Withania somnifera as a potential candidate to ameliorate high fat diet-induced anxiety and neuroinflammation

The leaves were collected from the seed-raised W. somnifera plant growing at the herbarium of the Department of Botanical and Environmental Sciences, Guru Nanak Dev University, Amritsar. The plant was identified and authenticated by the plant taxonomist and was deposited for reference purpose with voucher specimen number 401-24/07/1982. The previous studies from our lab have used water extract from the leaves of W. somnifera plant (ASH-WEX) [21‐23], but in the current study, we have used dry leaf powder of W. somnifera (ASH). The dose of ASH powder has been calculated based on previous in vivo reports from our lab [22, 24] using ASH-WEX (dry weight) at a dose of 140 mg/kg body weight of the animal. One milligram dry weight of ASH-WEX is equivalent to 6.80 mg of dry leaf powder, so the dose corresponds to 952 mg/kg body weight or ~ 1 g/kg body weight or 1 mg/g body weight. ASH-WEX has been standardized and characterized in the previous studies from our lab [24, 25].

Wistar albino young female rats in the age group of 3–4 months and weighing 130–150 g were used for all the experiments. The animals were housed in the group of three animals per cage under controlled environmental conditions (temperature 25 ± 2 °C and light/dark cycle of 12:12 h) with ad libitum supply of food and water. Animal care and procedures were followed in accordance with the guidelines of Institutional Animal Ethical Committee. The animals were categorized randomly into four groups: low fat diet group (LFD) fed with regular chow feed (2.5–5% fat, 75–77.5% carbohydrates, 20% proteins as macronutrients), high fat diet group (HFD) fed with diet containing 30% fat by weight (30% fat, 50% carbohydrates, 20% proteins as macronutrients), low fat diet plus extract group (LFDE) fed with regular chow feed supplemented with ASH, and high fat diet plus extract group (HFDE) fed with diet containing 30% fat by weight and supplemented with ASH (n = 10 ± 1 animals per group). Regular chow feed and high fat diet were mixed with ASH at a concentration of 1 mg/g body weight of animal. All animals were kept on the respective dietary regimen for a period of 12 weeks. The body weight of animals was monitored every week. Daily food intake for animals of all groups was also recorded. LFDE group was included in our study to elucidate any additional benefits of ASH in rats kept on normal diet. Since no significant changes were observed in the LFDE group, only preliminary data has been presented for this group.

The elevated plus maze (EPM) is a plus-shaped apparatus consisting of two opposing closed arms (50 cm × 10 cm × 40 cm), two opposing open arms (50 cm × 10 cm), and a central open area, elevated 50 cm above the ground. Individual rat from each group was placed in the central open area, facing one of the open arms at the beginning of the test. It was allowed to freely explore the maze for the duration of 5 min. The whole experiment was set up in dim light conditions and without any human disturbance. The behavior of each rat was recorded in a video camera. The whole apparatus was wiped with 70% ethanol before placing the next rat in the maze. Video recording of each rat was later analyzed for the number of crossings, number of entries, total time spent in each of the arms, and total number of head dips. An entry into an arm was counted only when all the four limbs of the rat were in that arm. A crossing was defined as the entry of a rat from an arm directly to its opposing arm. Also, total time spent in open and closed arms was recorded. A head dip was defined as looking down from the edge of an open arm or from central open area.

After subjecting the animals to EPM test, animals were sacrificed and used for immunohistochemical staining (n = 3–4 each group), Western blot analysis (n = 3–4 each group), and quantitative real-time PCR analysis (n = 3–4 each group).

