Early life adversity accelerates hypothalamic drive of pubertal timing in female rats with associated enhanced acoustic startle

Abstract

Early life adversity in the form of childhood maltreatment in humans or as modeled by maternal separation (MS) in rodents is often associated with an earlier emergence of puberty in females. Earlier pubertal initiation is an example of accelerated biological aging and predicts later risk for anxiety in women, especially in populations exposed to early life trauma. Here we investigated external pubertal markers as well as hypothalamic gene expression of pubertal regulators kisspeptin and gonadotropin-releasing hormone, to determine a biological substrate for MS-induced accelerated puberty. We further investigated a mechanism by which developmental stress might regulate pubertal timing. As kisspeptin and gonadotropin-releasing hormone secretion are typically inhibited by corticotropin releasing hormone at its receptor CRH-R1, we hypothesized that MS induces a downregulation of Crhr1 gene transcription in a cell-specific manner. Finally, we explored the association between pubertal timing and anxiety-like behavior in an acoustic startle paradigm, to drive future preclinical research linking accelerated puberty and anxiety. We replicated previous findings that MS leads to earlier puberty in females but not males, and found expression of kisspeptin and gonadotropin-releasing hormone mRNA to be prematurely increased in MS females. RNAscope confirmed increased expression of these genes, and further revealed that kisspeptin-expressing neurons in females were less likely to express Crhr1 after MS. Early puberty was associated with higher acoustic startle magnitude in females. Taken together, these findings indicate precocial maturation of central pubertal timing mechanisms after MS, as well as a potential role of CRH-R1 in these effects and an association with a translational measure of anxiety.

Emergence of puberty is a pivotal event in early adolescence that triggers both reproductive and non-reproductive processes in the brain and body. Pubertal timing is also a measure of biological aging, with genetic and environmental factors impacting the tempo at which puberty is reached. Importantly, an earlier age of pubertal initiation is associated with later life anxiety, especially in girls, and this has been associated with maturation of corticolimbic circuits mediating threat responsiveness (Miller et al., 2020). Early life adversity (ELA) is associated with accelerated pubertal development in girls (Bleil et al., 2017; Colich et al., 2020), and this accelerated puberty predicts anxiety symptomology later in adolescence and adulthood (Colich and McLaughlin, 2022; Stenson et al., 2021; Winer et al., 2017). However, clinical evidence is correlative and unavoidably confounded by psychosocial factors, therefore the presence of causative mechanisms or a biological substrate underpinning these effects is largely unknown.

Puberty is controlled by the activity of the hypothalamic-pituitary-gonadal (HPG) axis. Gonadotropin-releasing hormone (GnRH) neurons in the hypothalamus stimulate pituitary release of gonadotropins, which stimulate the gonads to release the sex steroid hormones testosterone and estrogen (Delemarre et al., 2008). The initiation of this process relies on the pulsatile release of GnRH and the subsequent release of lutenizing hormone and follicular stimulating hormone. GnRH neurons are influenced by a number of modulatory neuropeptides that respond to environmental and metabolic demands (Herbison, 2016). Kisspeptin is one such regulator and prerequisite for pubertal initiation, through actions at the receptor GPR54. Mice with mutations in kisspeptin or GPR54 fail to initiate puberty, and manipulations of kisspeptin/GPR54 affect fertility in adult animals (Seminara et al., 2003; Topaloglu et al., 2012). The arcuate (ARC) and anteroventral periventricular (AVPV) nuclei of the hypothalamus have a prominent representation of kisspeptin-expressing neurons, with kisspeptin in the AVPV being particularly critical for pubertal initiation and maintenance of normal cycling (Wang and Moenter, 2020); region-specific downregulation of kisspeptin in the AVPV, but not the ARC, caused delayed vaginal opening in females (Hu et al., 2015). Electrophysiological and neuroendocrine studies have revealed that the influence of kisspeptin on GnRH activation begins early in postnatal development, supporting their role in later reproductive development and the possibility of dysregulation by early life events (Bateman and Patisaul, 2008; Clarkson and Herbison, 2006; Han et al., 2005; Kauffman et al., 2007).

Notably, experiences of stress have downstream effects on the HPG axis. Evolutionary hypotheses posit that HPA and HPG crosstalk facilitates adaptation of individuals’ reproductive strategies to changes in the environment (Acevedo-Rodriguez et al., 2018). Classic theories suggest that stress impedes concurrent reproductive capacity by directing resources to the processes supporting immediate survival rather than energy-expensive ones, like reproduction (Hochberg and Belsky, 2013). Paradoxically, stress exposure during early postnatal development has been shown to stimulate the reproductive axis and lead to earlier puberty onset (Hochberg and Belsky, 2013; Moffitt et al., 1992; Strzelewicz et al., 2019). In rats, maternal separation ELA accelerates puberty (Cowan and Richardson, 2019; Grassi-Oliveira et al., 2016), providing a model for studying the ELA-attributable accelerated puberty observed in humans, but the mechanisms of these developmental impacts are not yet fully understood.

Here we test the hypothesis that seemingly paradoxical effects of early developmental stress on pubertal initiation involve the influences of the HPA neuropeptide corticotropin releasing hormone (CRH) on the HPG axis. CRH fibers and GnRH perikayra are in close proximity in the hypothalamus, and direct synaptic connections between CRH and GnRH have been identified (Dudas and Merchenthaler, 2002; MacLusky et al., 1988). Additionally, a subset of GnRH neurons express the CRH receptor type 1 (CRH-R1), and intra-hypothalamic administration of CRH suppresses GnRH secretion (Phumsatitpong et al., 2021). In typically-developing animals, stress suppresses reproductive functioning via CRH and glucocorticoid signaling on GnRH- and kisspeptin-expressing neurons in the anterior hypothalamus (Grachev et al., 2014). However, CRH-R1 and CRH-R2 antagonists do not completely prevent the effects of stress on puberty initiation (Li et al., 2006). Intermediate modulators, like kisspeptin neurons, may also contribute to the effects of stress on GnRH suppression. In fact, nearly all kisspeptin neurons in the AVPV and ARC co-express CRH-R1, and icv infusions of CRH reduces kisspeptin expression in the hypothalamus (Kinsey-Jones et al., 2009). It is therefore possible that heightened CRH activity during ELA impacts the ability of hypothalamic CRH-R1 to stave off puberty, resulting in acceleration of pubertal timing.

