. 2016, Fuller et al. 2017, Mihalas et al. 2017).

For the detection of follicular apoptosis, we de-paraffinised and rehydrated paraffin-embedded sections, as above. We pre-treated the sections with 20 µg/mL proteinase K for 15 min at RT and performed analysis for terminal deoxynucleotidyl transferase dUTP nick-end labelling (TUNEL) using ApopTag Fluorescein In Situ Apoptosis Detection Kit (Merck Millipore), according to the manufacturer’s instructions. We then counterstained the sections using Fluoroshield with DAPI mounting medium (Sigma-Aldrich). A positive control included sections treated with DNAse I (Qiagen) to induce nonspecific DNA fragmentation, and negative control staining was conducted without active TdT but including proteinase K digestion, to control for nonspecific incorporation of nucleotides. Slides were viewed under fluorescent microscope and follicles were classified as apoptotic if they contained a TUNEL-positive oocyte and/or ≥4 TUNEL-positive granulosa cells for primary, secondary and antral follicles or >1 TUNEL-positive granulosa cells for primordial follicles (Almog et al. 2001, Roti et al. 2012, Sominsky et al. 2016). TUNEL-positive granulosa cells were also primarily expressed and assessed in large follicle populations (Tingen et al. 2009, Liew et al. 2017).

Statistical analyses

Data were analysed by two-way analyses of variance (ANOVA) with Tukey post hoc tests, where significant interactions were found. We used repeated-measures ANOVAs to analyse body weight changes, with stress and d-Lys3 treatment as between factors and time as the repeated measure. We also conducted a chi-square test for the assessment of regularity of oestrous cyclicity. Data are presented as the mean ± s.e.m. Statistical significance was assumed when P < 0.05.

Results

Chronic predator stress reduces body weight and corticosterone, elevating acyl ghrelin

Chronic stress did not significantly affect the regularity of oestrous cyclicity during the 4–6 days of observation, with the incidence of regular cycle in non-stressed saline and d-Lys3-treated mice ranging between 62.5% (five out of eight mice) to 87.5% (seven out of eight mice) respectively, compared to 42.9% regular cyclicity in stressed saline (three out of seven mice), and 57.1% regular cyclicity in stressed d-Lys3-treated mice (four out of seven mice); (Fig. 1B; n = 7–8).

Chronic stress typically reduces body weights in rodents (Harris 2015). We therefore assessed changes in body weight during the first 3 weeks of the chronic stress protocol, until the commencement of daily vaginal smears. There were significant week × stress (F(2,50) = 3.88, P = 0.027) and week × drug (F(2,50) = 5.02, P = 0.010) interactions (Fig. 1C; n = 6–8). Post hoc tests revealed that stress caused a reduction in body weight by two weeks after the commencement of the experiment (F(1,25) = 7.19, P = 0.013), suggestive of the effectiveness of the stress paradigm, and d-Lys3 significantly increased weight gain at the beginning of the second and third experimental weeks (F(1,25) = 5.75, P = 0.024; F1,25 = 9.85, P = 0.004).

Four weeks of chronic predator stress led to a significant reduction in circulating corticosterone concentrations (F(1,25) = 6.54, P = 0.017; Fig. 1D; n = 6–8), reflective of a potential stress sensitisation that has been previously reported in similar stress models of predator exposure and is associated with poor stress reactivity (Herman 2013, Whitaker & Gilpin 2015). There was no effect of d-Lys3 treatment. As anticipated, acyl ghrelin levels were significantly increased in chronically stressed mice (F(1,22) = 7.74, P = 0.011; Fig. 1E; n = 4–8), with, again, no effect of d-Lys3.

GHSR mediates the growth hormone-releasing effects of acyl ghrelin (Kojima et al. 1999), and d-Lys3 has been shown to antagonise growth hormone-releasing properties of GHSR (Pinilla et al. 2003). Thus, we assessed the levels of circulating growth hormone levels. There was a significant stress × drug interaction (F(1,23) = 5.13, P = 0.033; Fig. 1F; n = 6–7), and post hoc analyses revealed growth hormone levels were significantly increased by stress in saline-treated mice, corresponding with increased acyl ghrelin levels in these animals. In d-Lys3-treated mice, despite a similar increase in acyl ghrelin, there was no increase in growth hormone levels.

