Glucoraphanin Ameliorates Obesity and Insulin Resistance Through Adipose Tissue Browning and Reduction of Metabolic Endotoxemia in Mice

Low-grade sustained inflammation, triggered by chronically high levels of proinflammatory cytokines and gut microbiota–derived circulatory lipopolysaccharide (LPS), links obesity with comorbidities such as insulin resistance and nonalcoholic fatty liver disease (NAFLD) (1,2). Although a number of pharmacological treatments for obesity and NAFLD have been tested, few drugs are clinically available owing to the lack of long-term efficacy and safety concerns (3,4). Thus, a novel therapeutic approach that would improve energy metabolism and reduce chronic inflammation in obesity is sorely needed.

Nuclear factor (erythroid-derived 2)–like 2 (Nrf2), a basic leucine zipper transcription factor, is widely expressed in human and mouse tissues and serves as a defense response against extrinsic and intrinsic stressors (5). Upon exposure to electrophilic and oxidative stress, Nrf2 detaches from its repressor Kelch-like ECH-associated protein 1 nuclear factor (Keap1), and is translocated from the cytoplasm into the nucleus. This translocation leads to the transcriptional activation of genes encoding phase 2 detoxifying and antioxidant enzymes (6). In addition to the ubiquitous induction of cytoprotective genes, Nrf2 regulates a large number of genes involved in glucose and lipid metabolism. In the liver, the constitutive activation of Nrf2 through Keap1 knockdown represses the expression of genes involved in gluconeogenesis (7) and lipogenesis (8), thereby alleviating obesity, diabetes, and hepatic steatosis. Accordingly, synthetic Nrf2 inducers, such as synthetic triterpenoid 2-cyano-3,12-dioxoolean-1,9-dien-28-oic acid (CDDO)-imidazolide (9), CDDO-methyl ester (known as bardoxolone methyl) (10), and dithiolethione analog oltipraz (11), have been shown to ameliorate high-fat diet (HFD)–induced obesity and diabetes. These synthetic Nrf2 inducers also decrease liver and adipose tissue lipogenesis and enhance glucose uptake in skeletal muscles. However, the mechanisms by which Nrf2 enhances energy metabolism in response to an HFD remain largely unknown. Although enhanced Nrf2 signaling has shown promising results in several animal studies, the synthetic Nrf2 inducers have caused adverse cardiac events and gastrointestinal toxicities in clinical trials (12,13). These observations prompted us to explore a safer Nrf2 inducer for the treatment of obesity, insulin resistance, and NAFLD.

Sulforaphane, an isothiocyanate derived from cruciferous vegetables, is one of the most potent naturally occurring Nrf2 inducers; this compound exhibits anticancer activity in cancer cell lines and in carcinogen-induced rodent models (14). Among the cruciferous vegetables, broccoli sprouts are the best source of glucoraphanin, a stable glucosinolate precursor of sulforaphane (15). In both rodents and humans, glucoraphanin is hydrolyzed by gut microbiota-derived myrosinase into bioactive sulforaphane before intestinal absorption (16). A recent clinical study demonstrated the safety of orally administered glucoraphanin (17). In the current study, we examined the dietary glucoraphanin-mediated modulation of systemic energy balance and the mitigation of chronic inflammation, insulin resistance, and NAFLD in diet-induced obese mice.

The sulforaphane precursor glucoraphanin was prepared as previously described (17) with minor modifications. Briefly, 1 day after germination from broccoli seeds (Caudill Seed Company, Louisville, KY), sprouts were boiled in water for 30 min. The water extract was mixed with dextrinized cornstarch and subsequently spray dried to yield an extract powder containing 135 mg of glucoraphanin per gram (0.31 mmol/g) (18). The total glucoraphanin titer in the resulting powder was determined by high-performance liquid chromatography as previously reported (19).

Male C57BL/6JSlc mice were purchased from Japan SLC (Hamamatsu, Japan) at 7 weeks of age. The Nrf2 knockout (Nrf2^−/−) mouse strain (RBRC01390; C57BL/6J background) was provided by RIKEN BioResource Center (Tsukuba, Japan) (6). After 1 week of acclimation, mice were fed normal chow (NC) (containing 2.2% dextrinized cornstarch, 10% kcal from fat, #D12450B; Research Diets, New Brunswick, NJ), NC containing 0.3% glucoraphanin (NC-GR) (containing 2.2% extract powder), an HFD (containing 2.2% dextrinized cornstarch, 60% kcal from fat, #D12492; Research Diets), or an HFD containing 0.3% glucoraphanin (HFD-GR) (containing 2.2% extract powder) for 14 weeks. Both the NC and the HFD containing cornstarch or glucoraphanin were prepared by Research Diets. All mice studied were maintained on a 12-h light/dark cycle at 24–26°C with free access to water and food. All animal procedures were performed in accordance with the Guidelines for the Care and Use of Laboratory Animals at Kanazawa University, Japan.