Precocial or accelerated puberty in girls is of growing concern across the world (Eckert-Lind et al., 2020), requiring new understanding of medical and mental health consequences. Correlational data from clinical studies (Belsky et al., 2015; Deardorff et al., 2021; Hartman et al., 2017) point to associations between heightened pubertal tempo and increased anxiety. Therefore, here we tested the following hypotheses and predictions to investigate mechanisms linking ELA to accelerated puberty and an association between accelerated puberty and heightened anxiety-like behavior in a rodent model: (1) If ELA exposure directly impacts pubertal timing through hypothalamic mechanisms in females, then female, but not male, rats subjected to maternal separation ELA will display premature expression of hypothalamic kisspeptin (Kiss1 mRNA) and GnRH (Gnrh mRNA) as well as earlier emergence of external puberty markers; (2) If accelerated pubertal timing following ELA is due to downregulation of CRH-R1 in the hypothalamus, then hypothalamic Crhr1 mRNA will be decreased in maternally separated females in Kiss1- and Gnrh-expressing neurons within the hypothalamus. We also explored an association between pubertal timing and baseline acoustic startle—a measure of emotional arousal (Hantsoo et al., 2018)—to guide future studies investigating causative mechanisms linking pubertal tempo and anxiety-like behaviors in a rodent model.

Methods

Animals and Maternal Separation

All experiments were performed in accordance with the 1996 Guide for the Care and Use of Laboratory Animals (NIH) with approval from the Institutional Animal Care and Use Committee at Northeastern University.

All experiments began with in-house mating of male and female Sprague-Dawley rats originally obtained from Charles River Laboratories (Wilmington, MA). One male and one female were caged together until pregnancy was confirmed by checking for the presence of sperm in a vaginal swab each morning for a maximum of 4 days. All females were nulliparous, and pregnant dams were housed singly upon confirmation of pregnancy. Each litter was randomly assigned to maternal separation (MS) or control (Con) treatment conditions. Parturition was checked daily, and the day of birth was denoted as P0. On P1, litters were culled to 10 pups with 5 males and 5 females when possible. Pup sex was determined by anogenital distance on P1. Subjects were identified by toe clips, which were performed on P5.

MS litters underwent maternal separations daily from P2-P20. From P2-P10, each pup was placed in an individual plastic cup containing pine shavings from the home cage to maintain a familiar odor. Cups were placed in a circulating water bath kept to 37°C for 3.5 hours (0930h-1300h). From P11-P20, when pups were able to adequately thermoregulate, each pup was placed in an individual mouse cage containing clean bedding mixed with a small handful of home cage bedding for 4 hours (0930h-1330h). MS dams remained in the home cage in a separate room for the duration of the separation. Con and MS pups were weighed on P9 and P20. To control for the experience of experimenter handling received by MS pups, while avoiding the introduction of a daily brief handling manipulation which has nuanced developmental effects (Raineki et al., 2014), Con pups were also briefly handled by an experimenter on P12 and P15 for 3 minutes each in addition to weighing on P9 and P20, but otherwise were left undisturbed.

On P21, all rats were weaned and pair-housed in standard caging with sex- and condition-matched cage mates. Standard laboratory conditions include a polycarbonate wire-top caged with pine shave bedding, a plexiglass tube for enrichment, and food and water available ad libitum. The facility was kept on a 12-hour light/dark cycle (light period between 0700–1900) with regulated temperature (22–23°C) and humidity (37%−53%). Post-weaning, animals were left undisturbed except for weekly cage cleanings. No more than 2 male and 2 females per litter were used in each experimental group. Therefore, a total of 12 MS litters and 12 Control litters were used in this study, and from individual litters 1–2 pups per sex were tested for pubertal status and acoustic startle, 1–2 pups per sex at each age were used for qPCR, and 1–2 females were taken at P20 for RNAscope, with remaining animals used in other ongoing studies.

Pubertal status measurement

One cohort of animals (n=10–11/group) was used to monitor pubertal status, then were undisturbed other than cage cleaning until assessment of acoustic startle response at P34. After weaning, animals’ pubertal status was monitored daily by visual inspection, during which animals were picked up individually by an experimenter and inspected for 15–30 seconds per day. For males, pubertal initiation was determined by the initial preputial separation, defined as any separation of the prepuce from the glans penis. Full puberty was defined as the ability to fully retract the prepuce. For females, pubertal initiation was defined as the appearance of a vaginal pinhole, which typically appears 2–5 days before full vaginal opening.

Validation of female puberty initiation

In a subset of 6 MS and 4 Con-reared females from a separate cohort, vaginal opening was rated prior to sacrifice and uteri were dissected and weighed to verify our designation of pubertal initiation. Uterine wet weight was normalized to total body weight.

Tissue Collection and Real-Time Quantitative Polymerase Chain Reaction (qPCR) to measure Gnrh, Kiss1, and Crhr1 expression in the hypothalamus

In a separate cohort, tissue was collected for qPCR at P10, 15, or 20 (n=8–11/group). Male and female MS and Con animals were mildly anesthetized with isoflurane and decapitated, and the brains were immediately extracted, frozen in cold 2-methylbutane, and stored at −80°C until tissue processing.

Brains were sectioned on the cryostat to obtain 300μm-thick hypothalamus punches (Fig. 1) from the anterior hypothalamus (AH) (for Gnrh and Kiss1) and posterior hypothalamus (PH) (for Kiss1). Kiss1 is most densely expressed in the AVPV and the ARC, and Gnrh is expressed in the medial preoptic area (MPOA). AH punches included both the MPOA and AVPV, while PH punches contained the ARC.

Figure 1.

