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. 2016 Jan 7;157(4):1535–1545. doi: 10.1210/en.2015-1638

Maternal Vitamin D Deficiency Programs Reproductive Dysfunction in Female Mice Offspring Through Adverse Effects on the Neuroendocrine Axis

Cari Nicholas 1, Joseph Davis 1, Thomas Fisher 1, Thalia Segal 1, Marilena Petti 1, Yan Sun 1, Andrew Wolfe 1, Genevieve Neal-Perry 1,✉
PMCID: PMC5393357  PMID: 26741195

Abstract

Vitamin D (VitD) deficiency affects more than 1 billion people worldwide with a higher prevalence in reproductive-aged women and children. The physiological effects of maternal VitD deficiency on the reproductive health of the offspring has not been studied. To determine whether maternal VitD deficiency affects reproductive physiology in female offspring, we monitored the reproductive physiology of C57BL/6J female offspring exposed to diet-induced maternal VitD deficiency at three specific developmental stages: 1) in utero, 2) preweaning, or 3) in utero and preweaning. We hypothesized that exposure to maternal VitD deficiency disrupts reproductive function in exposed female offspring. To test this hypothesis, we assessed vaginal opening and cytology and ovary and pituitary function as well as gonadotropin and gonadal steroid levels in female offspring. The in utero, preweaning, and in utero and preweaning VitD deficiency did not affect puberty. However, all female mice exposed to maternal VitD deficiency developed prolonged and irregular estrous cycles characterized by oligoovulation and extended periods of diestrus. Despite similar gonadal steroid levels and GnRH neuron density, females exposed to maternal VitD deficiency released less LH on the evening of proestrus. When compared with control female offspring, there was no significant difference in the ability of females exposed to maternal VitD deficiency to respond robustly to exogenous GnRH peptide or controlled ovarian hyperstimulation. These findings suggest that maternal VitD deficiency programs reproductive dysfunction in adult female offspring through adverse effects on hypothalamic function.

Vitamin D (VitD) is a fat-soluble secosteroid primarily obtained from dietary intake and conversion of 7-dehydrocholesterol in the skin by exposure to UV-B radiation from sunlight (1). VitD is further processed in the liver and subsequently in the kidney into its active form, 1α,25-dihydroxyvitamin D3. 1α,25-Dihydroxyvitamin D3 binds to the nuclear VitD receptor (VDR) and the membrane bound rapid response steroid receptor to regulate gene transcription and nongenomic responses, respectively (2, 3). VitD is well known for its role in the maintenance of calcium and phosphorus homeostasis for bone health, but it is hypothesized to regulate cell proliferation and differentiation and immune modulation as well as other biological processes (4–6).

VitD insufficiency and deficiency has reached near epidemic levels worldwide (7, 8). Several recent human as well as rodent studies show VDR is expressed throughout the hypothalamic-pituitary-gonadal (HPG) axis (9–12), suggesting a role for VDR signaling in reproductive function. Consistent with our hypothesis that VDR signaling regulates reproductive physiology, several studies report that VitD deficiency with coincident hypocalcemia is linked to poor reproductive outcomes. These outcomes include subfertility in males and females and is characterized by reduced pregnancy rates and smaller litter sizes (13–18). Additionally, transgenic mice with VDR deletion exhibit pubertal failure and primordial stage ovarian follicular arrest and hypergonadotropic hypogonadism (19–21). Transgenic mice with the deletion of Cyp27b1, the enzyme that regulates synthesis of the most active form of VitD, have a milder reproductive phenotype characterized by delayed puberty, oligoovulation, early stage ovarian follicular arrest, and eugonadotropism (22).

VitD deficiency is highly prevalent in pregnant women and their offspring (7). The concept that maternal under- as well as overnutrition may serve as a backdrop for developmental origins of adult disease such as diabetes, hypertensions, and obesity is well established (23–25). More recent data suggest that micronutrient deficiencies such as VitD deficiency may also serve as a backdrop for the developmental origins of a wide range of chronic diseases in adulthood that include immune dysfunction, type 2 diabetes, heart disease, and cancer (26, 27). Although VitD deficiency is associated with reproductive dysfunction in adults (28–31) and adolescents (22, 32), the effects of maternal VitD deficiency on the reproductive function of their offspring has not been investigated. This study uses cross-fostering and dietary manipulation to investigate the effect of maternal VitD deficiency during defined early developmental stages of life (in utero, preweaning, and combined in utero and preweaning) on reproductive physiology in the first generation of female offspring. This study provides novel data that suggest maternal VitD deficiency may program reproductive dysfunction in exposed female offspring.

Materials and Methods

Animal handling and dietary management

All experiments were in compliance and approved by the Institutional Animal Care and Use Committee at Albert Einstein College of Medicine of Yeshiva University. C57BL/6J male and female mice obtained from Jackson Laboratories housed in the barrier facility at 25°C on a 14-hour light, 10-hour dark schedule (6:00 am/8:00 pm). Fertile male breeders fed standard mouse chow (control; irradiated 0.81% calcium, 0.63% phosphorus, 2.2 IU/g VitD2, number 5053; Purina Mills) were bred with females maintained on a control or VitD-deficient diet (0.81% calcium, 0.63% phosphorus, 0 IU/g vitamin D2, number AIN-93M; Purina) for 7 weeks preconceptionally and subsequently throughout pregnancy and lactation. Females fed the VitD-deficient diet received 1.5% calcium gluconate dietary supplements (calcium D-gluconate monohydrate, sc-214651A; Santa Cruz Biotechnology) to maintain bone health and calcium homeostasis (22, 33). Weight and snout-to-rump lengths were measured weekly from birth to weaning, and the ponderal index was calculated (grams per cubic millimeter). Body weights were also determined in 8- to 10-week-old adult females.

