Gonadotropin-releasing hormone (GnRH) is a key regulatory molecule of the hypothalamus–pituitary (PIT)–gonadal (HPG) axis. Neurons in the hypothalamus release GnRH which acts downstream on the PIT to stimulate transcription and secretion of luteinizing hormone (LH) and follicle stimulating hormone (FSH). In turn, LH and FSH stimulate follicular maturation and release of the steroids estradiol (E 2) and progesterone (P). These steroids feed back to the HPG axis to maintain homeostatic regulation of reproductive function and behavior.
While GnRH is thought to be the master regulator of reproduction, its metabolic product GnRH-(1–5) is also shown to be biologically active. GnRH-(1–5) is produced after thimet oligopeptidase 1 (also known as EP24.15) cleaves the covalent bond linking the fifth and sixth amino acids of GnRH. GnRH-(1–5), like its parent peptide, is involved in the regulation of the HPG axis. Both GnRH gene expression and secretion are stimulated by GnRH-(1–5). Additionally, the facilitation of lordosis by GnRH is mediated by its metabolism to GnRH-(1–5). However, GnRH-(1–5) binds to alternative receptors than its parent GnRH peptide. The actions of GnRH-(1–5) are mediated through a pair of orphan G protein-coupled receptors, GPR101 and GPR173. The downstream signaling actions of GnRH-(1–5) occur through traditional G-protein signaling pathways (GPR101) and non-canonical pathways in which β-arrestin 2 is rapidly recruited (GPR173).
It is thought that during aging, changes in the hypothalamic GnRH system, as well as PIT and ovarian processes, are key components that contribute to reproductive senescence. The hypothalamic changes include decreased GnRH release and neural activation, as well as diminution of the preovulatory GnRH/LH surge. The subsequent PIT and ovarian changes result in diminished E 2 and P secretion and changes in the positive feedback system on GnRH-induced LH surge. Additionally, GnRH cleavage enzyme (EP24.15) immunoreactivity within the median eminence where GnRH axons terminate is sensitive to hormonal changes, as its expression decreases from the early proestrous period (high circulating E 2 and low LH) to the late proestrous period (low circulating E 2 and high LH). It is possible that GnRH-(1–5) may have additional peripheral effects. However, whether and how the GnRH-(1–5) signaling pathway changes during reproductive aging within the brain is unknown.
Utilizing a reproductive aging female rat model, we sought to determine the impact of age and hormone treatment duration and timing on genes crucial for GnRH-(1–5) signaling, particularly in the medial preoptic area (mPOA) and arcuate nucleus (ARC) of the hypothalamus. The mPOA is of interest as GnRH cell bodies are mainly found here, and it is a major site for the regulation of reproductive function. The ARC is responsible for regulating the negative feedback response to E 2, as well as assisting in generation of the pulsatile release of GnRH. Importantly, Gpr101, Gpr173, and EP24.15 are all expressed within these regions. In addition, gene expression within motor cortex (MC) and PIT were used as comparisons to the hypothalamus. Our goal is to provide mechanistic insights into GnRH and GnRH-(1–5) signaling pathway regulation by E 2 deficiency and treatment during reproductive aging.
Tissue samples assayed in this study were previously generated and utilized for separate publications. This study utilized the cDNA generated by these previous publications.
Female Sprague-Dawley rats (Harlan, Indianapolis, IN, USA) were purchased at 3–4 months [reproductively mature (MAT); virgin] and 10–11 months old [reproductively aging (AG); retired breeder]. On arrival, rats were pair housed at random with same-age partners in a controlled room temperature (22°C) and light cycle (12-h light, 12-h dark, lights on at 7 a.m.). Food and water were available ad libitum. All animal experiments were conducted following protocols approved by the Institutional Animal Care and Use Committee at the University of Texas at Austin and in accordance with The Guide for the Care and Use of Experimental Animals.
The animal procedures were previously described. Briefly, animals were acclimated to the new housing environment for 1 week prior to 2 weeks of estrous cycle monitoring by vaginal lavage with sterile saline. Females age 3–4 months with regular 4–5 days cycles were used for the MAT group. Females age 10–11 months with regular cycles (50%), irregular estrous cycles (30%), or persistent estrus (20%) were randomly assigned to different treatments for the AG groups. Upon determination of estrous cyclicity, all animals underwent bilateral ovariectomy (OVX) under isoflurane inhalation anesthesia, and each animal was administered a non-steroidal anti-inflammatory drug (Rimadyl; 5 mg/kg) at the beginning of surgery for analgesia. Animals were randomly assigned to one of eight treatment groups, allowing for the examination of different temporal regimens of hormone treatment. Animals within the same cage always received the same treatment.