Plates adapted from rat brain atlas (Paxinos and Watson, 1998) illustrating where punches were taken for qPCR (rostral face of 300 μm section) or RNAscope (most rostral section of series)

AH and PH punches were analyzed separately. Total RNA from each region was isolated using the RNAqueous-4PCR Total RNA Isolation Kit (Applied Biosystems, Foster City, CA, United States) and processed per the manufacturer’s instructions. cDNA was synthesized using the High Capacity cDNA Reverse Transcriptase Kit (Applied Biosystems, Foster City, CA, United States). cDNA concentration was quantified using Nanodrop 2000 Spectrophotometer (Thermo Fisher Scientific), and samples were diluted to 400 μg/μl. Amplification was performed using TaqMan Gene Expression Assays using gene-specific primers purchased from Thermo Fisher Scientific for Gnrh (NM_012767.2) and Kiss1 (NM_181692.1) in AH punches, and for Kiss1 in PH punches. Reactions were performed in triplicate on a StepOnePlus Real-Time PCR System (Applied Biosystems) using standard cycling conditions, as recommended by the manufacturer. 4ul of diluted cDNA was placed in 20μl reaction plate containing 16μl of master mix and 1x dilution of each primer. Reactions were performed with a holding stage of 50°C for 2 min and 95°C for 10 min, followed by 40 subsequent cycles of 15s at 95°C and 1 min at 60°C. The threshold cycle (CT) (number of cycles required to reach detection threshold) was determined for each reaction. Standard curves were generated for each primer to determine primer efficiency, and the Pflaffl method used to determine gene expression relative to the house-keeping control gene, β-actin (Thermo Fisher Scientific, AILJKN2). The male control reference was the P10 control-reared average value, and the female control reference was the P10 control-reared average value. Data are expressed as the base 2 logarithm of the relative gene expression ratio.

Subcellular identification of hypothalamic gene expression with RNAscope to analyze Crhr1 expression

In a separate cohort of animals, female rats were sacrificed on P20 by rapid decapitation and brains were extracted as described above (n=8/group). Brains were sectioned on a cryostat to a thickness of 20μm. A total of 9 serial sections from the anterior end of the AH were mounted onto 3 Fisherbrand ColorFrost Plus microscope slides and stored at −80°C until the RNAscope assay. One slide containing 3 sections separated by 60μm was assayed for each subject.

Sections were fixed with 4% paraformaldehyde for 15 minutes, then dehydrated in a series of EtOH solutions. Then, sections were pretreated with hydrogen peroxide and Protease IV provided by RNAscope Multiplex Fluorescent Reagent Kit, diluted 1:2 in 1x PBS. Sections were hybridized with a triple probe mix for Kiss1, Gnrh, and Crhr1, and probes were developed with channel-specific HRPs using Opal 520 (Kiss1), Opal 570 (Gnrh), and Opal 690 (Crhr1) and counterstained with DAPI.

The Zeiss Axio Imager M2 was used for microscopic image acquisition. 10 Z-stack images were acquired at 20X magnification for DAPI, GFP, DsRed, and Cy5 and the image stack maximum projections were saved for later analysis. A total of 18 images were taken in the AH (3/hemisphere/section).

The 4 channels were batch merged in ImageJ and uploaded to QuPath for spot and cell identification. First, the cell detection tool was used to identify DAPI stained nuclei. A radius of 10μm was allowed around all nuclei for the purpose of assigning subcellular RNAscope detections to cells. Subcellular Kiss1, Gnrh, and Crhr1 clusters were identified using the subcellular detection tool, and cells were classified as Kiss1, Gnrh, Crhr1, colabeled Kiss1: Crhr1, or colabeled Gnrh: Crhr1. Once cell types were identified in QUPath, detections and classifications were confirmed by a trained experimenter blind to experimental conditions. Outcome measures were cell count (normalized to DAPI), mean cell intensity, and number of spots estimated by QuPath for each probe.

Acoustic Startle

On P34, animals that were assessed for pubertal timing were tested in an acoustic startle paradigm (Med Associates product number: MED-ASR-PRO1). The startle cabinets were equipped with sound attenuating foam on all walls and doors. Each cabinet held a grid rod animal holder on a startle platform containing the load cell. The load cell and load cell amplifier were used to convert force on the platform to a voltage representing the startle response. Speakers for delivering white noise background and startle noise bursts were positioned 1” behind the animal holder. The grids rods on the back of the animal holder provide ventilation and do not interfere with sounds.

Because weights differ between ages and sexes, the startle boxes were calibrated to accommodate the weight of each rat being tested. Rats were transported to the testing room and left to acclimate for 10 minutes. After acclimation, each rat was placed into the animal holder in the startle cabinet. The experiment began with 5 minutes of white background noise to acclimate the animal to the startle cabinet. Then, 100 stimulus tones (50ms white noise tones of 95dB, 100dB, 105dB, or 110dB presented with a 3ms rise/fall time) were presented. Each session consisted of 25 presentations of stimuli at each dB level, in ascending order. The background noise level was set to 70dB throughout the experiment. At the end of 100 trials, rats were removed from the boxes and returned to their home cages. Boxes were cleaned with 40% EtOH between runs. Startle magnitudes are reported as the average peak magnitude across all white noise presentations.

Statistical analysis

Group sizes were determined using G*Power with effect sizes determined from previous studies measuring pubertal timing with sex × rearing ANOVA, and preliminary t-tests measuring Kiss1 and Gnrh gene expression. A total sample size of 35–40 over 2 sexes and 2 rearing groups was determined to yield power of 0.80 – 0.9. Sex × rearing 2-way ANOVA was used to compare ages of pubertal initiation or completion (vaginal opening or preputial separation). Weights at P9, P20, and P34 were compared using a 3-way ANOVA with age as a repeated measure and Greenhouse–Geisser correction for lack of sphericity. Relative gene expression from qPCR data were analyzed with a 3-way sex × age × rearing ANOVA. Post-hoc tests were conducted using the Šidák test to correct for multiple comparisons. For P20 RNAscope analyses, differences between Con and MS animals were determined via either 2-way mixed ANOVA (when Crhr1 colocalization was included as a factor), independent t-tests, or Mann-Whitney tests when unequal variances were detected. Uterine weights were compared with independent t-test. P-values and effect sizes are reported. Because sex × rearing interactions have revealed rearing group differences in acoustic startle magnitude in females but not males (Granata et al., 2022b) and we have observed that MS impacts pubertal timing in females but not males (Grassi-Oliveira et al., 2016), separate linear regressions in males and females were used to analyze the relationship between age of puberty initiation and startle magnitude. Multiple linear regression was first applied to MS and Con animals separately, and when rearing was not identified as a predictor, a single linear regression was applied to all males or all females.