Cross-fostering

Mice born to dams maintained on control diets and subsequently weaned to a control diet are defined as controls (Figure 1A). All pups were categorized by three developmental stages; in utero, preweaning, and both in utero and preweaning. Mice identified as in utero VitD deficient were generated by cross-fostering postnatal day (PND) 2 or 3 pups from VitD deficient to control dams (Figure 1B). Mice identified as preweaning VitD deficient were generated by cross-fostering PND 2 or 3 pups from control (Figure 1C) to VitD-deficient lactating dams. Mice identified as in utero and preweaning VitD deficient were maintained with their VitD-deficient mother until weaning on PND 21 (Figure 1D). On PND 21, all pups were weaned onto and maintained on a control diet until they were killed.

Figure 1. Generation of female offspring exposed to different developmental stages of VitD deficiency with cross-fostering.

Figure 1.

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A, Control mice. B, Female mice exposed to in utero VitD deficiency were generated by cross-fostering pups born to VitD-deficient dams to control dams on PND 2–3. C, Female mice exposed to preweaning VitD deficiency were generated by cross-fostering pups born to control dams to VitD-deficient dams on PND 2–3. D, Female mice exposed to in utero and preweaning VitD deficiency. On PND 21 all mice were weaned onto VitD-sufficient chow and were maintained on that diet until their death. To maintain calcium homeostasis, all mice fed VitD-deficient chow were supplemented with calcium D-gluconate monohydrate in their drinking water. Broken arrows refer to pups that were cross-fostered to another dam; solid arrows refers to pups that were maintained with birth dam.

Estradiol, Testosterone, VitD, and calcium levels during adulthood

Nonlittermate adult female offspring between 8 and 10 weeks old were killed and serum isolated and stored at −80°C until VitD and calcium levels were measured by the Einstein-Sinai Diabetes Research Center Biomarker and Analytical Research Core. VitD levels were measured by liquid chromatography/mass spectrometry with a sensitivity range of 5–100 ng/mL (34) and an intra- and interassay coefficient of variance (CV) of less than 10%. Calcium levels were measured using the Olympus calcium o-cresolphthalein-complexone absorbance assay with a sensitivity range of 0–18 mg/dL (35) and an intra- and interassay CV of less than 5%.

Serum was collected from individual 9- to 10-week-old nonlittermate female offspring killed during diestrus. Diestrous-stage estradiol and testosterone (T) were measured by the University of Virginia Center for Research in Reproduction Ligand Core using the ELISA. The intra- and interassay CV for estradiol was 6.1% and 8.9%, respectively. The intra- and interassay CV for T was 4.3% and 7.4%, respectively.

Puberty and estrous cyclicity

Pubertal transition was defined by vaginal opening and first estrus (36). Immediately after the vaginal opening, estrous cyclicity was evaluated by daily vaginal lavage and microscopic cytology observed with a light microscope set at ×10 magnification 6 d/wk for a minimum of 5 weeks. Estrous cycle stages were identified as estrus, diestrus, or proestrus (36). Estrous cycle length was measured by counting days between each proestrus event for a minimum of 5 weeks.

Ovarian histology

Groups of nonlittermate adult female mice (9–10 wk) were anesthetized using isoflurane (Butler Animal Health Supply; catalog number 029405), and the ovaries and uterus were removed in diestrus and weighed. When possible, trunk blood was collected for steroid hormone determination. Ovaries were fixed in paraformaldehyde overnight, transferred to 70% ethanol, and embedded in paraffin. Paraffin-embedded ovaries were sectioned every 10th section with a microtome and 6-μm sections collected for staining with hematoxylin and eosin. Follicles were classified and quantified using methods previously described (37). Total numbers of primordial, primary, secondary, preantral, and corpora lutea were determined by two counters blinded to the mouse diet (37). Duplicate follicle counts were avoided by counting oocytes with a visible nucleus. Sections were examined with a Zeiss AX imager microscope.

Response to ovarian superovulation

Eight- to 10-week-old nonlittermate female offspring exposed to control (n = 10) or in utero and preweaning VitD-deficient diets (n = 10) were superovulated in diestrus with 0.1 cc ip injections of equine chronic gonadotropin (eCG; 5 IU; Sigma-Aldrich) or injected with vehicle (saline) at 9:00 am, followed by 0.1 cc human chorionic gonadotropin (hCG; 5 IU; Sigma-Aldrich) 48 hours later. Mice superovulated with eCG were killed 16 hours after hCG, and oocytes deposited into the oviducts were counted by an individual blinded to the dietary status of the mouse.

GnRH neuron density

GnRH neuron density was assessed as previously described by our laboratory (38, 39). Briefly, transcardiac perfusion was performed using 4% paraformaldehyde in PBS (pH 6.8) at 8:00 pm in 9- to 10-week-old nonlittermate female offspring. Brains were collected and postfixed in 4% paraformaldehyde overnight at 4°C and then transferred into 30% sucrose. A freezing microtome was used to collect 30-μm coronal sections from each mouse between the organum vasculosum and the lamina terminalis. Sections were stored in cryoprotectant at −20°C until processed for immunolabeling with 1:5000 dilution of rabbit polyclonal GnRH antibody LR-5 provided by D. R. Benoit. To quantify GnRH immunoreactive (ir) neurons, five sections of preoptic area in the 1-in-6 series were viewed under a Zeiss Axioversion microscope by an evaluator who was blinded to the dietary groups. GnRH-ir cells were counted when the cell body was clearly identified and had brown cytoplasmic staining. The total numbers of the GnRH neurons counted per five sections are reported for the female mice exposed to the control or VitD-deficient diet in utero and before weaning.