For the expression study of GFAP, the animals (n = 3 each group) were perfused transcardially with 4% paraformaldehyde (PFA) in phosphate-buffered saline (PBS, 0.1 M). Brains were dissected out and stored in fixative (4% PFA) overnight at 4 °C. Subsequently, the brains were cryopreserved in grades of sucrose from 10–30% for 24 h each at 4 °C. Coronal sections of 35 μm thickness were cut using a cryostat microtome (Thermo Scientific Shandon, Waltham, MA, USA). The sections were collected in a petri dish and were given three washes with 0.1 M PBS pH 7.4, 15 min each, followed by incubation with 2% H₂O₂ for 15 min. The sections were then incubated with 0.3% Triton X-100 in 0.1 M PBS for 30 min for permeabilization followed by washing with phosphate-buffered saline with 0.1% Triton X-100 (0.1% PBST) thrice for 5 min each and incubation in 5% normal goat serum (NGS) in 0.1 M PBS for 30 min. The sections were then incubated with an antibody, i.e., rabbit IgG anti-GFAP (1:500) (Sigma Aldrich, St. Louis, MO, USA) in 5% NGS for 48 h at 4 °C. Sections were washed with 0.1% PBST thrice for 5 min each and incubated with a secondary antibody, i.e., goat anti-rabbit IgG-biotin conjugate (1:400) (Sigma Aldrich, St. Louis, MO, USA) in 0.1% PBST for 2 h at room temperature. Sections were washed with 0.1% PBST thrice for 5 min each. Further, the sections were incubated with ExtrAvidin-Peroxidase (1:400) (Sigma Aldrich, St. Louis, MO, USA) in 0.1% PBST for 2 h at room temperature, followed by diaminobenzidine (DAB) (Sigma Aldrich, St. Louis, MO, USA) for 15–20 min. Sections were mounted on the glass slide and air-dried overnight followed by dehydration through grades (50, 70, 90, and 100%) of ethanol. Tissue sections were then coverslipped using DPX permanently. Images of the sections were captured using Life Technologies EVOS FL microscope. To ensure the specificity of GFAP immunostaining, negative control was used in which the incubation with primary antibody (rabbit IgG anti-GFAP) was omitted.

Animals (n = 3–4 each group) were anaesthetized with sodium thiopentone injection and sacrificed by decapitation. Brains were dissected to isolate PC, hippocampus, and hypothalamus regions, which were homogenized in a buffer (1X Tris buffered saline, dithiothreitol (DTT), Na₃VO₄, protease inhibitor) using a tissue homogenizer at 4 °C. The homogenate was centrifuged at 10,000g for 10 min at 4 °C. The supernatant was separated and used for protein estimation by Bradford method. Equal quantity of protein from each sample was mixed with 6X sample buffer (0.25 M Tris-HCl (pH 6.8), 20% glycerol, 4% sodium dodecyl sulfate (SDS), 10% β-mercaptoethanol, and 1 mg bromophenol blue) and boiled for 2–3 min. 30-50 μg of protein sample was loaded in each well for electrophoretic separation by SDS-PAGE in 10–15% gels, followed by transfer onto a PVDF membrane (Hybond-P, GE Healthcare, USA) using the semidry TE70 PWR system (Amersham Biosciences, USA). Subsequently, the membrane was blocked (5% skimmed milk in 0.1% TBS-Tween 20) and immediately incubated with respective primary antibody (mouse anti-GFAP (1:2000); mouse anti-TNFα (1:1000); mouse anti-IL-1β (1:1000); mouse anti-IL-6 (1:1000); mouse anti-Bcl-xL (1:2000); rabbit anti-AP-1 (1:2000) (Sigma Aldrich, St. Louis, MO, USA); mouse anti-Iba1 (1:1000) (EMD Millipore, CA, USA); rabbit anti-OB-Rb (1:1000); mouse anti-PPARγ (1:1000) (Santa Cruz Biotechnology, Dallas, TX, USA); rabbit anti-phospho IKKα/β (1:1500); mouse anti-IKKα (1:1500); mouse anti-IKβα (1:1500); rabbit anti-NF-κB (1:1500) (NF-κB Pathway Sampler Kit, Cell Signaling Technology, MA, USA)) at 4 °C overnight, followed by three washings with 0.1% TBST and incubation with respective secondary antibody (goat anti-mouse for GFAP, TNFα, IL-1β, IL-6, Bcl-xL, Iba1, PPARγ, IKKα, and IKβα and goat anti-rabbit for AP-1, OB-Rb, phospho IKKα/β, and NF-κB) conjugated with horse radish peroxidase (HRP) (1:5000) (Merck Millipore, USA) for 2 h at room temperature. The membranes were also probed with anti-α-tubulin antibody (1:5000) (Sigma Aldrich, St. Louis, MO, USA) as an internal control for protein loading. Immunoreactive bands on membrane were visualized by ECL Prime Western Blotting Detection Reagent (GE Healthcare, Little Chalfont, UK) according to the manufacturer’s instructions using Image Quant LAS 4000 system (GE Healthcare, Little Chalfont, UK). Intensity analysis was done using AlphaEase FC software (Alpha Innotech, CA, USA). Change in expression of proteins of interest was taken as the average of integrated density values (IDV) obtained from at least three independent experiments.

