Age-matched male and female C57Bl6/J mice were used in all studies. 13-week old mice were given ad libitum access to a HFD (Research Diets D12451; 45% kcals fat, 20% kcals protein, 35% kcals carbohydrate; 5.8% fiber; 4.7 kcal/g metabolizable energy) and drinking water containing vehicle or 1 mg/mL DHF for 12-weeks. Water preparation for the conventional non-antibiotic treated group was as reported previously (38). For antibiotic (Abx) treatment, 11-week old female mice were given drinking water containing a cocktail of Abx including ampicillin (1 g/L), neomycin (1 g/L), metronidazole (1 g/L) (all from Sigma, St. Louis, MO, United States) and vancomycin (VWR Radnor, PA, United States) (0.5 g/L) in 1% sucrose and 0.17% DMSO in PBS for 2 weeks prior to HFD feeding. Bacterial ablation was confirmed as described below. Following this, mice were randomly assigned to Abx-control or Abx-DHF groups and received Abx throughout the 12-week HFD-feeding.

Body weights and food intake were measured weekly, and water intake was measured twice a week (38). Body composition was determined by NMR (Echo MRI, Columbus, OH, United States), at the start and end of HFD-feeding (38). The breeding and care of the animals were in accordance with the protocols approved by the Animal Care and Use Committee of Rutgers University, New Brunswick, New Jersey. For in vivo procedures, all efforts were made to minimize discomfort and suffering, in accordance with approved animal care protocols. Following 12 weeks of feeding, mice were euthanized by isoflurane overdose, followed by cardiac exsanguination. Tissues were rapidly collected, snap-frozen in liquid nitrogen, and stored at −80°C until further analyses.

Fresh fecal pellets were collected before and after 2 weeks of Abx-treatment and after 1, 6, and 12 weeks of HFD-feeding and stored at −80°C until further analysis. Fecal bacterial DNA was isolated using QIAamp DNA stool mini kit (QIAGEN, Germantown, MD, United States) following manufacturer’s instructions. Total fecal 16S rRNA was quantified by qPCR using forward primer 785F: 5′-GGA TTA GAT ACC CTG GTA and reverse primer 907R: 5′-CCG TCA ATT CCT TTR AGT TT, spanning the V5 region of 16S rRNA gene (40). A 16S rRNA plasmid DNA with known copy number was used to generate a standard curve to quantify bacterial DNA in samples, as previously described (40).

Energy expenditure was measured by indirect calorimetry (Oxymax, Columbus Instruments, Columbus, OH) after 8 weeks of feeding, as previously described (41). Briefly, oxygen consumption (VO₂) and carbon dioxide production (VCO₂) were determined in individually housed animals with ad libitum access to HFD and water; both DHF and Abx were maintained in drinking water of respective groups during the calorimetry studies. Animals were acclimatized to Oxymax chambers for 24 h, and VO₂ and VCO₂ were recorded for eight consecutive 12-h light and 12-h dark cycles. Metabolic variables were recorded every minute, with a room air reference taken following each cycle of measurements.

Oral glucose tolerance test (GTT) was performed after 10 weeks of HFD-feeding. Briefly, mice were fasted for 4 h, followed by oral gavage of 20% dextrose at a dose of 1 g/kg body weight. Blood was collected from the tail at 0, 20, 40, 90, and 120 min for assessment of plasma glucose using an Accu-Chek glucometer (Accu-Chek Performa, Roche Diagnostics, IN, United States).

Plasma free cholesterol was measured using the Wako Free Cholesterol E Microtiter kit, as per manufacturer’s instructions (FUJIFILM Healthcare, Lexington, MA, United States). Hepatic lipids were extracted by a modified Folch method and separated by thin-layer chromatography (TLC), as previously described (42). Briefly, 20 mg liver tissue was homogenized in chloroform:methanol (2:1), followed by phase separation by the addition of acidified saline and centrifugation. The organic phase was collected, dried under a nitrogen stream, reconstituted in chloroform:methanol (2:1), spotted onto Silica Gel 60 chromatography plates, and developed in heptane:isopropyl ether:acetic acid (60:40:3) until the solvent front reached 1 cm from the top of the plate. Plates were dried, dipped in 10% copper sulfate in 10% phosphoric acid, and charred at 110°C for 30 min.

Protein samples were prepared by homogenizing tissues in HEPES lysis buffer (50 mM pH 7.4 HEPES, 150 mM NaCl, 10 mM Na-pyrophosphate, 2 mM EDTA, 1% NP-40, 10% Glycerol) containing 1x protease and phosphatase inhibitor cocktail (Sigma-Aldrich). Equal amounts of protein were separated by SDS-PAGE, transferred on PVDF membranes, and immunoblotted with primary antibodies. Primary antibodies including PGC-1α (Novus Biologicals, Littleton, CO, United States), VDAC (Pierce Biotechnology, Waltham, MA, United States), UCP1, COX-IV (Abcam, Cambridge, United Kingdom), and Total TrkB, P-TrkB, PAMPK, Total AMPK, HSP60, SIRT1, and GAPDH (all from Cell Signaling Technology, Danvers, MA, United States) were used at manufacturer recommended dilutions. Following incubation with HRP- or near-IR dye-conjugated secondary antibodies, membranes were visualized using enhanced chemiluminescence or near-IR imaging on an Azure c600 imaging system (Azure Biosystems, United States) and quantified using ImageStudio Lite.