Serum gonadotropins

Groups of 9- to 10-week-old intact female mice from different litters were killed at approximately 8:00 pm and trunk blood collected during diestrus, proestrus, or estrus. Each mouse contributed one sample per estrous stage. The serum was isolated and stored at −80°C until FSH and LH levels were quantified with the MILLIPLEX MAP mouse pituitary panel as previously described in the report by Wu et al (40) (EMD Millipore). The lower limit of detection was 9.6 pg/mL and 96 pg/mL for LH and FSH, respectively. The intra- and interassay CV for LH was 14% and 6%, respectively. The intra- and interassay CV for FSH was 9% and 6%, respectively. LH and FSH are reported in nanograms per milliliter.

GnRH-induced gonadotropin release

Nine- to 10-week-old nonlittermate female offspring were ovariectomized and allowed to recover for 1 week before steroid priming. Female offspring of dams exposed to the control and VitD-deficient diets were steroid primed with two sc injections of 2 μg estradiol benzoate at 9:00 am given 24 hours apart and 500 μg progesterone sc injection 24 hours after the last estradiol benzoate injection (41). Mice were tail vein injected with 100 ng of GnRH peptide (L7134; Sigma-Aldrich), and serial blood samples were collected 2 hours after the progesterone injection (baseline) and 30 and 60 minutes after the GnRH injection. Serum was isolated and stored at −80°C until the FSH and LH levels were evaluated.

Statistical analysis

All statistical analyses were conducted with GraphPad Prism version 6.0 for Windows (GraphPad Software). VitD, calcium, body weight, puberty, and estrous cycle lengths were analyzed using a one-way ANOVA and post hoc Tukey's multiple comparison tests. A two-way ANOVA and post hoc analysis with Sidak's multiple correction tests were used to analyze ovarian follicles. A repeated-measures ANOVA was used to analyze the FSH and LH levels for the GnRH-induced gonadotropin release and body mass index. Unpaired Student's t tests were used to analyze ovarian and uterine weight, estradiol, T, FSH, and LH in the intact females.

Results

Maternal VitD deficiency does not affect ponderal index during early development or adulthood weight

To determine the effect of maternal VitD status on the ponderal index of offspring exposed to in utero, preweaning, combined in utero and preweaning VitD-deficient or control diets, the snout-to-rump length and weight were determined. The maternal diet did not significantly affect the ponderal index at birth or preweaning (Figure 2A). The maternal diet also did not affect the weight of adult offspring (Figure 2B).

Figure 2. Maternal VitD deficiency does not affect ponderal index and adult body weight in female offspring.

Figure 2.

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A, Ponderal index of female offspring (n = 8–12). B, Weight of 8- to 10-week-old adult offspring (n = 7–10). C, Age at vaginal opening (n = 10–34). D, Age at first estrus (n = 10–34). Mean ± SEM is reported.

Maternal VitD deficiency does not affect puberty

We previously demonstrated that peripubertal VitD deficiency delayed puberty in affected female mice (22). To determine the effect of maternal VitD deficiency on the pubertal transition, we assessed the age of vaginal opening and first estrus in female offspring exposed to control diets or VitD deficiency in utero, preweaning, or in utero and preweaning. Compared with control offspring, puberty was not affected by exposure to in utero, preweaning, or in utero and preweaning VitD deficiency (Figure 2, C and D).

Estrous cyclicity and maternal VitD deficiency

Serum calcium and VitD levels were measured to determine whether maternal VitD deficiency resulted in VitD deficiency or hypocalcemia in 9- to 10-week-old adult offspring. Compared with control offspring, serum calcium and VitD levels were equivalent in adult female offspring exposed to maternal VitD deficiency (Figure 3, A and B). Daily vaginal cytology was evaluated to determine the effect of maternal diet on estrous cycle patterns in female offspring. We observed regardless of when in development maternal VitD deficiency existed (in utero, preweaning, or combined in utero and preweaning VitD), affected adult female offspring exhibited prolonged estrous cycles characterized by oligoovulation and extended periods of diestrus (Figure 4, A–H; P < .05). However, females exposed to combined in utero and preweaning VitD deficiency exhibited the most severe phenotype with a 1.6-fold increase in estrous cycle length compared with control females (Figure 4A). We therefore limited the additional studies to groups of females exposed to in utero and preweaning VitD deficiency or control diets.

Figure 3. Serum VitD and calcium levels in control and adult female offspring exposed to maternal VitD-sufficient and -deficient diets are equivalent.

Figure 3.

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Serum calcium (n = 5) (A) and VitD (n = 5) (B) levels are shown. Mean ± SEM is reported.

Figure 4. Adult female offspring exposed to VitD deficiency in utero and preweaning have abnormal estrous cycle patterns.

Figure 4.

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A, Average estrous cycle length over a minimum of 5 weeks of daily vaginal smears. Average percentage of time spent in diestrus (B), proestrus (C), and estrus (D) per 5 days during a minimum of 5 weeks' monitoring time is shown. E, Representative estrous cycle patterns of female offspring exposed to a control diet. Representative estrous cycle patterns of female offspring exposed to in utero (F), preweaning (G), and in utero and preweaning VitD deficiency (H) are shown. *, **, ****, denotes P < 0.05, P < .005, and P < .0001, respectively. Mean ± SEM is reported (n = 12–17).

Gonadal steroids, ovarian physiology, and maternal VitD deficiency

Serum estradiol and T levels in diestrous-stage adult female offspring exposed to in utero and preweaning VitD deficiency were not significantly different from control female offspring (Figure 5, A and B). Similarly, compared with controls, there was no significant difference in the weight of diestrous-stage uterus collected from adult offspring exposed to control or in utero and preweaning VitD deficiency (Figure 5C). However, ovaries collected from female offspring exposed to in utero and preweaning VitD deficiency were more than 25% heavier than those collected from control females (P < .05) (Figure 5D). Compared with control females, ovaries collected from female offspring exposed to in utero and preweaning VitD deficiency had a significant increase in the number of primordial to secondary-stage follicles (Figure 5E; P < .05). There was no significant difference in the number of preantral follicles or corpus luteum. To determine whether exposure to in utero and preweaning VitD deficiency affected ovarian responsiveness to gonadotropins, we superovulated females with eCG and triggered ovulation with hCG. Female offspring exposed to control or in utero and preweaning VitD-deficient diets exhibited a robust response to superovulation and deposited oocytes into their oviducts. However, there was no significant difference in the number of oocytes deposited into the oviduct of females exposed to in utero and preweaning VitD deficiency compared with controls (26 ± 2.0 vs 18 ± 4; n = 10, P > .05).