Total RNA was extracted from the tissues using TRI reagent (Sigma Aldrich, St. Louis, MO, USA) according to the manufacturer’s instructions. For cDNA synthesis, 5 μg of RNA, 1 μL of 250 ng random hexamers (Life Technologies, Carlsbad, CA, USA), 1 μL of 10 mM dNTP mix (Sigma Aldrich, St. Louis, MO, USA), and nuclease-free sterile water up to 12 μL were added. The mixture was heated to 65 °C for 5 min and quick-chilled on ice. 4 µL of 5X first-strand buffer, 2 μL of 0.1 M DTT, and 1 μL of recombinant ribonuclease inhibitor (40 units/μL) (Life Technologies, Carlsbad, CA, USA) were added and incubated at 37 °C for 2 min. Then, 1 μL (200 units) of M-MLV reverse transcriptase (Life Technologies, Carlsbad, CA, USA) was added for 20 μL reaction volume. The mixture was kept at 25 °C for 10 min and at 37 °C for 50 min followed by inactivation of reaction by heating at 70 °C for 15 min in a thermal cycler. 50 µg of cDNA was then used to amplify gene of interest using Step One Plus Real Time PCR system (Applied Biosystems, Foster City, CA, USA). Each 5 μL reaction mixture comprised of 2.5 μL of 2X Power-SYBR Green Master Mix (Applied Biosystems, Foster City, CA, USA), 1 μL of 20X pre-designed Primer mix (Integrated DNA Technologies, Coralville, IA, USA), 1 μL of water, and 0.5 μL (50 μg) cDNA. GAPDH was used as an endogenous control for each gene of interest. The relative gene expression of each gene was calculated by “Livak method” [26] and represented as \( {2}^{-\Delta \Delta {\mathrm{C}}_{\mathrm{T}}} \).

The levels of pro-inflammatory cytokines TNFα, IL-1β, and IL-6 along with leptin and insulin were assayed using sandwich ELISA kits. The kits for inflammatory cytokines were procured from Sigma Aldrich (St. Louis, MO, USA) and for leptin and insulin from EMD Millipore Corporation (USA). The estimations were done based on colorimetric detection according to manufacturer’s protocol. The concentrations of TNFα, IL-1β, IL-6, leptin, and insulin in each sample were calculated using kit provided standards and Sigma Plot software.

A gradual increase in the body weight was observed in the rats of LFD, HFD, and HFDE group over the period of 12 weeks. On the completion of respective regimen for 12 weeks, LFD and HFD rats weighed 16.7% and 40.1%, respectively, more than their initial weight (Fig. 1a). HFDE rats also weighed 23.9% more than their initial weight, but on the contrary, LFDE rats showed 10.2% reduction of body weight after 12 weeks of feeding ASH-supplemented normal chow diet. Further, the amount of calories from fat was also calculated for each week. Highest calorie intake was observed in the HFDE group over the period of 12 weeks (Fig. 1b), though it was reduced by 40.63% at the end of the 12th week as compared to initial intake. The HFD group also showed gradual reduction in calorie intake over the 12-week period. However, LFD and LFDE groups showed consistent calorie intake from fat during the respective regimens of 12 weeks.

Prior to the EPM test, the animals were not given any training. Among the four groups, HFD rats spent minimum time in the open arm and maximum time in the closed arm as compared to LFD, LFDE, or HFDE rats (Fig. 1c). The HFDE group showed EPM profile comparable to LFD group. Further, HFD animals also showed less number of entries in both open and closed arms as compared to LFD animals (Fig. 1d). The number of crossings in the open arm was also reduced in HFD animals as compared to LFD animals (Fig. 1e). This behavior was reversed in HFDE animals, which showed highest number of entries and significantly high number of crossings (p ≤ 0.05) into the closed arm. Further, the number of head dips, which is an indicator of anxiety in rodents, was also reduced in HFD animals as compared to LFD animals, while HFDE animals showed head dips comparable to LFD animals (Fig. 1f). Overall, the behavior of LFDE animals was comparable to LFD animals.

