Animals were group-housed at Baylor College of Medicine (BCM) on a 12 h light/dark cycle with ad libitum access to water and food. The mice used for viral studies were bearing a floxed VDR allele (VDR^f/f) (18, 24). Additionally, we bred Sim1-cre mice (25) bearing a Rosa26-tdTOMATO allele onto the VDR^f/f mice. For the initial phenotyping experiments, mice bearing homozygous VDR floxed alleles were mated (VDR^f/f x VDR^{f/f;Sim1-cre;TOMATO}), creating experimental PVH VDR knockdown mice (KD; VDR^f/f;Sim1-cre) and control animals (VDR^f/f), without regard for the presence or absence of the TOMATO gene. However, for electrophysiological experiments, we mated heterozygous VDR^f/+ x VDR^{f/+;Sim1-cre;TOMATO} mice in order to have VDR^{+/+;Sim1-cre;TOMATO} controls which would allow for visualization of the PVH neurons. Animal numbers are stated in the figure legends. All studies used littermate controls. Mice were euthanized with ketamine/xylazine per BCM protocols. All studies were approved by the BCM Institutional Animal Care and Use Committee (IACUC).

Animals were studied in a lean state (chow diet) or an obese state. Obesity was induced by a western diet (45% fat, D12451, Research Diets, New Brunswick, NJ) for 8 weeks prior to glucose tolerance testing.

Some mice at 8-12 weeks of age received 20nL of a replication-defective adenovirus-associated virus (AAV) containing Cre-recombinase (AAV9.CMV.HI.eGFP-Cre.WPRE.SV40 9.82e12 gc/mL; University of Pennsylvania Vector Core, Chapel Hill, NC) or its control (AAV-CMV-GFP-9 4.0e12 gc/mL) injected bilaterally into the PVN (coordinates 0.94A/P, 4.75D/V, 0.20M/L) as previously published (18).

Intraperitoneal glucose tolerance tests (i.p. GTT) were performed as previously published (26). Mice were fasted for four hours and injected i.p. with 1.5 g/kg dextrose (D20W). A tail laceration was made, and blood glucose was measured on a glucometer in duplicate prior to dextrose administration (“0” minutes) and again 15, 30, 45, 60 and 120 min after dextrose administration. The insulin tolerance test (ITT) was performed similarly. Mice were fasted for four hours and injected i.p. with 1 unit/kg Humulin R U-100 (Eli Lilly, Indianapolis, IN). Blood glucose was measured from a tail vein laceration in duplicate on a glucometer prior to and at 15, 30, 45 and 60 minutes after insulin administration. Both GTT and ITT were performed in freely moving, conscious mice.

At euthanasia, the paraventricular hypothalamic nuclei were dissected through punch biopsy microscopically and rapidly frozen with liquid nitrogen. Tissue RNA was extracted using a Qiagen RNeasy kit. cDNA was isolated and real-time quantitative PCR (qPCR) was performed using a TaqMan 7900 sequence detection system with TaqMan universal PCR master mix and TaqMan gene expression assays (all from Applied Biosystems). Relative mRNA expression for the VDR using a primer/probe targeted to exons 3 and 4 of the VDR gene (Mm00437297_m1, Applied Biosystems) was calculated relative to the housekeeping gene L32 using the ΔΔCT method. Quantification of mRNA expression was performed as previously described (27).

Electrophysiology recordings were performed as previously published (26, 28, 29). Briefly, 10-12 week old Sim1-Cre;TdTOMATO mice were used where all neurons in the PVH would be labeled. Two male and female PVH VDR knockdown mice (VDR^{f/f;Sim1-Cre;TdTomato}) and control mice (VDR^{+/+;Sim1-cre;TOMATO}) were used for recordings. Separate mice were used for baseline and 1,25D₃-activated recordings. Mice were anesthetized with isoflurane and brains were dissected rapidly and immersed in ice-cold and oxygenated cutting solutions (in mM: 10 NaCl, 195 Sucrose, 2.5 KCl, 1.25 NaH₂PO₄, 7 MgCl₂, 25 NaHCO₃, 5 glucose, 0.5 CaCl₂, 2 sodium pyruvate. balanced with 95% O₂/5% CO₂). Coronal brain slices (220 μm) containing the PVH were cut with a Microm HM 650 V vibratome (Thermo Scientific) in oxygenated cutting solution. Slices were then incubated in oxygenated artificial cerebrospinal fluid (aCSF) (in mM: 126 NaCl, 2.5 KCl, 2.4 CaCl₂, 1.2 NaH2PO₄, 1.2 MgCl₂, 11.1 glucose, and 21.4 NaHCO₃, balanced with 95% O₂/5% CO₂, pH 7.4) to recover ~25 min at 32°C and subsequently for ≧1 h at room temperature before recording.