Figure 5. Female offspring exposed to maternal VitD deficiency have more early-stage follicles and larger ovaries than control female offspring.

Figure 5.

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A, Estradiol (n = 8–12). B, T (n = 4–5). C, Uterine weight (n = 8–10). D, Ovarian weight (n = 8–9). E, Quantity of primordial follicles, primary follicles, secondary follicles, preantral follicles, and corpus lutei (n = 12–21).

Hypothalamic-pituitary axis and maternal VitD deficiency

VDR is expressed in the pituitary and hypothalamus (9, 11, 42). To determine whether in utero and preweaning VitD deficiency exposure affects hypothalamic-pituitary function, gonadotropins levels during diestrus, proestrus, and estrus were quantified in intact female offspring. LH levels during diestrus (Figure 6A) and estrus were comparable between the two groups (Figure 6E). However, compared with control offspring, females exposed to in utero and preweaning VitD deficiency exhibited attenuated LH release on the evening of proestrus (P < .05) (Figure 6C). FSH levels were not affected (Figure 6, B, D, and F) by the maternal diet. We previously reported that GnRH neurons express VDR (22). To determine whether exposure to in utero and preweaning VitD deficiency affected GnRH neuronal density, we quantified GnRH neurons and found the density of GnRH neurons in female offspring exposed to in utero and preweaning VitD was not different from control female offspring (Figure 7, A and B). To determine whether maternal diet affected gonadotroph responsiveness to GnRH peptide, we performed a GnRH stimulation challenge with 100 ng of exogenous GnRH peptide in gonadectomized and estradiol- and progesterone-primed female offspring exposed to control or in utero and preweaning VitD-deficient diets. We did not observe a significant difference in GnRH-induced LH or FSH release between the dietary groups (Figure 7, C and D).

Figure 6. Serum LH levels on the day of proestrus are reduced in female offspring exposed to maternal VitD deficiency compared with control female offspring.

Figure 6.

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A, Diestrus LH level (n = 14–20). B, Diestrus FSH level (n = 14–22). C, Proestrus LH level (n = 4–5). D, Proestrus FSH level (n = 4–5). E, Estrus LH level (n = 10–18). F, Estrus FSH level (n = 10–18). *, P < .05. Mean ± SEM is reported.

Figure 7. GnRH density and pituitary responsiveness to exogenous GnRH peptide are not affected by maternal VitD deficiency.

Figure 7.

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A, GnRH-ir cells (n = 6–9). B, Representative sections of single-label immunohistochemistry (original magnification, ×40) showing GnRH neurons (brown cytoplasm). Hypothalamic sections reviewed corresponded to plates 25–32 of the Paxinos and Watson mouse atlas (62) and the hypothalamic region between the organum vasculosum of lamina terminalis and the medial preoptic area. Black arrows indicate GnRH-ir neurons. LH (C) and FSH (D) release in the female offspring exposed to in utero and preweaning control and VitD-deficient diets challenged with 100 ng GnRH peptide (n = 7–11). Mean ± SEM is reported.

Discussion

The present study demonstrates that female offspring exposed to maternal VitD (in utero and preweaning) deficiency have a normal pubertal transition. However, adult female offspring exposed to maternal VitD deficiency go on to exhibit estrous cycle dysfunction characterized by oligoovulation and extended periods of diestrus. Compared with control females, females offspring exposed to maternal VitD deficiency had equivalent birth, prepubertal, and adult weights as well as uterine weights and gonadal steroid levels. Although the ovaries from adult offspring exposed to maternal VitD deficiency were larger and had more early-stage follicles, they responded just as robustly as control females to superovulation. Female offspring exposed to maternal VitD deficiency also had intact negative feedback, as evidenced by equivalent serum levels of LH and FSH during diestrus and estrus. However, compared with controls, female offspring exposed to maternal VitD deficiency exhibited an attenuated preovulatory LH surge but a normal pituitary response to exogenous GnRH under estradiol positive feedback conditions. In aggregate, these data suggest maternal VitD deficiency programs estrous cycle dysfunction, in part, through adverse effects on hypothalamic function and subsequent adverse downstream effects on pituitary and ovarian physiology.

Puberty and VitD

We previously reported that peripubertal (postweaning) VitD deficiency delayed puberty (22). In contrast, female offspring in this study exposed to maternal VitD deficiency have normal puberty. These data suggest that peripubertal VitD signaling is necessary for an appropriately timed puberty in female mice. The mechanisms by which peripuberty and not maternal VitD deficiency affect puberty are unclear. A study by Walker et al (43) suggests the window of peripubertal development is characterized by increased VDR expression in the medial preoptic area, the brain region that houses GnRH neurons. These data raise the possibility that adequate peripubertal VDR signaling in GnRH neurons or other VDR-expressing neurons that affect GnRH function may be necessary for the timely activation of the HPG axis. Increased availability of VitD in female offspring weaned onto a VitD-supplemented diet most likely provided the threshold level of VDR signaling required for a normal pubertal transition in female offspring exposed to maternal VitD deficiency (22). Studies designed to characterize the reproductive phenotype of transgenic mice with conditional deletion of VDR in GnRH neurons will fill gaps in the knowledge about the interrelationship between VDR expression and signaling in GnRH neurons and pubertal timing.