Glial cells play an indispensable role during generation of immune response under various stressed conditions. High fat diet feeding led to upregulation in the expression of GFAP, an astroglial marker, in both hippocampus and PC regions of the brain (Fig. 2a). Western blotting and real-time PCR data also supported the immunostaining data of upregulation of GFAP in hippocampus and PC regions of HFD animals (Fig. 2b–d). The supplementation of high fat diet with ASH suppressed the change in GFAP expression in HFDE animals as observed by immunostaining, Western blotting, and real-time PCR study (Fig. 2). The expression of GFAP in the LFDE group was not different from the LFD group. These results are suggestive of reactive gliosis in HFD animals, which was effectively suppressed in ASH-fed HFDE animals. Reactive gliosis may be one of the initial events leading to chronic immune activation and inflammatory response. So, we further studied the expression of some inflammatory markers by both Western blotting and real-time PCR.

The expression of Iba1, a microglial cell-specific marker, was upregulated by 19.9% in the hippocampus region (p ≤ 0.05) as compared to LFD animals (Fig. 2b, c). The treatment with ASH suppressed the expression of Iba1 in HFDE group. PPARγ, which is a nuclear receptor controlling the transcription of genes responsible for growth and differentiation of adipocytes, showed significant upregulation (p ≤ 0.05) in the hippocampus region, whereas its expression was alleviated in the HFDE group (Fig. 2b, c).

Further, the mRNA levels of various inflammatory markers were studied by quantitative real-time PCR (Fig. 2d). ITGAM, which is a gene coding for Cd11b on macrophages/microglia, showed 1.6-fold upregulation in the hippocampus region of HFD animals, which was reduced to 1.2-fold in the HFDE group as compared to LFD animals. The mRNA levels of other inflammatory markers, such as iNOS (inducible nitric oxide synthase), MCP-1 (monocyte chemoattractant protein-1), and COX2 (Cyclooxygenase-2) also showed upregulation in the hippocampus region of HFD animals, which was alleviated in the HFDE group. Overall, the mRNA levels of different markers in the LFDE group were similar to the LFD group.

Further, the expression of Iba1 showed 23.5% increase in the PC region of HFD animals (p ≤ 0.05), which was suppressed with ASH supplementation in HFDE animals (Fig. 2b, c). The expression of PPARγ was not different in the PC region between the LFD and HFD group animals; however, it was reduced significantly (p ≤ 0.05) in HFDE animals (Fig. 2b, c). However, no difference in the mRNA levels of ITGAM, iNOS, MCP-1, and COX2 was observed in the PC region among the different groups of animals (Fig. 2d).

Since the preliminary data showed induction of inflammation in HFD animals, we further extended the work to study the expression of pro-inflammatory cytokines TNFα, IL-1β, and IL-6 by ELISA, Western blotting, and quantitative real-time PCR (Fig. 3). In ELISA-based assays, the circulating levels of TNFα, IL-1β, and IL-6 showed significant increase (p ≤ 0.001) in the HFD group animals as compared to the LFD group, which were alleviated in the HFDE animals (Fig. 3a). Further, the expression of TNFα, IL-1β, and IL-6 also showed significant increase (p ≤ 0.05) in the hippocampus region of the HFD group as compared to the LFD group at both translational (Western blotting Fig. 3b, c) and transcriptional levels (quantitative real-time PCR data Fig. 3d). However, the supplementation of ASH suppressed the HFD-induced increase of pro-inflammatory cytokines in the HFDE group (Fig. 3b–d). The expression of TNFα also showed upregulation in HFD animals as compared to LFD animals in the PC region (p ≤ 0.05), but it was reduced to near-control (LFD) level in both LFDE and HFDE animals as is evident from Western blotting (Fig. 3b, c) and real-time PCR results (Fig. 3d). No change was seen in the mRNA expression of IL-1β between the LFD and HFD groups, but it showed significant recovery in the LFDE and HFDE groups in the PC region at transcriptional level (Fig. 3d).