Ovarian expression of stress and ghrelin regulatory genes is unchanged with chronic stress

CRH, the main regulating hormone of the HPA axis, is also produced in the ovary (Mastorakos et al. 1993), and ovarian CRH receptors have been suggested to mediate ovulation and steroidogenesis (Wypior et al. 2011). Likewise, glucocorticoid receptors, important in the HPA axis-negative feedback, are also essential for mediating the effects of glucocorticoids on ovarian steroidogenesis (Tetsuka et al. 1999). Despite 4 weeks of predator stress, there were no differences in the gene expression of CRH or glucocorticoid receptors in mouse ovaries between any of our groups (Fig. 2A, B, C and D; n = 4–8). Despite stress-induced differences in circulating ghrelin (Fig. 1D), there were also no changes in the expression of Ghrl, Mboat4 and Ghsr mRNA (Fig. 2E, F and G; n = 4–8).

Figure 2
Figure 2

Ovarian expression of stress and ghrelin regulatory genes is unchanged. Chronic stress and GHSR antagonism did not affect ovarian gene expression of (A) corticotropin-releasing hormone receptor 1 (Crhr1), (B) corticotropin-releasing hormone receptor 2 (Crhr2), (C) glucocorticoid receptor (Nr3c1), (D) mineralocorticoid receptor (Nr3c2), (E) ghrelin (Ghrl), (F) ghrelin O-acyltransferase (Mboat4) and (G) growth hormone secretagogue receptor Ghsr. Data are mean ± s.e.m.

Citation: Journal of Endocrinology 241, 3; 10.1530/JOE-19-0109

Chronic stress and GHSR antagonism influence the expression of genes regulating ovarian responsiveness to FSH

Chronic stress and changes in ghrelin signalling have been previously shown to affect gonadotropin release and ovarian follicle development (reviewed in Sominsky et al. 2017). We therefore next assessed whether chronic stress and GHSR antagonism affected the release of gonadotropins, the expression of genes regulating ovarian sensitivity to gonadotropins and the potential for follicle proliferation. There were no differences in the levels of circulating LH or the expression of its receptor in the ovary (Fig. 3A and B; n = 5–8). While there were also no differences in the levels of circulating FSH (Fig. 3C; n = 6–8), there was a significant stress × drug interaction affecting the expression of the Fshr gene (F(1,21) = 9.81, P = 0.005; Fig. 3D; n = 5–8), with no post hoc differences, suggesting that d-Lys3 may increase the ovary’s capacity to respond to FSH under non-stressed conditions, but reduce this after chronic stress.

Figure 3
Figure 3

Chronic stress and GHSR antagonism influence the expression of genes regulating ovarian responsiveness to FSH. Chronic stress and GHSR antagonism had no effect on (A) circulating luteinising hormone (LH), (B) gene expression of LH receptor (Lhcgr) and (C) circulating follicle-stimulating hormone (FSH). (D) Gene expression of ovarian FSH receptor is affected by chronic stress and GHSR antagonism. (E) There were no changes in the gene expression of inhibin βA subunit (Inhba), but (F) an effect of chronic stress and GHSR antagonism on the expression of inhibin βB subunit (Inhbb). (G) The expression of the expression of Nppc, a gene encoding for CNP was significantly reduced in d-Lys3-treated mice. (H) There were no differences in the percentage of Ki-67-positive proliferating follicles. Data are mean ± s.e.m. Parentheses and $ × # represent a significant stress × drug interaction with no post hoc differences; parentheses and # represent main effect of drug. $,#P < 0.05.

Citation: Journal of Endocrinology 241, 3; 10.1530/JOE-19-0109

Inhibin A and inhibin B are produced by antral follicles and are stimulated by the secondary surge of FSH during the oestrous cycle, with different secretion patterns (Welt and Schneyer 2001, Kenny & Woodruff 2006). Both are important in regulating granulosa cell Fshr expression (Lu et al. 2009). There were no significant differences in the expression of Inhba (Fig. 3E). However, there was a significant stress × drug interaction on the expression of Inhbb (F(1,21) = 4.17, P = 0.054; Fig. 3F; n = 5–8), consistent with the above effects of stress and d-Lys3 on the expression of Fshr, and hence, a potential increase in ovarian sensitivity to FSH. We also saw a significant d-Lys3 treatment-induced reduction in the expression of Nppc, a gene encoding for C-type natriuretic peptide (CNP) that is produced by granulosa cells of secondary and antral follicles in response to FSH stimulation, inducing follicle maturation, while also maintaining oocyte meiotic arrest (reviewed in Hsueh et al. 2015) (F(1,21) = 12.68, P = 0.002; Fig. 3G; n = 6–7). This latter finding indicates that GHSR antagonism may lead to precocious resumption of meiosis.