VitD and adult female reproductive physiology

Chronic VitD deficiency in adulthood results in estrous cycle dysfunction characterized by extended periods of diestrus and oligoovulation (22). Similarly, females exposed to isolated in utero as well as preweaning VitD deficiency have an adult reproductive phenotype that is also characterized by extended periods of diestrus and oligoovulation. Of special note, estrous cycle dysfunction observed in adult females exposed to chronic VitD deficiency is rescued after 6–8 weeks of dietary VitD addback (22). In contrast, estrous cycle dysfunction persists in adult female offspring exposed solely to in utero as well as preweaning VitD deficiency and even with normcalcemia and VitD sufficiency. These data suggest that, to program normal adult HPG axis physiology, females must have adequate VDR signaling during the critical stages of early life development.

To further explore the effect of the maternal VitD deficiency during discrete stages of development on adult HPG axis function and to identify critical windows during which VitD supplements might prevent HPG axis dysfunction in adult offspring, we used cross-fostering. However, regardless of when in development (in utero vs preweaning vs in utero and preweaning) females were exposed to VitD deficiency, they exhibited estrous cycle dysfunction. The inability to link reproductive dysfunction to discrete periods of development most likely reflects the fact that in mice, brain development occurs both prenatally and postnatally (44).

Maternal VitD deficiency, estrous cyclicity, and ovarian follicular development

Reduced responsiveness of the pituitary to GnRH peptide or ovarian resistance to gonadotropins may disrupt the LH surge mechanism as well as estrous cycling. Analogous to control offspring, females exposed to maternal VitD deficiency respond robustly to exogenous gonadotropins. Thus, it is unlikely ovarian resistance to gonadotropins caused the abnormal LH surge pattern or estrous cycle phenotype observed in female offspring exposed to maternal VitD deficiency. Additionally, the maternal diet did not affect the baseline LH or FSH levels during diestrus or estrus or GnRH-induced LH or FSH release from female offspring. These data suggest maternal VitD deficiency did not adversely affect gonadotrope responsiveness to steroid-negative feedback conditions or GnRH peptide sensitivity during positive feedback conditions. GnRH peptide regulates FSH and LH synthesis and release from gonadotrophs in the anterior pituitary. However, it is important to note that FSH and LH synthesis and release are differentially regulated by the steroid environment as well as pulsatile release of GnRH (45). Specifically, LH synthesis and release is primarily regulated by the gonadal steroid environment and rapid pulsatile GnRH pulses (46), whereas FSH synthesis and release are primarily regulated by activin, inhibin, follistatin, and slow GnRH pulses (47, 48). Prolonged intervals of diestrus observed in mice exposed to maternal VitD deficiency with intact responsiveness to exogenous GnRH peptide suggests reduced endogenous GnRH release or altered pulse frequency (49). Reduced GnRH pulse frequency may adversely affect the LH surge amplitude and FSH release patterns (45–48). Taken together, our data suggest LH surge dysfunction observed in females exposed to maternal VitD most likely result from hypothalamic and GnRH neuron dysfunction. Future studies designed to investigate LH and FSH pulse frequency may provide insight into how maternal VitD deficiency affects estrous cycling and the preovulatory LH surge.

GnRH neurons, LH surge dysregulation, and maternal VitD deficiency

The exact mechanism by which maternal VitD deficiency programs estrous cycle and LH surge dysfunction in female offspring remains to be elucidated. As previously stated, a key observation of this study was females exposed to maternal VitD deficiency exhibited estrous cycle dysfunction and attenuated preovulatory LH surges. Attenuated preovulatory LH release is associated with reduced GnRH neuron density (50). Relevant to this point, female offspring were exposed to VitD deficiency during the following: 1) peak neurogenesis (around embryonic d 12.5 in mice) for the hypothalamic preoptic area (51), the brain region that houses GnRH and other neurons critical for reproduction; and 2) when GnRH neurons migrate to the hypothalamus (around embryonic d 16 [52]). Moreover, we found GT1–7 cells, immortalized GnRH neurons, express VDR (22), and we have preliminary data that suggest VDR colocalizes in GnRH neurons of adult mice (data not shown). We therefore determined whether maternal VitD deficiency disrupted GnRH neuronal development. Compared with control female offspring, we did not observe an effect of the maternal diet on GnRH neuron density. These data would argue against an abnormal density of GnRH neurons as the primary cause for estrous cycle or LH surge dysfunction.

A putative VDR element consensus (AGGTTC AG AGGTTCA) is found in the gene promoter region of Gnrh1 (National Center of Biotechnical Information: 14714 and Mouse Genome Informatics: 95789) using MAST in MEME Suite 4.10.1 (data not shown). Thus, it is quite possible that VitD regulates GnRH mRNA expression, GnRH peptide synthesis, and GnRH neuron function. Alternatively, a possible mechanism by which VDR may regulate GnRH neuron function is through effects on calcium channel signaling. Specifically, calcium is important for GnRH neuron function (53) and GnRH neurons express L-type calcium channels, which are regulated by VDR (54). It is possible that early-life VitD deficiency dysregulates GnRH neuron activity through programmed effects on L-type calcium channel function (45, 55, 56). Lastly, it is also possible that maternal VitD deficiency modifies GnRH neuron responsiveness to afferent input from kisspeptinergic, glutamatergic, or γ-aminobutyric acidergic neurons (57, 58). Studies that investigate the effect of VitD deficiency on GnRH neuron responsiveness, activation, and baseline electrophysiology will shed insight on acute as well as programmed effects of VDR signaling on GnRH neuron function.