All data are expressed as mean ± SEM. Statistical analysis are carried out using student’s t- test for two group comparisons or, for multi-group comparisons using one-way ANOVA, followed by post hoc analysis (Bonferroni) in Graph Pad Prism (version 9.3.1 for Windows, GraphPad Software, La Jolla, CA, United States). p-values < 0.05 were considered significant.

We previously reported that mice supplemented with oral DHF for 12 weeks accumulated differential amounts of body weight and fat mass on a high-fat diet (HFD), dependent on sex. Female mice gained significantly less weight and adiposity when supplemented with oral DHF, but male mice were resistant to this metabolic protection afforded by DHF supplementation. We also reported that DHF-induced early and stable remodeling of the gut microbiome in female but not male mice, corresponding with metabolic protection or lack thereof, upon DHF supplementation (38).

To examine a causal role for the gut microbiome in mediating the metabolic effects of DHF in females, we generated an experimental model of gut microbiome ablation by treatment with a cocktail of broad-spectrum antibiotics (Abx) for 2 weeks prior to DHF supplementation and HFD-feeding. Similar protocols have been previously reported to stably and consistently ablate the gut microbial community (43–46). Germ-free mice were not utilized in order to avoid the developmental and immune defects associated with this model (47). Two weeks of Abx-treatment did not result in any body weight changes. Gut microbiome ablation was confirmed by comparing fecal bacterial load at baseline (Week-2) and after 2 weeks of Abx-treatment (Week 0) to a standard curve generated with a 16S rRNA plasmid DNA with known copy number (40). Oral Abx-supplementation decreased the fecal 16S rRNA copy number by >3 log₁₀ fold, with similar reductions maintained throughout the course of the study.

Following this, all conventionally raised i.e., which did not receive Abx-treatment and Abx-treated mice received HFD and were segregated into groups receiving vehicle or DHF, with or without oral Abx, for an additional 12 weeks. As we have previously reported (38), DHF treatment reduced weight gain in female mice that were conventionally raised (Figure 1A). Abx-treatment alone has been demonstrated to attenuate weight gain (48–52), independent of changes in food and water intake and solely dependent on absence of gut microbiome. However, Abx-treatment alone did not attenuate HFD-induced weight gain to the same extent as DHF supplementation (Figure 1A). Furthermore, DHF treatment in Abx-treated animals did not have any further impact on body weights (Figure 1A). Similarly, DHF treatment reduced fat mass in conventional mice but had no effect on fat or lean mass in Abx-treated animals (Figures 1B,C).

Food intake was similar across all groups throughout the treatment period (Figure 1D). A significant reduction in water intake was observed with Abx-treatment (Figure 1E), which may be attributable to taste aversion due to antibiotics. This resulted in a daily DHF intake of 0.26 ± 0.01 mg/d/g BW in conventional females and 0.16 ± 0.01 mg/d/g BW in Abx-treated animals. It is important to note that the lower daily dose was still over 2-3-fold higher than a previously reported low-dose of DHF that conferred protection against weight gain in female mice (37, 51). Thus, the lack of protection from weight gain in the Abx-DHF group is not attributable to reduced DHF intake but rather to the concurrent Abx administration. These data indicate that reductions in HFD-induced weight gain due to oral DHF require an intact gut microbiome.

Gut microbiome ablation has been previously reported to significantly increase cecal weights (52–54). These results were recapitulated in the current study, and DHF did not induce any additional effects on cecal weights (Figure 1F).

Plasma cholesterol was significantly reduced by DHF in the conventional group (Figure 2A). Abx-treatment alone did not reduce plasma cholesterol levels. Importantly, DHF treatment in the Abx-group had no impact on plasma cholesterol, unlike in the conventional group (Figure 2A). Hepatic lipid accumulation was measured by thin-layer chromatography and revealed a significant reduction in levels of diacylglycerols (DAG), triacylglycerols (TAG), and cholesterol esters (CE) in DHF-treated females (Figures 2B,C). However, Abx-DHF animals had no significant reductions in hepatic lipids, relative to Abx-controls. Taken together, these data indicate that reductions in plasma and hepatic lipids by DHF administration require an intact gut microbiome.