Despite these indications of chronic stress and d-Lys3 exposure altering the capacity for follicle maturation, follicle proliferation was not likely affected by stress and GHSR antagonism. There were no differences in the immunohistochemical expression of Ki-67 (Fig. 3H, n = 5–8), a marker of cell proliferation expressed primarily in the granulosa cells of secondary and antral follicles in adult ovary (Scalercio et al. 2015).

Chronic stress and GHSR antagonism do not affect ovarian apoptosis

Given the potential for changes in FSH sensitivity to alter the development of large follicles and to affect apoptosis, we assessed the expression of ovarian apoptotic markers. There were no differences in the expression of Bax, Bcl or Casp3 mRNA, nor were there differences in the percentage of TUNEL-positive follicles between the groups (Fig. 4A, B, C, D and E; n = 5–8).

Figure 4
Figure 4

Chronic stress and GHSR antagonism had no effect on ovarian apoptosis. There were no changes in gene expression of apoptosis regulators (A) Bax, (B) Bcl2 and (C) caspase 3 (Casp3). (D) There were no differences in the percentage of TUNEL-positive follicles. (E) Representative images of TUNEL (green). Scale bars = 50 µm. Data are mean ± s.e.m. A full colour version of this figure is available at https://doi.org/10.1530/JOE-19-0109.

Citation: Journal of Endocrinology 241, 3; 10.1530/JOE-19-0109

Chronic stress induces a decline in the number of primordial follicles, partially rescued by GHSR antagonism

Since primordial and primary follicle death is most accurately estimated by follicle counts (Tingen et al. 2009, Liew et al. 2017), we also performed the assessment of follicle numbers. We saw a significant stress × drug interaction affecting the number of primordial follicles (F(1,24) = 4.84, P = 0.036; Fig. 5A; n = 6–8). Post hoc analyses indicated chronic stress significantly reduced primordial follicles in saline-, but not d-Lys3-treated mice. There was also a tendency for a reduction in the number of early primary follicles, a transitional primordial follicle population, in the ovaries of stressed mice. This occurred irrespective of drug treatment (F(1,24) = 3.95, P = 0.059; Fig. 5B; n = 6–8). There were no differences in the numbers of primary follicles (Fig. 5C; n = 6–8). Consistent with the lack of an effect of stress or d-Lys3 on apoptotic markers primarily expressed in larger follicle populations, there were also no differences in the numbers of ovulatory, antral and secondary follicles (Fig. 5D, E and F) or in follicle atresia (Fig. 5G), which was restricted to the populations of secondary and antral follicles, as previously described (Tingen et al. 2009).

Figure 5
Figure 5

GHSR antagonism partially reversed chronic stress-induced decline of the primordial follicle reserve. (A) Chronic stress induced a decline in the number of primordial follicles in chronically stressed saline-treated, but not d-Lys3-treated mice. (B) There was a trend for a reduced presence of early primary follicles in the ovaries of chronically stressed mice (P = 0.059). There were no differences in the presence of (C) primary, (D) secondary, (E) antral, (F) ovulatory or (G) atretic follicles. Data are mean ± s.e.m. Connecting lines with * represent post hoc differences. *P < 0.05.

Citation: Journal of Endocrinology 241, 3; 10.1530/JOE-19-0109

There were no differences in the volumes of ovarian cortex, medulla or CL (data not shown). Similarly, no significant differences were found in the proportion of these ovarian divisions of the total ovarian volume (Table 4; n = 6–8). Primordial follicle and oocyte diameters were significantly increased in chronically stressed mice (F(1,24) = 6.04, P = 0.022; F(1,24) = 5.13, P = 0.033; Tables 5 and 6; n = 6–8), but no further differences were evident in other follicle populations.

Table 5

Mean follicle diameters (µm2).

Follicle stageNS SalNS d-Lys3Stress SalStress d-Lys3Statistics
Primordial17.6 ± 0.517.4 ± 0.318.0 ± 0.4 18.9 ± 0.4$
Early primary23.3 ± 0.623.8 ± 0.625.6 ± 0.924.0 ± 1.4ns
Primary43.9 ± 2.945.0 ± 1.540.4 ± 1.646.5 ± 1.6ns
Secondary133.2 ± 6.2135.5 ± 5.9125.0 ± 3.8128.2 ± 5.9ns
Antral297.7 ± 23.4284.1 ± 10.9271.3 ± 25.5291.8 ± 15.7ns
Ovulatory403.3 ± 11.4406.3 ± 16.9419.9 ± 9.9446.9 ± 27.1ns

N = 6–8. Data are mean ± s.e.m.