Results

Maternally separated animals gained more weight between P20-P34

Three-way sex × rearing × age ANOVA revealed main effects of age (F_1.1,44.9 = 2361; p < 0.0001; η²= = 0.95) and sex (F_1,37 = 11.94; p = 0.0014; η²= 0.005), as well as age × sex (F_2,74 = 24.97; p < 0.001; η²= 0.01) and age × rearing (F_2,74 = 4.596; p = 0.013; η²= 0.001) interactions. Notably, extremely small effect sizes suggest that these were low-powered analyses. Separate three-way ANOVAs comparing weight at P9 and P20 (during MS) revealed a main effect of age (F_1,37 = 1861; p < 0.001; η²= 0.891) but no interactions or main effects of sex or rearing, while 3-way ANOVA comparing weight at P20 and P34 revealed a main effect of age (F_1,37 = 1855; p < 0.001; η²= 0.912) and an age × sex interaction with a small effect size (F_1,37 = 6.011; p < 0.001; η²= 0.011). Two-way ANOVA within each sex revealed effects of age in both females (F_1,19 = 2327; p < 0.001; η²= 0.936) and males (F_1,18 = 655.5; p < 0.001; η²= 0.941) (Fig. 2a). Comparing the change in weight (weight gain) with 2-way sex × rearing ANOVAs revealed no effects of sex or rearing on weight gained from P9-P20, while a comparison of weight gained P20–35 revealed main effects of both sex (F_1,37 = 25.29; p < 0.001; η²= 0.356) and rearing (F_1,37 = 8.24; p = 0.007; η²= 0.116) (Fig. 2b).

Figure 2.

Effects of maternal separation on body weight at P9, P20, and P34. (a) Body weight of males (top) and females (bottom) at P9 and P20 (during maternal separation). Violin plots of medians and distributions for each group at each age; n = 10–11. (b) Body weight of males (top) and females (bottom) at P20 and P34 (following maternal separation). Violin plots of medians and distributions for each group at each age; n = 10–11. (c) Weight gain from P9–20 (left) and from P20–34 (right) in males and females; n = 10–11. *p < 0.05 main effects of age and rearing. Individual data points with means ± SEM are shown. MS: maternal separation; Con: control.

Exposure to maternal separation yielded earlier pubertal initiation and completion in females

While a 2-way interaction was not significant (p = 0.210; η²= 0.04), both a main effect of sex (F_1,36 = 16.72; p = 0.0002; η ²= 0.4) and of rearing (F_1,36 = 5.82; p = 0.021; η ²= 0.14) were found on initiation of puberty, with females (p = 0.022) but not males (p = 0.608) showing earlier puberty initiation following MS (Fig. 3a).

Figure 3.

Effects of maternal separation on puberty initiation and completion. (a) Females exposed to maternal separation displayed earlier initiation of puberty, as measured by the initiation of vaginal opening; n = 9–11. (b) Females exposed to maternal separation displayed earlier puberty completion, as measured by full vaginal opening; n = 9–11. (c) Uterine weights (normalized to body weight) of females rated with initiation of vaginal opening (pinhole opening) were higher than those rated with no vaginal opening (pre-initiation); n = 4–6. MS: maternal separation; Con: control. *p < 0.05. Individual data points with means ± SEM are shown.

Pubertal completion as assessed by full vaginal opening or full preputial separation was similarly affected by MS. While a 2-way interaction was not significant (p = 0.473; η²= 0.003), both a main effect of sex (F_1,33 = 1118.9; p < 0.0001; η²= 0.745) and of rearing (F_1,33 = 7.10; p = 0.012; η²= 0.044) were found, with females (p = 0.026) but not males (p = 0.380) showing earlier completion of puberty following MS (Fig. 3b).

Uterine weight, a proxy for estrogen levels, was increased in females with the vaginal pinhole on P30 compared to those with no visual sign of pubertal initiation (t₈ = 3.22; p = .012; η²= 0.56) (Fig. 3c).

Exposure to maternal separation yielded accelerated increases of AVPV Kiss1 and MPOA Gnrh in females (See Table 1 for full statistics).

Table 1.

Full statistical information from ANOVAs comparing hypothalamic expression of Gnrh and Kiss1 in MS and Con males and females. PH: qPCR performed on posterior hypothalamus tissue punches, containing the arcuate nucleus; AH: qPCR performed on anterior hypothalamus tissue punches, containing the anteroventral periventricular nucleus and the medial preoptic area. Shaded cells indicate significant main effects or interactions