Further, we estimated the circulating levels of leptin and insulin in these groups using ELISA-based assays. The level of leptin significantly increased in the HFD group (p ≤ 0.01) as compared to LFD animals and was partially recovered in the HFDE group (Fig. 4a, left panel). The level of leptin in the LFDE group was not different from the LFD group. Along with the increase in circulating level of leptin, the expression of leptin receptor OB-Rb showed decrease in HFD animals in both the hippocampus and PC regions of the brain as shown by both Western blot (Fig. 4b) and real-time PCR analysis (Fig. 4c, d), whereas, the expression of OB-Rb in HFDE and LFDE animals was similar to the LFD group. The circulating level of insulin also showed increase in the HFD animals (p = 0.06), which was recovered to near-control (LFD) level in HFDE animals (Fig. 4a, right panel).

Since SOCS1 and SOCS3 have been implicated in the development of leptin and insulin resistance, we further evaluated the mRNA expression level of these genes. Real-time PCR analysis showed significant upregulation (p ≤ 0.05) of both SOCS1 and SOCS3 in the hippocampus and PC regions of the brain, which was alleviated in HFDE animals (Fig. 4c, d). Leptin is known to activate JAK2-STAT3 signaling pathway, which regulates the expression of SOCS3. So, we further elucidated the differences in mRNA expression of JAK2 and STAT3. The expression of both JAK2 and STAT3 showed significant upregulation (p ≤ 0.05) in the hippocampus region of HFD animals, which was ameliorated by ASH in the HFDE group (Fig. 4c). However, the PC region did not show any change in the expression of JAK2 and STAT3 (Fig. 4d). We further studied the mRNA expression level of insulin receptor substrates IRS1 and IRS2. Consistent with the high level of circulating insulin in the serum, the mRNA levels of IRS1 and IRS2 showed downregulation in HFD animals in both the hippocampus and PC regions, which was restored to the normal level with ASH supplementation in HFDE animals (Fig. 4c, d).

We further extended the study to elucidate the effect of ASH on NF-κB pathway by both Western blot and quantitative real-time PCR (Fig. 5). The phosphorylated form of IKKα/β showed slight upregulation in the hippocampus region of the HFD group as compared to the LFD group, which was partially reduced with ASH supplementation in the HFDE group (Fig. 5a, c). Further, IKKα, which is known to be activated during CNS insult, showed significant (p ≤ 0.05) increase in the hippocampus region of the HFD group animals which was ameliorated in ASH-fed HFDE animals (Fig. 5a, c). IKβα, which is responsible for transcriptional activation of NF-κB, showed upregulation in the hippocampus region of HFD animals in both protein (Fig. 5a, c) and mRNA expression (Fig. 5e), which was suppressed in ASH-fed HFDE animals. However, the PC region of HFD animals showed upregulation only in mRNA expression (Fig. 5e), which was ameliorated in HFDE animals. NF-κB did not show any significant change in hippocampus region among the four groups (Fig. 5a, c, and e). However, its mRNA expression was significantly (p ≤ 0.05) downregulated in the PC region of the LFDE and HFDE groups (Fig. 5e). TLR4, which activates NF-κB pathway via MyD88 protein, showed significant (p ≤ 0.05) upregulation in the hippocampus region of HFD animals (Fig. 5e), but its expression was alleviated in HFDE animals. TLR4 also showed significant (p ≤ 0.05) downregulation in the PC region of HFDE animals (Fig. 5e). MyD88 showed slight upregulation in the HFD animals in the PC region only (Fig. 5e). LFDE animals also showed slight upregulation in the expression of MyD88 in both the hippocampus and PC regions (Fig. 5e) of the brain, but the change was not statistically significant.