After 3 weeks of feeding, mice were individually housed in an indirect calorimeter chamber at 24–26°C (Oxymax; Columbus Instruments, Columbus, OH). Calorimetry, daily body weight, and daily food intake data were acquired during a 3-day acclimation period followed by a 2-day experimental period. VO₂ and VCO₂ were measured in each chamber every 20 min. The respiratory exchange ratio (RER = VCO₂/VO₂) was calculated with use of Oxymax software. Energy expenditure was calculated as VO₂ × (3.815 + [1.232 × RER]) and normalized to the body mass of each mouse.

Metabolic parameters, body fat composition, insulin sensitivity, and glucose tolerance were assessed as previously described (20). Plasma LPS levels were analyzed with a Limulus Amebocyte Lysate assay kit (QCL-1000; Lonza, Allendale, NJ). Plasma LPS-binding protein (LBP) levels were determined with an ELISA kit (Enzo Life Sciences, Farmingdale, NY). Immunoblotting was performed with primary antibodies (Supplementary Table 1) as previously described (20). mRNA expression levels were determined by quantitative real-time PCR that used SYBR Green with the primers (Supplementary Table 2) as previously described (20).

Stromal vascular fractions from inguinal white adipose tissue (WAT) of 7-week-old wild-type and Nrf2^−/− mice were prepared as previously reported (21). At confluence, stromal vascular fraction cells were induced for 2 days with differentiation medium containing DMEM/F-12 supplemented with 10% FBS, 20 nmol/L insulin, 1 nmol/L T3, 5 μmol/L dexamethasone, 500 μmol/L isobutylmethylxanthine, 125 μmol/L indomethacin, and 0.5 μmol/L rosiglitazone (all from Sigma-Aldrich, St. Louis, MO). Induced cells were subsequently cultured in maintenance medium (DMEM/F-12 containing 10% FBS, 20 nmol/L insulin, and 1 nmol/L T3) for 5 days and treated with DMSO or sulforaphane (Toronto Research Chemicals, Toronto, Ontario, Canada) at the indicated concentrations for 48 h.

Cells from the liver and epididymal WAT were prepared as previously described (22). Isolated cells were incubated with Fc-Block (BD Bioscience, San Jose, CA) and subsequently incubated with fluorochrome-conjugated antibodies (Supplementary Table 1). Flow cytometry was performed on a FACSAria II (BD Bioscience), and the data were analyzed with FlowJo software (Tree Star, Ashland, OR).

Metagenomic DNA was extracted from mouse cecal content with a QIAamp DNA Stool Mini Kit (QIAGEN, Hilden, Germany). The V1–V2 region of the 16S rRNA gene was amplified by using primer sets as previously reported (23). Mixed samples were prepared by pooling approximately equal amounts of PCR amplicons from each sample and subjected to a GS Junior System (Roche Diagnostics, Basel, Switzerland) for subsequent 454 sequencing. Preprocessing and taxonomic assignment of sequencing reads were conducted as described previously (23) and separated by unique bar codes. The 16S rRNA sequence database was constructed by retrieving 16S sequences of bacterial isolates (1,200–2,384 bases in length) from the Ribosomal Database Project Release 10.27. We used 4,000 filter-passed reads of 16S sequences for the operational taxonomic unit (OTU) analysis of each sample. Clustering of 16S sequence reads with identity scores >96% into OTUs was performed by using the UCLUST algorithm (www.drive5.com). Representative sequences with identity scores >96% for each OTU were assigned to bacterial species by using the BLAST algorithm. Principal component analysis with EZR software (www.jichi.ac.jp/saitama-sct/SaitamaHP.files/statmedEN.html) was applied for assessment of alterations of cecal bacterial phylum associated with the diets.

Data were expressed as mean ± SEM. P < 0.05 was considered statistically significant. Statistical differences between groups were determined by a two-tailed Student t test. An overall difference among more than two groups was determined by one-way ANOVA. If one-way ANOVAs were significant, differences between individual groups were estimated by Bonferroni post hoc test. All calculations were performed with SPSS version 19.0 statistical software (IBM Corporation, Armonk, NY).