For whole-cell recording, slices were transferred to the recording chamber at room temperature and perfused continuously with aCSF bubbled with 95% O₂/5% CO₂ to ensure adequate oxygenation of slices. tdTomato⁺ neurons and tdTomato^- neurons were identified by using epifluorescence and IR-DIC imaging on an upright microscope (Eclipse FN-1, Nikon) equipped with a moveable stage (MP-285, Sutter Instrument). Patch pipettes with resistances of 3–5 MΩ were filled with intracellular solution (adjusted to pH 7.3) containing (in mM: 128 K gluconate, 10 KCl, 10 HEPES, 0.1 EGTA, 2 MgCl2, 0.3 Na-GTP and 3 Mg-ATP). Recordings were made using a MultiClamp 700B amplifier (Axon Instrument), sampled using Digidata 1440A and analyzed offline with pClamp 10.3 software (Axon Instrument). Series resistance was monitored during the recording, and the values were generally <10 MΩ and were not compensated. The liquid junction potential was +12.5 mV, and was corrected after the experiment. Data were excluded if the series resistance increased more than 20% during the experiment or without overshoot for action potential. Currents were amplified, filtered at 1 kHz, and digitized at 20 kHz. Current clamp was engaged to test neural firing frequency at the baseline and after puff delivery (Picospritzer III, Parker Hannifin) of VDR agonist 1,25-dihydroxyvitamin D₃ (1,25D₃) (500 ms at a concentration of 1 μM) (30). The resting membrane potential and firing frequency values were averaged within 2-min bin at the baseline or after 1,25D₃ puff. In some experiments, the aCSF solution also contained 1 μm tetrodotoxin (TTX) and a cocktail of fast synaptic inhibitors, namely bicuculline (50 μM; a GABA receptor antagonist), D-AP5 (30 μM; an NMDA receptor antagonist) and CNQX (30 μM; an AMPA receptor antagonist) to block the majority of presynaptic inputs.

To measure mEPSC, the internal recording solution contained 125 mM CsCH3SO3, 10 mM CsCl, 5 mM NaCl, 2 mM MgCl2, 1 mM EGTA, 10 mM HEPES, 5 mM (Mg)ATP, 250 and 0.3 mM (Na)2GTP (pH 7.3 with NaOH) (31). The mEPSCs were recorded in whole-cell voltage-clamp mode, by holding the membrane potential a t Vh = −60 mV in the presence of 1μM TTX and 50 μM bicuculline.

For GTT, ITT, and body weight curves, data were analyzed with a paired 2-way ANOVA and Sidak post-hoc test. For AUC and qPCR measurements, data were analyzed with unpaired t-tests. The data for electrophysiology are presented individually for each recorded neuron and data were analyzed with unpaired 1-way ANOVA for baseline averages. Pre- and post-vitamin D treatment comparisons were made by paired 2-way ANOVA. P < 0.05 was considered to be statistically significant.

We demonstrated previously that male mice with PVH VDR viral knockdown had impaired glucose tolerance when obese, but not when lean (18). However, female mice with viral PVH VDR knockdown did not differ in glucose tolerance compared to control virus when lean ( Figure 1A ) or obese ( Figure 1B ). Importantly, these female mice were littermates to our previously published male cohort and thus, were of the same age and exposed to the same housing conditions as their male counterparts. Thus, external factors are unlikely to have influenced the observed sexual dimorphic difference in glucose tolerance. Furthermore, viral knockdown of VDR within the PVH in females did not alter their body weight in chow or DIO conditions ( Figure 1C ). We analyzed expression data from our previous male cohort and found that males and females had similar expression levels of VDR within the PVH and that virus administration resulted in a similar knockdown of the VDR ( Figure 1D ). Thus, the difference in glucose tolerance between male and female mice resulting from PVH VDR loss does not seem to be due to a difference in levels of the VDR but rather a sexually dimorphic function of the VDR within the PVH.