Gene	ANOVA Results		P value	η2	Post-hoc multiple comparisions
Kiss1 (AH)	Main Effect Age	F (3, 104) = 19.39	P<0.0001	0.306	*P20 Female Con-MS; p = 0.014
	Main Effect Sex	F (1, 104) = 0.1892	P=0.6645	0.001
	Main Effect Rearing	F (1, 104) = 4.850	P=0.0299	0.026
	Age × Sex	F (3, 104) = 1.449	P=0.2330	0.023
	Age × Rearing	F (3, 104) = 3.826	P=0.0121	0.060
	Sex × Rearing	F (1, 104) = 1.912	P=0.1697	0.010
	Age × Sex × Rearing	F (3, 104) = 1.735	P=0.1644	0.027
Gnrh (AH)	Main Effect Age	F (3, 114) = 5.107	P=0.0024	0.087	*P20 Female Con-MS; p = 0.01
	Main Effect Sex	F (1, 114) = 23.42	P<0.0001	0.133
	Main Effect Rearing	F (1, 114) = 0.8903	P=0.3474	0.005
	Age × Sex	F (3, 114) = 1.412	P=0.2430	0.024
	Age × Rearing	F (3, 114) = 3.026	P=0.0325	0.052
	Sex × Rearing	F (1, 114) = 0.4552	P=0.5012	0.003
	Age × Sex × Rearing	F (3, 114) = 2.727	P=0.0474	0.047
Kiss1 (PH)	Main Effect Age	F (3, 115) = 12.86	P<0.0001	0.236
	Main Effect Sex	F (1, 115) = 1.622	P=0.2054	0.010
	Main Effect Rearing	F (1, 115) = 0.005692	P=0.9400	0.000
	Age × Sex	F (3, 115) = 1.129	P=0.3405	0.021
	Age × Rearing	F (3, 115) = 1.328	P=0.2687	0.024
	Sex × Rearing	F (1, 115) = 0.02566	P=0.8730	0.000
	Age × Sex × Rearing	F (3, 115) = 0.2941	P=0.8296	0.005

3-way ANOVA comparing AVPV Kiss1 revealed a main effect of age (p < 0.0001), a main effect of rearing (p = 0.03), and an age × rearing interaction (p = 0.012) (Fig 4). Post-hoc analysis found that MS increased Kiss1 specifically at P20 in females (p = 0.014). 3-way ANOVA comparing MPOA Gnrh revealed a sex × rearing × age interaction (p = 0.047), as well as a trend-level age × rearing interaction (p = 0.052), and main effects of age (p = 0.0024) and sex (p < 0.0001). Post-hoc analysis found that MS increased Gnrh specifically at P20 in females (p = 0.01). 3-way ANOVA comparing PH Kiss1 revealed only a main effect of age (p < 0.001).

Figure 4.

Effects of maternal separation on hypothalamic Kiss1 and Gnrh gene expression (n = 9–11). Sexes are illustrated separately for clarity, with males above and females below, however main effects were determined with a sex × age × rearing ANOVA. Main effects of age, rearing, or sex are designated with asterisks, and if interactions with rearing were observed, group differences in post-hoc comparisons are shown. * p < 0.05; **p<0.001. MS: maternal separation; Con: control.; PH: posterior hypothalamus; AH: anterior hypothalamus. Individual data points with means ± SEM are shown.

Exposure to maternal separation increased number of cells expressing puberty-regulating genes and decreased the percentage of Kiss1 neurons expressing Crhr1 (See Table 2 for full statistics).

Table 2.

Full statistical information from RNAscope analyses comparing cell-specific expression of Gnrh, Kiss1, and Crhr1 in MS and Con males and females. Shaded cells and bold text indicate significant main effects or interactions

Measure	Test	Effect of Rearing
Gnrh+ cells/DAPI+ cells	Mann-Whitney	U = 7; p = 0.014
Kiss1+ cells/DAPI+ cells	Independent t	t8 = 0.434; p = 0.676 η2 = 0.022
Crhr1+ cells/DAPI+ cells	Independent t	t13 = 0.581; p = 0.571 η2 = 0.025
Gnrh + Crhr1 cells/DAPI+ cells	Mann-Whitney	U = 15; p = 0.152
Kiss1 + Crhr1 cells/DAPI+ cells	Independent t	t13 = 0.181; p = 0.859 η2 = 0.002
% Gnrh cells coexpressing Crhr1	Independent t	t12 = 0.954; p = 0.358 η2 = 0.07
% Kiss1 cells coexpressing Crhr1	Independent t	t12 = 2.0; p = 0.048 η2 = 0.287
Gnrh spot estimate	Mann-Whitney	U = 11; p = 0.051
Kiss1 spot estimate	Mann-Whitney	U = 8; p = 0.042
Crhr1 spot estimate	Mann-Whitney	U = 24; p = 0.694
Mean intensity of Gnrh	Independent t	t13 = 0.364; p = 0.722 η2 = 0.01
Mean intensity of Kiss1	Independent t	t10 = 0.739; p = 0.477 η2 = 0.05
Mean intensity of Crhr1	Independent t	t13 = 0.432; p = 0.673 η2 = 0.01
Mean Gnrh intensity in Crhr1+ vs Crhr1- cells	Mixed ANOVA	Rearing × Crhr1: F1,16 = 0.51; p = 0.48; η2 = 0.03 Rearing Main Effect: F1,16 = 2.14; p = 0.156; η2 = 0.02 Crhr1 Main Effect: : F1,16 = 12.09; p = 0.002; η2 = 0.43
Mean Kiss1 intensity in Crhr1+ vs Crhr1- cells	Mixed ANOVA	Rearing × Crhr1: F1,16 = 0.51; p = 0.48; η2 = 0.03 Rearing Main Effect: F1,16 = 2.14; p = 0.156; η2 = 0.02 Crhr1 Main Effect: : F1,16 = 12.09; p = 0.002; η2 = 0.43

RNAscope images at 20x are presented in Fig 5 to illustrate Kiss1 and Gnrh colocalization with Crhr1, and to illustrate spots and clusters. T-tests were conducted for each individual RNAscope outcome variable to determine the effects of MS at P20 in females. In order to determine whether the MS impact on puberty-regulating genes was associated with Crhr1, 2-way Mixed ANOVAs were conducted to analyze differential effects of MS on Gnrh or Kiss1 expression in Crhr1+ versus Crhr1- neurons (Fig 6). The total number of Kiss1, Gnrh, Crhr1, and colocalized Kiss1/ Crhr1 and Gnrh/ Crhr1 within the AH were counted and normalized to the DAPI count. Due to unequal variances, Mann-Whitney U was used to compare Gnrh+ cells in MS versus Con animals, as well as all spot estimate analyses. MS increased the number of Gnrh+ cells (p = 0.0168) at P20, and decreased the percentage of Kiss1 cells co-labeled for Crhr1 (p = 0.048).

Figure 5.