Nonneuronal cells, such as semaphorin 3A-expressing endothelial cells in the median eminence, regulate morphological aspects of GnRH neurons including axon extension toward the vascular plexus of the hypophyseal portal system (59). Shortening of axons can cause reduced GnRH release into the portal system and result in reduced circulatory LH, similar to females exposed to maternal VitD deficiency. Developmental VitD deficiency is reported to reduce RhoA protein expression in the brain (60). RhoA, a small GTPase, regulates neuronal growth cone collapse by interacting with semaphorin 3A (61). Similar to our adult female offspring exposed to maternal VitD deficiency, rats treated with neutralizing antibodies for semaphorin 3A or its receptor neurophilin-1 exhibit fewer proestrous events and extended periods of diestrus (59). Hence, it is quite possible that maternal VitD deficiency may also affect the function of nonneuronal cells that influence GnRH neuron function and morphology.

Conclusion

It is well documented that undernutrition disrupts puberty and adversely affects adult reproductive capacity. Our study reports novel findings that demonstrate that maternal deficiency of the micronutrient VitD programs HPG axis dysfunction that is characterized by oligoovulation and attenuated preovulatory LH surges in intact and VitD-sufficient adult female offspring. Although maternal VitD deficiency did not adversely affect GnRH neuron density, our data suggest the abnormal reproductive phenotype of females exposed to maternal VitD deficiency reflects a hypothalamic defect that most likely is due, in part, to programmed dysregulation of GnRH neurons or neuronal populations that provide afferent inputs to GnRH neurons. Abnormal GnRH neurophysiology subsequently causes downstream pituitary defects and ovulatory dysfunction. Given the potential long-term reproductive consequences of maternal VitD deficiency, additional studies are needed to understand the mechanisms by which maternal VitD deficiency disrupts the regulation of the reproductive axis and fertility of female offspring.

Acknowledgments

Disclosure Summary: The authors have nothing to disclose.

Footnotes

Abbreviations:
CV
coefficient of variance
eCG
equine chronic gonadotropin
hCG
human chorionic gonadotropin
HPG
hypothalamic-pituitary-gonadal
ir
immunoreactive
PND
postnatal day
VDR
VitD receptor
VitD
vitamin D.