To determine if the effects of PVH VDRs were consistent across other models, we tested the effects of genetic deletion of PVH VDR. The single-minded homolog 1 gene (Sim1) is expressed significantly in the PVH and parts of the amygdala. By mating VDR^f/f mice with Sim1-Cre (25), we established mice with loss of VDR within the PVH from birth. Once again, PVH VDR knockdown did not result in any glucose or insulin tolerance abnormalities in females ( Figures 2A, B ). However, PVH VDR knockdown had a pronounced deleterious effect on glucose tolerance in males, as evidenced in their glucose curves ( Figure 2C ; main effect of genotype, p < 0.05) and AUC ( Figure 2C inset). We did not observe significant insulin resistance during an ITT ( Figure 2D ). Before starting high-fat diet, mice were tested for glucose tolerance under chow-fed conditions. We observed no effect of PVH VDR loss in either sex when animals were on chow ( Figures 2E, F ), which is similar to our viral data shown in Figure 1 and our prior publication (18). Body-weight trajectories on a high-fat diet did not differ between the groups ( Figure 2G ). VDR expression was significantly decreased in the PVH of VDR^f/f;Sim1-Cre mice but did not differ between sexes ( Figure 2H ).

We and others have published that vitamin D has rapid actions on neuronal activity (18, 30, 32). However, whether VDRs affect basal levels of neuronal function within the hypothalamus were unknown. In DIO VDR^f/fSim1-Cre mice and their controls, we tested the role of VDRs on PVH neuronal function. Loss of PVH VDR decreased the basal firing frequency in male mice but not in female mice ( Figure 3A ). However, we noticed that female control mice had lower firing frequency compared to male controls. Loss of PVH VDR also hyperpolarized neuronal membranes in male but not female mice ( Figure 3B ). We did not observe differences between baseline levels in male and female controls, indicating a role for the VDR within the PVH on neuronal function that was apparent in males but not females. Since the loss of PVH VDR decreased neuronal firing frequency and hyperpolarized neurons, we tested whether loss of PVH VDR impacts miniature excitatory post- synaptic current (mEPSC), a key factor in the generation of action potentials. Interestingly, we did not find any effect of PVH VDR on the frequency of mEPSCs in either males or females ( Figures 3C, E ). Yet, male animals with PVH VDR loss had significantly lower mEPSC amplitude while females with PVH VDR loss had no effect ( Figures 3D, E ). These results indicate that VDR is required to maintain normal post-synaptic responses of male PVH neurons to excitatory inputs (likely via glutamate), but female PVH neurons do not require this VDR function.

We previously published that vitamin D has acute activating effects on neurons in the hypothalamus (18, 26). Thus, we next tested the loss of the VDR within the PVH had any sex-specific effects on vitamin D-mediated actions on electrical activity. Active vitamin D (1,25-dihydroxyvitamin D3; 1,25D₃) increased the firing frequency in 10 of 15 (66.7%) PVH neurons (identified by Sim1-Cre mediated Tomato florescence) in males and 8 of 15 (53.3%) PVH neurons in females ( Figure 4A ). However, vitamin D had reduced effects in mice with PVH VDR KD, only activating 2 of 12 (16.5%) male neurons and 3 of 16 (18.7%) female neurons. There was no difference in the magnitude of change in firing frequency after 1,25D₃ stimulation in control male and female mice ( Figure 4B ). Similarly, 1,25D3 depolarized 10 of 15 PVH neurons in males and 8 of 15 PVH neurons in females ( Figure 4C ). Loss of PVH VDR neurons decreased the ability of 1,25D3 to depolarize PVH neurons. Only 2 of 16 neurons in males and 3 of 16 neurons in females were depolarized by vitamin D in animals with PVH VDR knockdown ( Figure 4C ). There was no difference in the change in resting membrane potential between the sexes ( Figure 4D ). To confirm that the effects of vitamin D on neuronal function were due to direct actions of vitamin D on these PVH neurons, we treated neurons from male mice with vitamin D (1,25D₃) in the presence of synaptic blockers. We found that in the presence of synaptic blockade, vitamin D depolarized 8 of 11 (72.7%) PVH neurons in control animals and was unable to depolarize any neurons in animals with PVH knockdown ( Figure 4E ). This data indicated that vitamin D (1,25D₃) directly acts on VDR in PVH SIM-1 neurons to activate their activities.