$Represents main effect of stress; P < 0.05.

ns: not significant.

Table 6

Mean oocyte diameters (µm2).

Follicle stageNS SalNS d-Lys3Stress SalStress d-Lys3Statistics
Primordial10.8 ± 0.310.2 ± 0.311.4 ± 0.311.1 ± 0.3$
Early primary13.2 ± 0.313.3 ± 0.413.8 ± 0.512.5 ± 1.1ns
Primary22.8 ± 1.423.6 ± 1.020.5 ± 0.7922.3 ± 1.1ns
Secondary57.8 ± 1.660.1 ± 0.957.2 ± 1.155.9 ± 1.3ns
Antral60.9 ± 3.269.2 ± 2.564.9 ± 4.064.2 ± 1.6ns
Ovulatory67.3 ± 3.5 66.3 ± 4.268.3 ± 3.270.2 ± 2.1ns

N = 6–8. Data are mean ± s.e.m.

$Represents main effect of stress; P < 0.05.

ns, not significant.

Chronic stress leads to a significant increase in the expression of AMH

Since we saw a reduction in the primordial follicle pool with stress, we examined the expression of other factors regulating primordial follicle development. We saw no differences in the expression of bone-morphogenetic protein 15 (Bmp15) and growth differentiation factor 9 (Gdf9) – oocyte-derived growth factors that regulate primordial follicle recruitment (Fig. 6A and B; n = 5–8). However, there was a reduction in the expression of transcriptional factor forkhead box O3 (Foxo3) in the ovaries of d-Lys3 treated mice (F(1,22) = 4.8, P = 0.039; Fig. 6C; n = 5–8). Interestingly, despite a significant reduction in the size of the primordial follicle pool in stressed mice, there was a significant increase in the expression of Amh mRNA in the ovaries of this group (F(1,21) = 7.44, P = 0.013; Fig. 6D; n = 5–8), and a similar chronic stress-induced increase in the fluorescence intensity of AMH (F(1,23) = 28.1, P < 0.001; n = 6–7). AMH, expressed in granulosa cells of primary, secondary and small antral follicles, controls and inhibits premature recruitment of primordial follicles (Durlinger et al. 1999, 2002). Since AMH is most strongly expressed in granulosa cells of secondary and early antral follicles, diminishing its expression at the later preovulatory stage (Visser et al. 2006), we additionally assessed AMH fluorescence intensity specifically in these follicle types where we also found a significant effect of stress (F(1,23) = 32.25, P < 0.001, Fig. 6E and F; n = 6–7). AMH’s increased expression in the ovaries of chronically stressed mice indicates possible compensatory signalling to prevent further depletion of primordial follicles. Together, these findings suggest that despite notable effects of chronic stress on ovarian health, there are compensatory mechanisms in place to curtail the extent of this. They further suggest acyl ghrelin may play a role in the effects of stress on the ovary.

Figure 6
Figure 6

Chronic stress increases the expression of ovarian AMH. There were no effects of stress and GHSR antagonism on the expression of (A) bone-morphogenetic protein 15 (Bmp15) and (B) growth differentiation factor 9 (Gdf9). (C) Treatment with GHSR antagonist significantly reduced ovarian expression of forkhead box O3 (Foxo3). Chronic stress led to a significant increase in (D) ovarian gene expression of anti-Müllerian hormone (Amh) and (E) fluorescence intensity of AMH-positive follicles. (F) Representative images of AMH staining (red). Scale bars = 50 µm. Data are mean ± s.e.m. Parentheses and $ represent main effect of stress; parentheses and # represent an effect of drug. $,#P < 0.05. A full colour version of this figure is available at https://doi.org/10.1530/JOE-19-0109.

Citation: Journal of Endocrinology 241, 3; 10.1530/JOE-19-0109

Discussion

Here, we aimed to investigate the role of acyl ghrelin in stress-induced suppression of indices of reproductive capacity, with a particular focus on the ovary. Our findings show that chronic predator exposure, an ethologically relevant model of stress, elevated acyl ghrelin levels, as expected. While there were no stress-induced changes in circulating gonadotropins, GHSR antagonism stress-dependently affected the expression of genes regulating ovarian sensitivity to gonadotropins. It also rescued stress-induced depletion of primordial follicles, a finite ovarian follicle population. These data therefore suggest that antagonism of the ghrelin receptor during chronic stress attenuates the detrimental effects of stress on the ovarian primordial follicle pool, findings that have implications for long-term reproductive success.