References

  • 1. Holick MF, Frommer JE, McNeill SC, Richtand NM, Henley JW, Potts JT Jr. Photometabolism of 7-dehydrocholesterol to previtamin D3 in skin. Biochem Biophys Res Commun. 1977;76:107–114. [DOI] [PubMed] [Google Scholar]
  • 2. Christakos S, Raval-Pandya M, Wernyj RP, Yang W. Genomic mechanisms involved in the pleiotropic actions of 1,25-dihydroxyvitamin D3. Biochem J. 1996;316(Pt 2):361–371. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 3. Mizwicki MT, Norman AW. The vitamin D sterol-vitamin D receptor ensemble model offers unique insights into both genomic and rapid-response signaling. Science Signal. 2009;2:re4. [DOI] [PubMed] [Google Scholar]
  • 4. Cui X, McGrath JJ, Burne TH, Mackay-Sim A, Eyles DW. Maternal vitamin D depletion alters neurogenesis in the developing rat brain. Int J Dev Neurosci. 2007;25:227–232. [DOI] [PubMed] [Google Scholar]
  • 5. Deluca HF. History of the discovery of vitamin D and its active metabolites. BoneKEy Rep. 2014;3:479. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 6. Eyles D, Brown J, Mackay-Sim A, McGrath J, Feron F. Vitamin D3 and brain development. Neuroscience. 2003;118:641–653. [DOI] [PubMed] [Google Scholar]
  • 7. Johnson DD, Wagner CL, Hulsey TC, McNeil RB, Ebeling M, Hollis BW. Vitamin D deficiency and insufficiency is common during pregnancy. Am J Perinatol. 2011;28:7–12. [DOI] [PubMed] [Google Scholar]
  • 8. van Schoor NM, Lips P. Worldwide vitamin D status. Best Pract Res Clin Endocrinol Metab. 2011;25:671–680. [DOI] [PubMed] [Google Scholar]
  • 9. Perez-Fernandez R, Alonso M, Segura C, Munoz I, Garcia-Caballero T, Diguez C. Vitamin D receptor gene expression in human pituitary gland. Life Sci. 1997;60:35–42. [DOI] [PubMed] [Google Scholar]
  • 10. Shahbazi M, Jeddi-Tehrani M, Zareie M, et al. Expression profiling of vitamin D receptor in placenta, decidua and ovary of pregnant mice. Placenta. 2011;32:657–664. [DOI] [PubMed] [Google Scholar]
  • 11. Veenstra TD, Prufer K, Koenigsberger C, Brimijoin SW, Grande JP, Kumar R. 1,25-Dihydroxyvitamin D3 receptors in the central nervous system of the rat embryo. Brain Res. 1998;804:193–205. [DOI] [PubMed] [Google Scholar]
  • 12. Blomberg Jensen M, Nielsen JE, Jorgensen A, et al. Vitamin D receptor and vitamin D metabolizing enzymes are expressed in the human male reproductive tract. Hum Reprod (Oxford, England). 2010;25:1303–1311. [DOI] [PubMed] [Google Scholar]
  • 13. Hickie JP, Lavigne DM, Woodward WD. Reduced fecundity of vitamin D deficient rats. Comp Biochem Physiol A Comp Physiol. 1983;74:923–925. [DOI] [PubMed] [Google Scholar]
  • 14. Johnson LE, DeLuca HF. Reproductive defects are corrected in vitamin d-deficient female rats fed a high calcium, phosphorus and lactose diet. J Nutr. 2002;132:2270–2273. [DOI] [PubMed] [Google Scholar]
  • 15. Halloran BP, DeLuca HF. Effect of vitamin D deficiency on fertility and reproductive capacity in the female rat. J Nutr. 1980;110:1573–1580. [DOI] [PubMed] [Google Scholar]
  • 16. Kwiecinksi GG, Petrie GI, DeLuca HF. 1,25-Dihydroxyvitamin D3 restores fertility of vitamin D-deficient female rats. Am J Physiol. 1989;256:E483–E487. [DOI] [PubMed] [Google Scholar]
  • 17. Kwiecinski GG, Petrie GI, DeLuca HF. Vitamin D is necessary for reproductive functions of the male rat. J Nutr. 1989;119:741–744. [DOI] [PubMed] [Google Scholar]
  • 18. Uhland AM, Kwiecinski GG, DeLuca HF. Normalization of serum calcium restores fertility in vitamin D-deficient male rats. J Nutr. 1992;122:1338–1344. [DOI] [PubMed] [Google Scholar]
  • 19. Yoshizawa T, Handa Y, Uematsu Y, et al. Mice lacking the vitamin D receptor exhibit impaired bone formation, uterine hypoplasia and growth retardation after weaning. Nat Genet. 1997;16:391–396. [DOI] [PubMed] [Google Scholar]
  • 20. Kinuta K, Tanaka H, Moriwake T, Aya K, Kato S, Seino Y. Vitamin D is an important factor in estrogen biosynthesis of both female and male gonads. Endocrinology. 2000;141:1317–1324. [DOI] [PubMed] [Google Scholar]
  • 21. Johnson AL, Zinser GM, Waltz SE. Loss of vitamin D receptor signaling from the mammary epithelium or adipose tissue alters pubertal glandular development. Am J Physiol Endocrinol Metab. 2014;307:E674–E685. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 22. Dicken CL, Israel DD, Davis JB, et al. Peripubertal vitamin D(3) deficiency delays puberty and disrupts the estrous cycle in adult female mice. Biol Reprod. 2012;87:51. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 23. Barker DJ, Osmond C, Forsen TJ, Kajantie E, Eriksson JG. Trajectories of growth among children who have coronary events as adults. N Engl J Med. 2005;353:1802–1809. [DOI] [PubMed] [Google Scholar]
  • 24. El Hajj N, Schneider E, Lehnen H, Haaf T. Epigenetics and life-long consequences of an adverse nutritional and diabetic intrauterine environment. Reproduction. 2014;148:R111–R120. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 25. Hales CN, Barker DJ. The thrifty phenotype hypothesis. Br Med Bull. 2001;60:5–20. [DOI] [PubMed] [Google Scholar]
  • 26. Harvey L, Burne TH, McGrath JJ, Eyles DW. Developmental vitamin D3 deficiency induces alterations in immune organ morphology and function in adult offspring. J Steroid Biochem Mol Biol. 2010;121:239–242. [DOI] [PubMed] [Google Scholar]
  • 27. Kaludjerovic J, Vieth R. Relationship between vitamin D during perinatal development and health. J Midwifery Womens Health. 2010;55:550–560. [DOI] [PubMed] [Google Scholar]
  • 28. Luk J, Torrealday S, Neal Perry G, Pal L. Relevance of vitamin D in reproduction. Hum Reprod (Oxford, England). 2012;27:3015–3027. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 29. Mahmoudi T. Genetic variation in the vitamin D receptor and polycystic ovary syndrome risk. Fertil Steril. 2009;92:1381–1383. [DOI] [PubMed] [Google Scholar]
  • 30. Ozkan S, Jindal S, Greenseid K, Shu J, Zeitlian G, Hickmon C, Pal L. Replete vitamin D stores predict reproductive success following in vitro fertilization. Fertil Steril. 2010;94:1314–1319. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 31. Paffoni A, Ferrari S, Vigano P, et al. Vitamin D deficiency and infertility: insights from in vitro fertilization cycles. J Clin Endocrinol Metab. 2014;99:E2372–E2376. [DOI] [PubMed] [Google Scholar]
  • 32. Villamor E, Marin C, Mora-Plazas M, Baylin A. Vitamin D deficiency and age at menarche: a prospective study. Am J Clin Nutr. 2011;94:1020–1025. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 33. Panda DK, Miao D, Bolivar I, et al. Inactivation of the 25-hydroxyvitamin D 1α-hydroxylase and vitamin D receptor demonstrates independent and interdependent effects of calcium and vitamin D on skeletal and mineral homeostasis. J Biol Chem. 2004;279:16754–16766. [DOI] [PubMed] [Google Scholar]