Representative RNAscope photomicrographs of the anterior hypothalamus at P20. (a) Crhr1 (white) expressing cells merged with DAPI (blue). Arrow indicates example Crhr1 expressing cell. (b) Kiss1 (green) expressing cells merged with DAPI (blue). Arrow indicates Kiss1 expressing cell that also expresses Crhr1. Chevron indicates Kiss1 cell that does not also express Crhr1. (c) Merged image of Crhr1 (white), Kiss1 (green), and DAPI (blue). Arrow indicates Crhr1+Kiss1+ cell. Chevron indicates Crhr1-Kiss1+ cell. (d) Crhr1 (white) expressing cells merged with DAPI (blue). Arrow indicates example Crhr1 expressing cell. (e) Gnrh (magenta) expressing cells merged with DAPI (blue). Arrow indicates Gnrh expressing cell that also expresses Crhr1. Chevron indicates Gnrh cell that does not also express Crhr1. (f) Merged image of Crhr1 (white), Gnrh (magenta), and DAPI blue. Arrow indicates Crhr1+Gnrh+ cell. Chevron indicates Crhr1-Gnrh+ cell. Scale bar = 500μm

Figure 6.

Effects of maternal separation on cells co-expressing Kiss1 or Gnrh with Crhr1 in the anterior hypothalamus at P20. (a) Kiss1 expressing cells, normalized to DAPI-stained nuclei; n = 7–8. (b) Kiss1 + Crhr1 co-expressing cells, normalized to DAPI-stained nuclei; n = 7–8. (c) Percent Kiss1-expressing cells co-expressing Crhr1; n = 6–8. (d) Gnrh expressing cells, normalized to DAPI-positive cells; n = 7–8. (e) Gnrh + Crhr1 co-expressing cells, normalized to DAPI-stained nuclei; n = 7–8. (f) Percent Gnrh-expressing cells co-expressing Crhr1; n=7. (g) Crhr1 expressing cells, normalized to DAPI-stained nuclei; n = 7–8. MS: maternal separation; Con: control. *p < 0.05. Individual data points with means ± SEM are shown.

Because the presence of multiple mRNA transcripts in close proximity appear as clusters rather than individual dots, QuPath uses estimation algorithms to provide a relative measure of the number of spots present in a cluster, dependent on spot size and cluster size. MS increased the total spot estimate of Kiss1 (p = 0.0417), with a trend-level effect on Gnrh (p = 0.051) and no effect on Crhr1 (p = 0.6943) (Fig 7a,d,g). Cell mean intensities were also calculated and compared based on rearing condition (Fig 7b,e,h). Overall intensities of Gnrh, Kiss1, and Crhr1 were not affected by MS. Intensities were then stratified to determine specific intensities of Kiss1 or Gnrh in cells co-expressing Crhr1 versus cells expressing only Kiss1 or Gnrh and analyzed with Mixed 2-way ANOVA (Fig 7c,f). While no interactions or main effects of rearing were observed, a main effect of Crhr1 co-expression revealed that Crhr1+ cells displayed lower intensities of both Gnrh and Kiss1.

Figure 7.

Effects of maternal separation on spot estimates and intensity per cell of Kiss1 or Gnrh in Crhr1 positive cells in the anterior hypothalamus at P20. (a) Kiss1 spot estimate/cell; n=7–8; *p<0.05 difference from Con. (b) Kiss1 mean intensity/cell; n = 7–8. (c) Kiss1 mean intensity in Crhr1 positive or Crhr1 negative cells; *p<0.05 main effect of CRH-R1 expression. (d) Gnrh spot estimate/cell; n = 7–8. (e) Gnrh mean intensity/cell; n = 7–8. (f) Gnrh mean intensity in Crhr1 positive or Crhr1 negative cells; *p<0.05 main effect of Crhr1 expression. (g) Crhr1 spot estimate/cell; n = 7–8. (h) Crhr1 mean intensity/cell; n = 7–8.

Pubertal timing was correlated with ASR magnitude at P34 in females

In males, there was no correlation between the age of pubertal initiation and P34 startle magnitude (R² = 0.0119, F_1,16 = 0.193; p = 0.6660) nor between the age of full preputial separation and startle (R² = 0.1327, F_1,14 = 2.142; p = 0.165) (Fig. 8a). In females, the age of vaginal pinhole appearance was negatively correlated with the magnitude of P34 acoustic startle (R² = 0.4142, F_1,19 = 13.44; p = 0.0016), as was the age of full vaginal opening with the magnitude of P34 acoustic startle R² = 0.398; F_1,19 = 12.56; p = 0.002) (Fig. 8b). Effects of MS on P34 acoustic startle in these subjects were published previously (Granata et al., 2022) but without relating to puberty.

Figure 8.

Relationship between pubertal timing and acoustic startle magnitude in adolescence. Linear regressions were run to determine relationships between the age of pubertal initiation (left) or completion (right) and acoustic startle magnitude at P34 in males (a) or females (b). Bold lines represent slopes significantly different from zero (p < 0.05).

Discussion

This study sought to examine the neuroendocrine mechanisms underlying the accelerated pubertal onset in ELA-exposed females and to test if there is a correlation between pubertal timing and acoustic startle response. We tracked external markers of pubertal initiation in MS- and Con-reared rats and related their pubertal timing to startle magnitude in an acoustic startle paradigm. In a separate cohort of rats, we determined the developmental trajectory of the expression of puberty related genes in both the anterior and posterior hypothalamus of MS- and Con-reared rats. Lastly, we performed RNAscope on female rats on P20 to determine the cell-specific co-expression of Crhr1 with Kiss1 and Gnrh.

We observed that MS-exposed females displayed an earlier age of pubertal initiation. In the present study levels of circulating hormones were not measured, however uterine weight can be used as a proxy for estrogen levels (Bray et al., 1976). Therefore, the uterine weight in females with and without a vaginal pinhole were compared, and females with a pinhole had a significantly higher uterine weight compared to those without a vaginal pinhole. These findings support that the pinhole is a valid external marker of puberty onset. We did not observe accelerated puberty in MS-exposed males, which in line with previous reports of accelerated pubertal timing in females and either no effects or delayed preputial separation in MS-exposed males (Cowan and Richardson, 2019; Grassi-Oliveira et al., 2016). This sex difference has been underexplored but could be related to a higher degree of CRH-R1 regulation of kisspeptin in females, compared to males (Rosinger et al., 2019). Differences in maternal behavior towards females versus males under adverse rearing conditions is another potential avenue for investigating female-specific effects of ELA on pubertal timing.