Stress is a ubiquitous facet of our lives, and the way we deal with and manage stress has important implications for our psychological and physical health. Chronic and unmanageable stress predisposes some individuals to develop depression and anxiety, to suppress immune function and to increase susceptibility to infections or inflammatory diseases (McEwen 2004, Glaser & Kiecolt-Glaser 2005, Webster Marketon & Glaser 2008). Stress-induced hormonal changes have also been repeatedly shown to inhibit HPG axis activity (reviewed in Whirledge & Cidlowski 2013, Sominsky et al. 2017). Interestingly, in our study, the overall changes to the reproductive axis were relatively minimal, despite a month-long daily predator exposure. Together with suppressed circulating corticosterone, these findings may indicate a unique response to repeated predator exposure relative to other reported stressors. Chronic predator stress protocols, whereby rodents are exposed to predators or predator-odour, are commonly used to model a post-traumatic stress disorder (PTSD). These chronic predator stress models have high ethological and face validity as they are able to reproduce the individual variability in trauma and stress vulnerability seen in humans (reviewed in Goswami et al. 2013, Zoladz & Diamond 2016). As such, PTSD in humans is often characterised by abnormally low basal levels of cortisol (Yehuda 2002, 2005, Gill et al. 2008). Similarly, in rats, exposure to chronic predator stress significantly reduces corticosterone levels at baseline (Zoladz et al. 2012). Blunted corticosterone responses to stress and increased negative feedback of the HPA axis have also been shown in other animal models of PTSD, but only in female and not male rats (Louvart et al. 2005, Louvart et al. 2006). Female rodents in general appear to show resilience to the effects of chronic stress on several behavioural parameters when compared to males (Luine et al. 2017).

A reduction in corticosterone levels in our study was similar in both saline and d-Lys3-treated stressed mice. This finding potentially indicates that ghrelin’s interactions with stress to affect ovarian follicles are independent of corticosterone, however examples from the literature illustrate that this effect is complex. In other models of chronic stress, where stress increases circulating corticosterone, GHSR KO male mice have an attenuated corticosterone response relative to their WT counterparts (Patterson et al. 2010, Chuang et al. 2011), although this effect is not always seen (Patterson et al. 2013a). Abizaid and colleagues have demonstrated chronic infusions of d-Lys3 into the paraventricular nucleus of the hypothalamus (PVN) increase circulating corticosterone, in male mice given chronic social defeat stress (Patterson et al. 2013b). However, others have shown immobilisation stress increases corticosterone to a similar degree after d-Lys3 (i.p.) as it does after vehicle injections (Meyer et al. 2014). Therefore, our current findings of suppressed corticosterone levels in chronically stressed female saline- and d-Lys3-treated mice may reflect the stress paradigm we used, but may also be specific to species or sex, elements that would be interesting to explore in future studies. In this context, we would also like to acknowledge the different housing conditions of non-stressed and stressed mice, with the latter being single-housed for the duration of the experiment to avoid social transmission of stress as well as consolation behaviours between the cage-mates (Bruchey et al. 2010, Ben-Ami Bartal et al. 2011, Burkett et al. 2016). This is particularly relevant to female mice that have an ability to buffer synaptic changes after stress in the presence of a partner (Sterley et al. 2018). On the other hand, since female mice also show increased stress reactivity to social isolation (Senst et al. 2016), we avoided changing the housing conditions of control mice. There is therefore a possibility that single-housing conditions of stressed mice contributed to the effects of chronic predator stress in this study.

As anticipated, acyl ghrelin levels were increased in chronically stressed mice, corresponding with previous findings in different models of chronic stress (Lutter et al. 2008, Ochi et al. 2008, Kodomari et al. 2009, Meyer et al. 2014, Yousufzai et al. 2018). The changes in circulating corticosterone and acyl ghrelin were not reflected in the mRNA expression of CRH or glucocorticoid receptors, nor in the expression of genes encoding ghrelin, ghrelin acylating enzyme (ghrelin O-acyltransferase) or GHSR; all of which remained normal after chronic stress. It is, however, unknown to what degree local expression of these genes in the ovary mediates the effects of circulating versus locally produced hormones (Caminos et al. 2003, Miller et al. 2005, Wypior et al. 2011).