At the time of surgery, Silastic capsules containing either 100% cholesterol (VEH) or 5% 17β-estradiol/95% cholesterol (E 2) were implanted subcutaneously between the shoulder blades. Delivery of E 2 via Silastic capsule was previously shown to last for at least 6 months without loss of integrity. After OVX, all animals received new identifiers to allow for blinded data collection. Groups 1 and 2 consisted of MAT animals treated with VEH or E 2 for 3 months, respectively (MAT-V3 and MAT-E3). Groups 3 and 4 consisted of AG animals treated with VEH or E 2 for 3 months, respectively (AG-V3 and AG-E3). Groups 5 and 6 consisted of AG animals treated with VEH or E 2 for 6 months, respectively (AG-V6 and AG-E6). Groups 7 and 8 consisted of AG animals treated with E 2 followed by VEH for 3 months each (group 7, AG-E3/V3) or VEH followed by E 2 for 3 months each (group 8, AG-V3/E3).
After 3 or 6 months of hormone treatment, animals were euthanized by rapid decapitation between 1 and 3 p.m. (4–6 h before lights off at 7 p.m.). Brains were quickly extracted and briefly cooled on ice. Coronal brain sections (eight total) were taken at 1 mm intervals throughout the entire hypothalamus using an ice-cold brain matrix (Ted Pella, Inc., Redding, CA, USA). Sections were quickly immersed in 1.5 mL of RNAlater (cat. no. AM7021M, Invitrogen, Waltham, MA, USA) and stored overnight at 4°C. After overnight storage, each section was mounted on plain glass slides and stored at −20°C before micropunching. Additionally, at time of euthanasia, the PIT was removed, immersed in RNAlater overnight at 4°C, and stored at −20°C prior to RNA extraction.
Brains treated with RNAlater were thawed once for micropunching. Hypothalamic regions containing the mPOA (bregma −0.26 to −1.80) and ARC (bregma −2.12 to −4.52) were micropunched using a 1.2-mm diameter punch (cat. no. 57399, Stoelting, Wood Dale, IL, USA). As a control (non-hypothalamic) region, MC was punched, using a 1.2-mm diameter punch, from regions related to motor control of the forelimbs and forepaws. The entire PIT was used for extraction of RNA. Total RNA, from all regions and the PIT, was extracted using the RNeasy Mini Kit (cat. no. 74104, Qiagen, Valencia, CA, USA) according to the manufacturer’s protocol. DNase digestion was performed on-column using the RNase-Free DNase Set (cat. no. 79254, Qiagen). RNA was eluted in 30 µL RNase-free water, and RNA quality and concentration were determined with the Agilent RNA 6000 Nano kit (cat. no. 5067-1511, Agilent Technologies, Santa Clara, CA, USA) on the bioanalyzer. For each region, 200 ng of total RNA was reverse-transcribed to single-stranded cDNA using a high capacity cDNA reverse transcription kit (cat. no. 4374966, Applied Biosystems, Foster City, CA, USA).
The mRNA expression of Gpr101, Gpr173, EP24.15, and the control gene Gapdh was assessed in each region by real-time PCR using the iQ SYBR Green Supermix (cat. no. 1708884, Bio-Rad, Hercules, CA, USA). Additionally, mRNA expression of Gnrh1 and Gnrhr were assessed in the mPOA and PIT, respectively. Each sample was assayed in duplicate using 400 nM of the appropriate primer pair on the CFX Connect Real-time System (Bio-Rad). Mature adult male rat hypothalamus was included in each assay as an intra- (0.281 ± 0.044%) and interassay (0.601 ± 0.084%) control. The following cycling parameters were used: initial denaturation and enzyme activation at 95°C for 3 min followed by 40 cycles of denaturation (95°C, 15 s), annealing (60°C, 30 s), extension (72°C, 30 s), and reading. Melt curve analysis was conducted after each real-time reaction to demonstrate the presence of a single amplicon. Amplified products were purified using the QIAquick PCR Purification Kit (cat. no. 28104, Qiagen) and verified post-purification by agarose gel analysis and sequenced with the ABI 3500xL Genetic Analyzer (Applied Biosystems). Sequences were verified using NCBI BLAST and comparing sequences to the Reference RNA sequences (refseq_rna) database. Primers specific for Gapdh, Gnrh1, Gnrhr, Gpr101, Gpr173, and EP24.15 are shown in Table 1. Relative expression of each gene was determined using the delta delta C t (ΔΔ C t) method, normalizing each sample to Gapdh. In a previous publication, additional brain regions from these animals were analyzed and Gapdh was used as the housekeeping gene. In both that study and the current study, there were no effects of age or E 2 treatment on the expression of Gapdh. Therefore, Gapdh was determined to be a valid normalizing gene. All data were expressed relative to the mature, vehicle-treated group (MAT-V3).