Some theories posit that ELA comprising deprivation early in life causes delayed puberty while exposure to threat-related ELA causes accelerated puberty; a recent meta-analysis of human studies found that ELA in general was associated with accelerated puberty, but further analysis indicated that the association was specific to experiences characterized by threat and not to experiences characterized by deprivation (Colich et al., 2020). However, studies investigating the effects of ELA on puberty in rodents are less clear. For example, in mice ELA in the form of limited bedding and nesting (Manzano Nieves et al., 2019) and neonatal lipopolysaccharide exposure (Knox et al., 2009) have been shown to delay vaginal opening, while in rats, limited bedding and nesting does not change pubertal timing in females (Eck et al 2021 (Eck et al., 2020). However, studies using MS as a model of ELA reliably demonstrate pubertal acceleration in female rats (Cowan and Richardson, 2019; Grassi-Oliveira et al., 2016). Importantly, rodent models of ELA vary in their relative contributions of deprivation, threat, and unpredictability (Brenhouse, 2023; McLaughlin et al., 2014). For example, while deprivation is thought to be the predominant experience in MS, we have previously shown that MS dams display unpredictable behavior following dam-pup reunion (Granata et al., 2022a) and MS alters development of circuits involved in threat processing (Honeycutt et al., 2020). Some children who experience early adversity may not exclusively experience deprivation or threat, but rather a combination of the two (McLaughlin and Sheridan, 2016). The inconsistency in the rodent literature about how ELA impacts puberty may reflect how different ELA paradigms encompass a range of threat and deprivation and that these models are not interchangeable. MS remains an informative paradigm for assessing how ELA leads to early puberty, however the individual contributions of threat and deprivation in this model are yet to be characterized.

While earlier vaginal opening indicates an impact of MS on an external phenotype indicating puberty initiation, effects of ELA on central mechanisms controlling puberty have not been characterized. In typically developing animals we observed a developmental increase in Kiss1 and Gnrh expression by P30 in both the PH and AH, which was expected given previous literature that these genes are more highly expressed leading up to puberty (Navarro et al., 2004; Semaan and Kauffman, 2015). While these developmental changes corroborate previous literature, a limitation in these analyses to note is that the housekeeping gene β-actin undergoes age-related changes during development, with a peak in the rat cerebrum at P10 followed by progressively decreased expression (Poddar et al., 1996). However, the increase in Kiss1 and Gnrh expression occurred earlier in MS-exposed females than in Con-reared females in the AH, and we did not observe differences in actin expression between MS and Con tissue (data not shown). These findings are important because they are the first to show that the accelerated development of external pubertal markers following MS are preceded by accelerated development at the hypothalamic level. These differences in Kiss1 and Gnrh gene expression between MS and Con females were seen in the AH, but not the PH, which is likely due to their roles in pubertal timing versus reproductive cycles respectively (Beale et al., 2014; Hu et al., 2015). Interestingly, neonatal lipopolysaccharide, which delays puberty, decreases Kiss1 expression prior to puberty (Knox et al., 2009). These findings in conjunction with our data indicate that while different forms of ELA confer different perturbations to pubertal timing, both delays and accelerations in puberty are preceded by hypothalamic Kiss1 expression changes. Future studies attempting to prevent the accelerated pubertal development in MS-exposed females can use Kiss1 and Gnrh gene expression in the AH as an outcome measure.

Given the understanding that stress decreases concurrent Gnrh and Kiss1 expression in the hypothalamus with corresponding effects on reproductive vitality and behavior via actions at CRH-R1, (Kinsey-Jones et al., 2009; Rivier et al., 1986), we hypothesized that ELA leads to earlier pubertal onset at both the level of the hypothalamus and of physical appearance via changes in Crhr1 expression. As the intensities of Kiss1 and Gnrh were lower in Crhr1 positive cells compared to negative cells, we postulate that CRH action in both MS- and Con-reared animals suppressed Kiss1 and Gnrh expression. However, since the percentage of Kiss1 cells co-expressing Crhr1 is lower in MS-females than Con-females, it is possible that a smaller percentage of Kiss1 cells have the machinery to be inhibited by CRH. Therefore, this may lead to increased overall Kiss1 production, as seen in our Kiss1 spot estimate as well as the increased Kiss1 expression in the AH using qPCR. Downstream, the excess Kiss1 production in MS animals likely leads to greater GnRH release, which we demonstrate with the increased number of Gnrh+ cells and the increase in Gnrh expression in the AH in MS-females compared to Con-females. In adult mice, CRH-R1 is more highly expressed by kisspeptin-positive neurons in females than in males in the AVPV (Rosinger et al., 2019). While we did not perform RNAscope in both males and females, the differences in baseline co-expression of Crhr1 and Kiss1 could explain why females but not males undergo early puberty after MS. These findings lead to the overall hypothesis that an MS-induced change in the proportion of kisspeptin cells expressing CRH-R1 leads to an inability of CRH to prevent kisspeptin release onto GnRH neurons; however, this hypothesis needs to be tested directly.

Because previous studies and evidence from human literature suggest that pubertal timing and ELA relate to anxiety symptomology, we revisited a dataset from some of our previously published work to correlate pubertal timing with acoustic startle response (Granata et al., 2022b). In female animals there was a significant negative correlation between day of puberty initiation and baseline startle magnitude, as well as between day of puberty completion and baseline acoustic startle magnitude. These results support evidence from the human literature that early puberty is associated with psychiatric disorders, namely depression and anxiety. For example, Toffol and colleagues (Toffol et al., 2014) demonstrated that the age of menarche correlated with psychiatric diagnosis, such that young age at menarche was associated with increased risk of recent mental health disorder, major depressive episode, major depressive disorder, and anxiety disorder. Others have shown that children undergoing precocious puberty show greater depression scores with girls especially having higher anxiety scores (Huang et al., 2021). These relationships between early puberty and psychiatric diagnoses are important to untangle, especially given the declining age of puberty onset. For example, a systematic review and meta-analysis found that the age of thelarche has decreased three months per decade from 1977 to 2013 (Eckert-Lind et al., 2020). It remains unclear if this decline in pubertal age simply calls for reassessment of what is considered precocious puberty or if this early puberty may leave individuals susceptible to experiencing anxiety.