While the inhibitory effects of stress on GnRH pulsatility and pituitary gonadotropins have been well documented (Whirledge & Cidlowski 2010), here we did not see changes in circulating LH and FSH. Although LH pulsatility was not assessed in this study, there was also no indication that oestrous cyclicity was affected by stress or GHSR antagonism. In this study, the monitoring of oestrous cyclicity was limited to the last four to six consecutive days, to minimise additional handling that could have disturbed otherwise non-stressed controls. This short time of observation allowed us to control for cycle stage at cull. However, this time frame may not have been sufficient to obtain a complete profile of cycle regularity (Caligioni 2009). Disrupted oestrous cyclicity and inhibition of pituitary gonadotropins is often (reviewed in Whirledge & Cidlowski 2010), but not always, (Geraghty et al. 2015, Shimamoto et al. 2015) an outcome of chronic stress exposure in female rodents. As such, Geraghty et al. have shown that despite normal cycling and no changes in LH secretion, chronic stress in rats induced other reproductive-related effects, including reduced fertility and poor pregnancy outcomes (Geraghty et al. 2015). Gonadotropin-treatment alone is also not sufficient to completely rescue chronic stress-induced suppression of ovarian follicle development (Wu et al. 2012b).

We did see, however, a change in the expression of genes regulating the potential of the ovary to respond to FSH: Fshr and Inhbb. FSH is crucial for follicle maturation, driving the transition of secondary follicles into the antral stage and mediating the survival of antral follicles. The effects of FSH are tightly regulated by its receptor and changes in the expression of the FSH receptor control the responsiveness of follicles to FSH and hence determine their survival and the likelihood of ovulation (McGee & Hsueh 2000).

In addition to its receptor, the impact of FSH on the growing follicles is mediated by activins and inhibins, with activins stimulating and inhibins reducing pituitary FSH synthesis and release. Here, we saw that stress interacted with the d-Lys3 to affect the expression of both Inhbb, a gene encoding inhibin βB subunit that is restricted to small antral follicles (Roberts et al. 1993) and Fshr. These changes indicate that under conditions of stress, GHSR antagonism elicits different responses to those under non-stressed conditions. Statistically, the lack of post hoc differences was not driven by insufficient effect sizes or low power, and the significant interaction effects suggest that the effects of stress were strongly dependent on drug and vice versa. In other words, in saline-treated animals stress increased the expression of Fshr and Inhbb, while an opposite pattern is seen in d-Lys3-treated stressed mice. As such, these data indicate a potential for GHSR antagonism to stress-dependently alter the expression of these genes that contribute to the regulation of follicle sensitivity to FSH. However, caution should be exercised in interpretation of these findings, since these demonstrate only changes in mRNA. While the role of ghrelin in the regulation of female fertility is still emerging, its different roles under different conditions have been demonstrated in at least one study, whereby both exogenous ghrelin and inhibition of endogenous ghrelin by d-Lys3 exerted negative effects on fertilisation and early embryo development (Luque et al. 2014). These data suggest appropriate concentrations of ghrelin are required for optimal reproductive success. However, future research will need to determine whether these assumptions also pertain to the divergent effects of d-Lys3 as seen in our study, in the context of stress.

In addition to FSH, CNP, encoded by Nppc, is another factor regulating antral follicle maturation. CNP prevents premature maturation of oocytes and is significantly decreased after the ovulatory LH surge (Kawamura et al. 2011). Decreased expression of Nppc in d-Lys3-treated mice may therefore indicate premature resumption of meiosis. Overall, these findings suggest that chronic stress and GHSR antagonism independently influence the expression of genes regulating follicle maturation and the potential for ovulation.

The above changes in ovarian gene expression were not associated with changes in ovarian atresia, apoptosis or proliferation and did not affect the large follicle counts. This is somewhat surprising since chronic unpredictable stress in mice has been shown to increase the presence of atretic secondary and antral follicles, reducing their healthy populations (Wu et al. 2012b). Chronic restraint stress has also been demonstrated to increase the presence of apoptotic markers in some, but not all ovarian cell types (Liang et al. 2013). It is important to note, however, that there are no current apoptotic markers that accurately detect primordial follicle atresia in adult ovary, and the presence of apoptosis in this follicle population is most accurately reflected by follicle counts (Tingen et al. 2009, Liew et al. 2017). Indeed, we saw a significant reduction of primordial follicles in chronically stressed mice. The non-renewing primordial follicles are the most critical component of the ovary, and the size of this initial follicle pool is a major predictor of the female reproductive lifespan (Bristol-Gould et al. 2006). Once established, this population of follicles develops through to primary, secondary and antral stages. Most antral follicles will undergo atresia, while only a few will respond to postpubertal gonadotropin stimulation and reach the ovulatory stage. Earlier reduction of this reserve thus strongly predicts earlier reproductive decline and hence premature reproductive senescence (McGee & Hsueh 2000).