Since NF-κB is also known to be instrumental in mediating cell survival pathway, we further studied the expression of apoptotic marker proteins. AP-1 showed significant upregulation in HFD animals as compared to the LFD group in both the brain regions at translational level (Fig. 5b, d), but its expression showed near-control level in ASH-supplemented HFDE animals. However, no change in mRNA expression of AP-1 was observed in HFD animals in either of the brain regions at transcriptional level (Fig. 5e), but it was reduced to 0.6-fold in the hippocampus region in the HFDE group. The PC region did not show any change in mRNA expression of AP-1 (Fig. 5e). Further, Bcl-xL, which is an anti-apoptotic protein, showed downregulation in the HFD animals as compared to LFD animals in both Western blot (Fig. 5b, d) and real-time PCR analysis (Fig. 5e), but the expression of Bcl-xL increased significantly in both the brain regions in LFDE and HFDE animals after supplementation with ASH (Fig. 5b, d, and e). BAD, the pro-apoptotic marker, did not show any change in either of the brain regions (Fig. 5e) in the HFD group, but it was reduced to 0.6-fold in the hippocampus region in the HFDE group. The PC region showed slight upregulation in mRNA expression of BAD in the HFDE group, but it was not statistically significant (Fig. 5e).

As the hypothalamus is most affected by obesity-associated neuroinflammation, so we also evaluated the expression of some inflammatory markers in this region. The expression of Iba1 showed significant increase by 24.9% in the HFD fed group as compared to the LFD group (Fig. 6a, b), while it was significantly reduced to 47.7% with ASH supplementation in HFDE group animals. The increase in expression of Iba1 in the HFD group was also complemented by significant upregulation in the expression of pro-inflammatory cytokines TNFα and IL-1β (p ≤ 0.05) (Fig. 6a, b). However, ASH supplementation effectively suppressed the HFD-induced expression of these cytokines as evident from their near-control expression in the HFDE group.

Further, mRNA expression of iNOS significantly increased by 12.2-fold in the HFD fed group (p ≤ 0.05) as compared to LFD fed rats (Fig. 6c), but it was notably reduced to near-control level with ASH supplementation in the HFDE group. The mRNA levels of other inflammatory markers, such as COX2, TNFα, and IL-1β also showed upregulation in the hypothalamus region of HFD fed animals (p ≤ 0.05) (Fig. 6c). The transcriptional expression of COX2 was suppressed to some extent only in the HFDE group, but TNFα and IL-1β showed near-control levels with HFD+ASH regimen (Fig. 6c). The mRNA expressions of iNOS, COX2, and IL-1β in the LFDE group were not different from the LFD group, but TNFα showed 78% reduction in the LFDE group animals as compared to the LFD group.

Since hypothalamus is the main site for leptin action, we evaluated the effect of HFD on leptin signaling pathway in this region. The expression of OB-Rb, the long form of leptin receptor, showed reduced expression in the HFD fed group as compared to the LFD fed group at both transcriptional (Fig. 7c) and translational (Fig. 7a, b) levels. This also corresponded with the higher circulating level of leptin in this group. However, the expression of OB-Rb was restored to near-control level with ASH supplementation in the HFDE group animals. Since leptin is known to activate JAK2-STAT3-SOCS3 pathway [27], we further elucidated the mRNA expression level of these markers in the hypothalamus region. All the three markers, viz., JAK2, STAT3, and SOCS3, showed significant upregulation in the HFD-fed animals (p ≤ 0.05) as compared to the LFD group (Fig. 7c). However, ASH supplementation effectively suppressed the stress-induced expression of these markers. The LFDE group also showed significant reduction in expression of STAT3 and SOCS3 as compared to the LFD group, but expression of JAK2 was enhanced significantly (Fig. 7c).

Considering the significant inflammatory changes observed in the hypothalamus region of the HFD group animals, we further analyzed the expression level of NF-κB pathway marker proteins. The upstream markers in NF-κB signaling pathway, IKKα and IKβα, showed significant upregulation by 50 and 130%, respectively, in HFD group animals as compared to the LFD group (Fig. 8a, b). The expression of these marker proteins was attenuated to some extent in the HFDE group. NF-κB also showed upregulated expression in the HFD fed group at both transcriptional (Fig. 8c) and translational (Fig. 8a, b) levels. However, it was only reduced to some extent at the translational level (Fig. 8a, b), but mRNA level of NF-κB showed significant reduction in the HFDE group (Fig. 8c). The expression of NF-κB in the LFDE group was similar to the LFD group. Further, the mRNA level of Bcl-xL, the anti-apoptotic marker protein, showed 24% reduction in the HFD-fed group as compared to the LFD group (Fig. 8c), but it was restored to near-control level in the HFDE group. The LFDE group animals also showed higher expression of Bcl-xL as compared to the LFD group animals.