Many environmental factors can alter pubertal timing. For example, nutritional status, particularly leptin, ghrelin, insulin, and IGF-1 can all impact puberty and reproduction status after puberty (Abreu and Kaiser, 2016). Previously we demonstrated that MS-animals weigh less than Con animals in the pre-weaning period, however they did not differ in weight by P55 (Grassi-Oliveira et al., 2016). In contrast, both mice and rats exposed to limited bedding and nesting display lower weights compared to their control reared counterparts throughout development and into adulthood (Eck et al., 2020; Manzano Nieves et al., 2019). While the present cohort of rats did not show any difference in weight during the pre-weaning period, MS rats gained significantly more weight than Con rats between P20 and P34. Indeed, greater body weight can accelerate puberty, as shown with neonatal overfeeding models (Stopa et al., 2021). It is possible that rapid weight gain leads to earlier puberty, however the hypothalamic changes in Kiss1 and Gnrh expression in this study are already apparent by P20. Furthermore, others have shown that blocking leptin delays puberty, without affecting body mass (Mela et al., 2016). While the differences in weight gain between P20 and P34 alone cannot explain the early puberty in MS animals, there is still a possibility that leptin signaling contributes to altered pubertal timing. Together, there is much to uncover about the relationships between ELA, early puberty, and metabolic processes related to weight gain.

Future Directions and Limitations

The current study assessed the transcriptional changes of puberty related genes in the hypothalamus, however, there are some limitations of this study that could be addressed in future experiments. One limitation to the present study is that the main outcome measures are RNA-based, which does not provide direct information about changes in protein levels. In particular, we found that there is a reduction in the percentage of Kiss1 cells coexpressing Crhr1, which we postulate indicates a downregulation of CRH-R1. This is plausible given that excess agonism can desensitize G-protein-coupled receptors (GPCR) through receptor phosphorylation, beta-arrestin binding, and receptor internalization, and this has been shown to be possible with excess CRH signaling at CRH-R1 (Hauger et al., 2003; Holmes et al., 2006; Perry et al., 2005; Roseboom et al., 2001; Teli et al., 2005). Other studies indicate the short-term desensitization of GPCRs can be followed by longer-term downregulation of the receptor, which involves transcriptional changes (Bouvier et al., 1989). The reduction in the percent of Kiss1 cells co-expressing Crhr1 mRNA could be due to this longer-term downregulation, however future studies could confirm that MS induces downregulation of the CRH-R1 protein and not just of the transcript. Additionally, the functional relevance of CRH-R1 protein reduction was not tested in the here, but could be tested in the future. Work in adult females shows that CRH infusion into the AVPV of the hypothalamus decreases Kiss1 expression (Kinsey-Jones et al., 2009), and that CRH administration during juvenility delays puberty (Kinsey-Jones et al., 2010). In contrast, during the MS paradigm, animals exhibit heightened CRH levels in the hypothalamus (Roque et al., 2022) which may lead to downregulation of CRH-R1 on KISS1 cells and ultimately accelerated pubertal timing. Future studies could investigate if CRH-R1 antagonism during MS off-sets the excess CRH signaling and prevents the downregulation of CRH-R1 in KISS1 cells and accelerated puberty in females. Additionally, measurement of the Kisspeptin receptor GPR54 could provide further insight into effects of ELA that we did not investigate here.

Another limitation of the present study is that we were underpowered to perform groupwise analyses based on pubertal status and therefore could not test the contributions of current pubertal status on acoustic startle response. The correlations between early puberty and acoustic startle response could be driven by hormonal surges related to puberty status at the time of testing. The presence of pubertal hormones have indeed been shown to play an important role in adolescent anxiety as measured with the marble-burying test (Boivin et al., 2017) as well as the development of adult stress-induced anxiety (Woodward et al., 2023). In humans, advanced pubertal status in girls was associated with higher social anxiety (Deardorff et al., 2007) and in pubertally advanced girls, resting state functional connectivity between the vmPFC and amygdala was negatively correlated with anxiety symptoms (Ladouceur et al., 2023). Future studies could assess acoustic startle response at different stages during puberty and correlate the response level of hormones at the time of testing. We also did not assess if the relationship between early puberty and acoustic startle response persists into later adolescence or adulthood. In the human literature, evidence about whether negative effects of early puberty persist into adulthood are contradictory, with some investigations demonstrating that the detrimental effects are attenuated in adulthood (Copeland et al., 2010) and others demonstrating lifetime prevalence of psychiatric disorders and impaired psychosocial functioning (Graber et al., 2004). Future work should assess if early puberty, induced pharmacologically or environmentally, lead to anxiety-like phenotypes in adulthood.

Conclusions

The present study replicates previous work that ELA leads to accelerated pubertal onset in females. We show for the first time that Kiss1 and Gnrh expression in the AH increases between P10 and P30 in both male and female rats, however MS-exposed females display this increase earlier than Con-reared females. On P20, MS-exposed females show a greater number of Gnrh cells, an increase in the number of Kiss1 transcripts, and a decrease in the percentage of Kiss1 cells expressing Crhr1, as compared to Con-reared females. These findings point to the possibility that ELA leads to downregulation of CRH-R1 on kisspeptin producing neurons, which may cause premature puberty onset via kisspeptin and GnRH signaling. Lastly, earlier puberty onset is associated with enhanced acoustic startle in adolescence. These results support an impact of ELA on central regulation of pubertal timing, and highlight the need to understand the mechanisms by which early puberty might lead to anxiety symptoms, especially as the average age of puberty is decreasing.