The stress-induced reduction in primordial follicles per mm2 of ovary was not apparent in mice treated with the GHSR antagonist. Similarly, our raw follicle counts revealed a stress-dependent effect of d-Lys3 treatment, suggesting a potential protection from the effects of chronic stress. While acyl ghrelin has been shown to inhibit apoptosis in certain cell types (Granata et al. 2007, Zhu et al. 2014), the evidence for its role in mediating ovarian apoptosis is still limited. Although some studies have provided evidence for an anti-apoptotic activity of acyl ghrelin in ovarian cells (Rak et al. 2009, Rak-Mardyla & Gregoraszczuk 2010), that is potentially GHSR independent (Rak et al. 2009, Sirotkin et al. 2011), others suggest increased levels of ghrelin disrupt ovarian steroidogenesis, inhibit ovarian development and impair embryo development, through cumulus cell apoptosis and DNA damage during in vitro maturation (Roa et al. 2010, Rak-Mardyla et al. 2015, Sirini et al. 2017). Interestingly, however, despite an apparent potential for the GHSR antagonist to rescue the decline in the number of primordial follicles, d-Lys3-treated mice had a reduced expression of the transcription factor Foxo3. Changes in Foxo3 gene expression in oocytes regulate primordial follicle quiescence, and depletion of this gene leads to increased activation and early depletion of the primordial pool (Castrillon et al. 2003). The Foxo3 gene is also expressed in granulosa cells, where together with Foxo1 it mediates follicle development through a complex interaction with activin-regulated genes (Liu et al. 2013). The function of Foxo3 in granulosa cells is different from its role in oocytes. Thus, the current findings that are derived from the analysis of whole ovary may have limited interpretation, particularly considering there were no effects of GHSR antagonism or stress on the expression of Bmp15 and Gdf9, oocyte-derived factors that promote early folliculogenesis, granulosa cell proliferation and FSH-dependent follicle development (Hsueh et al. 2015). It is also essential to mention that a systemic route of d-Lys3 administration suggests any local ovarian effects of the antagonist may be driven by its action at the level of the hypothalamus, the pituitary or other sites. It is also plausible that GHSR antagonism-driven physiological responses, such as an increase in body weight that we see in our study, may also play role in the intraovarian changes. We saw, however, a significant increase in AMH gene and protein expression in the ovaries of chronically stressed mice. AMH is produced by granulosa cells of primary to small antral follicles, inhibiting the recruitment of primordial follicles into the growing pool and reducing secondary and small antral follicle sensitivity to FSH (Durlinger et al. 1999, 2002). Therefore, this increase in ovarian AMH appears to be compensatory, possibly counteracting the depletion of the primordial follicle pool and changes in genes that contribute to FSH sensitivity.

Overall, our data indicate that chronic stress has subtle but potentially critical effects on female reproductive health and that at least some of these effects may be mediated by stress-induced acyl ghrelin. Although depletion of the primordial reserve can be compensated for by a slower rate of primordial follicle recruitment into the growing pool, in the long term, this may lead to premature reproductive senescence. While we have not assessed mating potential and pregnancy in the current study, others have shown chronic stress in mice impairs oocyte developmental potential and reduces blastocyst formation (Wu et al. 2012a, Gao et al. 2016). Even 4 days after cessation of stress and despite normal regularity of oestrous cyclicity, chronically stressed rats exhibit fewer successful mating events, fewer pregnancies and increased embryo resorption (Geraghty et al. 2015). Nevertheless, the long-term consequences of chronic stress on fertility and the role of ghrelin in mediating these effects remain to be further elucidated. It is also important to note that although young and otherwise healthy individuals may experience only temporary and potentially reversible effects of stress, our findings may have significant implications for those already suffering from underlying fertility issues, for whom even a minor perturbation of ovarian function may impede pregnancy success.

Declaration of interest

The authors declare that there is no conflict of interest that could be perceived as prejudicing the impartiality of the research reported.

Funding

This work was supported by an RMIT Vice-Chancellor’s Postdoctoral Fellowship and an RMIT University Grant Development Support to L S. S J S is a recipient of a National Health and Medical Research Council Career Development Fellowship II (APP1128646).

Acknowledgements

The authors would like to thank Dr Wendell Cockshaw for his statistical advice. Sarah J Spencer and Luba Sominsky: Joint senior authorship.