  • 34. Rogatsky E, Browne S, Cai M, Jayatillake H, Stein D. Quantitative analysis of 25-OH vitamin D using supported liquid extraction and liquid chromatography-mass spectrometry. J Chromatograph Separat Techniq. 2014;5(3):1–5. [Google Scholar]
  • 35. Stern J, Lewis WH. The colorimetric estimation of calcium in serum with ocresolphthalein complexone. Clin Chim Acta. 1957;2:576–580. [DOI] [PubMed] [Google Scholar]
  • 36. Cohen PE, Zhu L, Nishimura K, Pollard JW. Colony-stimulating factor 1 regulation of neuroendocrine pathways that control gonadal function in mice. Endocrinology. 2002;143:1413–1422. [DOI] [PubMed] [Google Scholar]
  • 37. Myers M, Britt KL, Wreford NG, Ebling FJ, Kerr JB. Methods for quantifying follicular numbers within the mouse ovary. Reproduction. 2004;127:569–580. [DOI] [PubMed] [Google Scholar]
  • 38. Sun Y, Todd BJ, Thornton K, Etgen AM, Neal-Perry G. Differential effects of hypothalamic IGF-I on gonadotropin releasing hormone neuronal activation during steroid-induced LH surges in young and middle-aged female rats. Endocrinology. 2011;152:4276–4287. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 39. Neal-Perry G, Yao D, Shu J, Sun Y, Etgen AM. Insulin-like growth factor-I regulates LH release by modulation of kisspeptin and NMDA-mediated neurotransmission in young and middle-aged female rats. Endocrinology. 2014;155:1827–1837. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 40. Wu S, Divall S, Hoffman GE, Le WW, Wagner KU, Wolfe A. Jak2 is necessary for neuroendocrine control of female reproduction. J Neurosci. 2011;31:184–192. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 41. Neal-Perry GS, Zeevalk GD, Santoro NF, Etgen AM. Attenuation of preoptic area glutamate release correlates with reduced luteinizing hormone secretion in middle-aged female rats. Endocrinology. 2005;146:4331–4339. [DOI] [PubMed] [Google Scholar]
  • 42. Eyles DW, Smith S, Kinobe R, Hewison M, McGrath JJ. Distribution of the vitamin D receptor and 1α-hydroxylase in human brain. J Chem Neuroanat. 2005;29:21–30. [DOI] [PubMed] [Google Scholar]
  • 43. Walker DM, Juenger TE, Gore AC. Developmental profiles of neuroendocrine gene expression in the preoptic area of male rats. Endocrinology. 2009;150:2308–2316. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 44. Luskin MB. Restricted proliferation and migration of postnatally generated neurons derived from the forebrain subventricular zone. Neuron. 1993;11:173–189. [DOI] [PubMed] [Google Scholar]
  • 45. Kirk SE, Dalkin AC, Yasin M, Haisenleder DJ, Marshall JC. Gonadotropin-releasing hormone pulse frequency regulates expression of pituitary follistatin messenger ribonucleic acid: a mechanism for differential gonadotrope function. Endocrinology. 1994;135:876–880. [DOI] [PubMed] [Google Scholar]
  • 46. Burger LL, Haisenleder DJ, Marshall JC. GnRH pulse frequency differentially regulates steroidogenic factor 1 (SF1), dosage-sensitive sex reversal-AHC critical region on the X chromosome gene 1 (DAX1), and serum response factor (SRF): potential mechanism for GnRH pulse frequency regulation of LH beta transcription in the rat. Endocrine. 2011;39:212–219. [DOI] [PubMed] [Google Scholar]
  • 47. McNeilly AS, Crawford JL, Taragnat C, Nicol L, McNeilly JR. The differential secretion of FSH and LH: regulation through genes, feedback and packaging. Reproduction (Cambridge, England) Suppl. 2003;61:463–476. [PubMed] [Google Scholar]
  • 48. Burger LL, Haisenleder DJ, Aylor KW, Marshall JC. Regulation of intracellular signaling cascades by GNRH pulse frequency in the rat pituitary: roles for CaMK II, ERK, and JNK activation. Biol Reprod. 2008;79:947–953. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 49. Levine JE, Ramirez VD. Luteinizing hormone-releasing hormone release during the rat estrous cycle and after ovariectomy, as estimated with push-pull cannulae. Endocrinology. 1982;111:1439–1448. [DOI] [PubMed] [Google Scholar]
  • 50. Herbison AE, Porteous R, Pape JR, Mora JM, Hurst PR. Gonadotropin-releasing hormone neuron requirements for puberty, ovulation, and fertility. Endocrinology. 2008;149:597–604. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 51. Finlay BL, Darlington RB. Linked regularities in the development and evolution of mammalian brains. Science. 1995;268:1578–1584. [DOI] [PubMed] [Google Scholar]
  • 52. Schwanzel-Fukuda M, Pfaff DW. Origin of luteinizing hormone-releasing hormone neurons. Nature. 1989;338:161–164. [DOI] [PubMed] [Google Scholar]
  • 53. Nunemaker CS, DeFazio RA, Moenter SM. Calcium current subtypes in GnRH neurons. Biol Reprod. 2003;69:1914–1922. [DOI] [PubMed] [Google Scholar]
  • 54. Gezen-Ak D, Dursun E, Yilmazer S. The effects of vitamin D receptor silencing on the expression of LVSCC-A1C and LVSCC-A1D and the release of NGF in cortical neurons. PLoS One. 2011;6:e17553. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 55. Krsmanovic LZ, Stojilkovic SS, Merelli F, Dufour SM, Virmani MA, Catt KJ. Calcium signaling and episodic secretion of gonadotropin-releasing hormone in hypothalamic neurons. Proc Natl Acad Sci USA. 1992;89:8462–8466. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 56. Nunez L, Villalobos C, Boockfor FR, Frawley LS. The relationship between pulsatile secretion and calcium dynamics in single, living gonadotropin-releasing hormone neurons. Endocrinology. 2000;141:2012–2017. [DOI] [PubMed] [Google Scholar]
  • 57. Feron F, Burne TH, Brown J, et al. Developmental vitamin D3 deficiency alters the adult rat brain. Brain Res Bull. 2005;65:141–148. [DOI] [PubMed] [Google Scholar]
  • 58. Brewer LD, Thibault V, Chen KC, Langub MC, Landfield PW, Porter NM. Vitamin D hormone confers neuroprotection in parallel with downregulation of L-type calcium channel expression in hippocampal neurons. J Neurosci. 2001;21:98–108. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 59. Giacobini P, Parkash J, Campagne C, et al. Brain endothelial cells control fertility through ovarian-steroid-dependent release of semaphorin 3A. PLoS Biol. 2014;12:e1001808. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 60. Almeras L, Eyles D, Benech P, et al. Developmental vitamin D deficiency alters brain protein expression in the adult rat: implications for neuropsychiatric disorders. Proteomics. 2007;7:769–780. [DOI] [PubMed] [Google Scholar]
  • 61. Wu KY, Hengst U, Cox LJ, et al. Local translation of RhoA regulates growth cone collapse. Nature. 2005;436:1020–1024. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 62. Paxinos G, Watson C. The Rat Brain in Stereotaxic Coordinates. Elsevier Academic Press; 2007:6. [Google Scholar]

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