To examine the mechanisms of protection from HFD-induced obesity in DHF-fed females, energy expenditure was measured by indirect calorimetry after 8 weeks of HFD-feeding. Significant increases in O₂ consumption and CO₂ respiration were observed in conventionally-raised female DHF-fed mice (Figures 3A,B). Similar increases were not apparent in Abx-treated DHF-fed females (Figures 3C,D). Similarly, regression of energy expenditure against body weight indicated a steeper regression slope in DHF-fed females, relative to control females (Figure 3E; slope: 0.0070 for Control vs. 0.0179 for DHF females) in the conventionally-raised mice, although this difference was not statistically significant. However, this same trend was not observed for DHF-fed females following Abx-treatment (Figure 3F; slope: 0.0011 for Abx-Control vs. 0.0035 for Abx-DHF). We interpret these data to collectively indicate a modest increase in energy expenditure upon DHF supplementation, which is lost in Abx-treated mice. These data are consistent with the protection from weight gain observed in DHF-fed females and lack of protection upon Abx-treatment (Figure 1A) and point to an important role for the gut microbiome in mediating DHF-induced alterations in energy balance.

Consistent with obesity resistance, DHF supplementation of conventional females ameliorated glucose tolerance relative to control females (Figure 3G). Abx-treatment resulted in lower glucose area under the curve (AUC) relative to conventional controls, possibly due to the reduced weight gain in these mice. However, DHF treatment did not improve glucose tolerance in Abx-treated mice, consistent with body weight changes and further implicating a role for the gut microbiome in mediating the improvements in glucose tolerance upon DHF supplementation (Figures 3H,I).

7,8-dihydroxyflavone is an activator of tropomyosin-related kinase receptor B (TrkB) which is activated by phosphorylation at discrete tyrosine residues. We measured activation of TrkB by DHF in both male and female mice (32, 34). A previous study reported that oral DHF increased skeletal TrkB phosphorylation in female mice, without any report on the effects of DHF in males (39). Apart from the brain, peripheral expression of TrkB was reported to be greatest in brown and white adipose tissues, with very low expression reported in liver or skeletal muscle (53). Since brown adipose tissue (BAT) plays a critical role in maintaining energy homeostasis, we focused our current investigations on this tissue. Given the prior report on the role of DHF in activating energy expenditure in skeletal muscle, we also investigated the impact of Abx and DHF in gastrocnemius tissue.

In BAT, we observed that DHF induced TrkB-phosphorylation in female, but not male mice (Figures 4A,B). Further, phosphorylation and activation of AMP-activated protein kinase (AMPK), one of the downstream targets of TrkB, was also elevated in female but not male mice, consistent with sexually-dimorphic TrkB activation in these tissues (Figures 4A,B). In addition, expression of deacetylase, Sirtuin 1 (SIRT1), and its downstream target Peroxisome proliferator activated receptor-gamma coactivator (PGC-1a), were also increased in BAT of DHF-fed female, but not male mice, indicating increased BAT mitochondrial content and metabolism in these animals (Figure 4C,D). Consistent with activation of SIRT1-PGC-1a signaling, DHF supplementation also significantly increased expression of uncoupling protein-1 (UCP1) in BAT of female but not male mice (Figures 4C,D). Given the role of PGC-1α in modulating mitochondrial mass, we measured expression of mitochondrial content markers including COX-IV, VDAC and HSP60. No differences in mitochondrial content were noted (Figures 4C,D).

Similar to BAT, activation of TrkB was significantly increased in gastrocnemius of female, but not male, DHF-fed mice (Figures 5A,B). While AMPK phosphorylation was significantly elevated in BAT of female DHF-fed mice, only a trend for elevation was observed in female gastrocnemius (Figure 5B). Expression of PGC-1α and SIRT1 were also slightly elevated in gastrocnemius of female mice, but again not to the same extent as observed in BAT (Figure 5D). Since a previous study reported a role for DHF in activating UCP1 in skeletal muscle (39), we measured expression of UCP1 in gastrocnemius, despite its low expression in this tissue. We observed a slight but significant increase in UCP1 expression in gastrocnemius of DHF-fed female mice, but not male mice (Figures 5C,D). Again, this 1.5-fold elevation of UCP1 protein expression in gastrocnemius was not as great as the 3-fold elevation observed in BAT, where the role of UCP1 in modulating energy homeostasis is more clearly established. Similar to BAT, no significant changes in mitochondrial content were evident in gastrocnemius of male or female DHF-fed mice (Figures 5C,D).

Taken together, these data indicate clear increases in TrkB activation, UCP1 induction, and activation of the SIRT1-PGC-1α -AMPK axis in BAT and skeletal muscle of DHF-fed female but not male mice, consistent with the sexually-dimorphic effects of DHF on energy homeostasis and consequently on HFD-induced weight gain.

We next examined these markers in Abx-treated females. Interestingly, upon Abx-treatment, no induction of any of these markers was apparent in either BAT (Figures 6A,B) or gastrocnemius (Figures 7A,B) of female mice. Phosphorylation of TrkB and AMPK, and expression of UCP1, SIRT1, and PGC-1a were static across control and DHF-treated mice upon Abx-treatment. These data clearly indicate, for the first time, that an intact gut microbiome is necessary for induction of TrkB as well as for downstream cellular signaling by DHF.