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      Society for Endocrinology

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    Effects of chronic stress and GHSR antagonism on oestrous cycle, body weight, circulating corticosterone and acyl ghrelin. (A) Study design. Female mice were treated with d-Lys3 (2.74 µg/kg, i.p.) or saline, and 30 min later were subjected to 30 min chronic predator stress or left undisturbed in their home cage. This procedure was repeated daily for 4 weeks. (B) Chronic stress and GHSR antagonism did not affect the regularity of oestrous cyclicity. (C) Exposure to chronic stress caused a significant reduction in body weight by two weeks after the commencement of stress, while d-Lys3 treatment significantly increased weight gain at the beginning of the second and third experimental weeks. (D) Chronic predator stress significantly reduced concentrations of circulating corticosterone, and (E) significantly increased the levels of acyl ghrelin. (F) Chronic stress increases the release of growth hormone (GH) in saline-treated but not d-Lys3-treated mice. Data are mean ± s.e.m. Parentheses and $ represent main effect of stress; Parentheses and # represent an effect of drug. Connecting lines with * represent post hoc differences. $,#,*P < 0.05.

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    Ovarian expression of stress and ghrelin regulatory genes is unchanged. Chronic stress and GHSR antagonism did not affect ovarian gene expression of (A) corticotropin-releasing hormone receptor 1 (Crhr1), (B) corticotropin-releasing hormone receptor 2 (Crhr2), (C) glucocorticoid receptor (Nr3c1), (D) mineralocorticoid receptor (Nr3c2), (E) ghrelin (Ghrl), (F) ghrelin O-acyltransferase (Mboat4) and (G) growth hormone secretagogue receptor Ghsr. Data are mean ± s.e.m.

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    Chronic stress and GHSR antagonism influence the expression of genes regulating ovarian responsiveness to FSH. Chronic stress and GHSR antagonism had no effect on (A) circulating luteinising hormone (LH), (B) gene expression of LH receptor (Lhcgr) and (C) circulating follicle-stimulating hormone (FSH). (D) Gene expression of ovarian FSH receptor is affected by chronic stress and GHSR antagonism. (E) There were no changes in the gene expression of inhibin βA subunit (Inhba), but (F) an effect of chronic stress and GHSR antagonism on the expression of inhibin βB subunit (Inhbb). (G) The expression of the expression of Nppc, a gene encoding for CNP was significantly reduced in d-Lys3-treated mice. (H) There were no differences in the percentage of Ki-67-positive proliferating follicles. Data are mean ± s.e.m. Parentheses and $ × # represent a significant stress × drug interaction with no post hoc differences; parentheses and # represent main effect of drug. $,#P < 0.05.

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    Chronic stress and GHSR antagonism had no effect on ovarian apoptosis. There were no changes in gene expression of apoptosis regulators (A) Bax, (B) Bcl2 and (C) caspase 3 (Casp3). (D) There were no differences in the percentage of TUNEL-positive follicles. (E) Representative images of TUNEL (green). Scale bars = 50 µm. Data are mean ± s.e.m. A full colour version of this figure is available at https://doi.org/10.1530/JOE-19-0109.

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    GHSR antagonism partially reversed chronic stress-induced decline of the primordial follicle reserve. (A) Chronic stress induced a decline in the number of primordial follicles in chronically stressed saline-treated, but not d-Lys3-treated mice. (B) There was a trend for a reduced presence of early primary follicles in the ovaries of chronically stressed mice (P = 0.059). There were no differences in the presence of (C) primary, (D) secondary, (E) antral, (F) ovulatory or (G) atretic follicles. Data are mean ± s.e.m. Connecting lines with * represent post hoc differences. *P < 0.05.

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    Chronic stress increases the expression of ovarian AMH. There were no effects of stress and GHSR antagonism on the expression of (A) bone-morphogenetic protein 15 (Bmp15) and (B) growth differentiation factor 9 (Gdf9). (C) Treatment with GHSR antagonist significantly reduced ovarian expression of forkhead box O3 (Foxo3). Chronic stress led to a significant increase in (D) ovarian gene expression of anti-Müllerian hormone (Amh) and (E) fluorescence intensity of AMH-positive follicles. (F) Representative images of AMH staining (red). Scale bars = 50 µm. Data are mean ± s.e.m. Parentheses and $ represent main effect of stress; parentheses and # represent an effect of drug. $,#P < 0.05. A full colour version of this figure is available at https://doi.org/10.1530/JOE-19-0109.

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