To investigate the effects of glucoraphanin on systemic energy balance, we examined the body weight of wild-type mice fed NC or an HFD supplemented with glucoraphanin or vehicle (i.e., cornstarch only). Glucoraphanin reduced weight gain only in HFD-fed mice without affecting food intake (Fig. 1A and Supplementary Fig. 1A). This reduction was not accompanied by evidence of gross toxicity. We determined the plasma concentration of sulforaphane in NC-GR and HFD-GR mice, but not in NC or HFD mice, indicating that glucoraphanin was absorbed as a sulforaphane after food consumption (Supplementary Fig. 1B). The reduction of weight gain in HFD-GR mice was largely attributed to decreased fat mass, not to lean mass (Fig. 1B). To assess energy expenditure, we placed the mice in indirect calorimetry cages after 3 weeks of feeding before an evident change in the body mass of HFD-fed mice was observed (29.9 ± 0.5 vs. 28.5 ± 0.5 g in HFD vs. HFD-GR, respectively). HFD-GR mice exhibited consistently higher VO₂ and VCO₂ than HFD controls (Fig. 1C and D), leading to increased energy expenditure (Fig. 1E); however, they displayed a similar RER (Fig. 1F), suggesting that glucoraphanin supplementation enhanced sugar and fat use under HFD conditions. In NC-GR mice, these parameters of energy balance were not affected (Fig. 1B–F). Consistent with increased energy expenditure, glucoraphanin increased the core body temperature of HFD-fed mice by ∼0.5°C (Fig. 1G).

After 14 weeks of feeding, glucoraphanin supplementation did not affect plasma triglyceride, total cholesterol, and free fatty acid (FFA) levels in either NC- or HFD-fed mice (Table 1). In NC-GR mice, blood glucose levels were not altered by glucoraphanin, but HFD-GR mice exhibited significantly lower fasted blood glucose compared with vehicle-treated controls (Table 1). Additionally, glucoraphanin significantly decreased plasma insulin concentrations in HFD-fed mice under both fasted and fed conditions, resulting in lower HOMA of insulin resistance (HOMA-IR) (Table 1). During the insulin tolerance test (ITT), glucoraphanin significantly enhanced the reduction in blood glucose levels in HFD-fed mice but not in NC-fed mice compared with the vehicle-treated controls (Fig. 2A). Glucoraphanin improved glucose tolerance in HFD-fed mice during the glucose tolerance test (GTT), but had no effect in NC-fed mice (Fig. 2B). Insulin secretion during the GTT was not affected by glucoraphanin (data not shown). In line with increased insulin sensitivity, insulin-stimulated Akt phosphorylation on Ser⁴⁷³ was enhanced by glucoraphanin in the liver, muscle, and epididymal WAT of HFD-fed mice (Fig. 2C).

Although the Keap1-Nrf2 pathway is a well-known target of sulforaphane, this isothiocyanate has also been reported to modulate different biological pathways independent of the Keap1-Nrf2 pathway (24,25). To determine whether the antiobesity and insulin-sensitizing effects of glucoraphanin are mediated through Nrf2, the effects of glucoraphanin on energy balance and glucose metabolism were assessed in NC- and HFD-fed Nrf2^−/− mice. Although food intake and plasma concentration of sulforaphane in NC-GR or HFD-GR Nrf2^−/− mice were comparable with that detected in wild-type NC-GR or HFD-GR mice (Supplementary Fig. 1C and D), the effects of glucoraphanin on weight gain after HFD feeding (Fig. 3A), VO₂ (Fig. 3B), VCO₂ (Fig. 3C), energy expenditure (Fig. 3D), RER (Fig. 3E), rectal temperature (Fig. 3F), insulin sensitivity (Fig. 3G), and glucose tolerance (Fig. 3H) were abolished by the Nrf2 deficiency. These data are consistent with comparable plasma metabolic parameters between HFD-GR and HFD mice. These metabolic parameters include lipids, blood glucose, insulin, HOMA-IR, and liver enzymes such as alanine transaminase (ALT) and aspartate transaminase (AST) (Supplementary Table 3).

The increased energy expenditure and body temperature of HFD-GR mice suggest an increase in adaptive thermogenesis. However, glucoraphanin supplementation had little effect on the size and number of lipid droplets in the intrascapular brown adipose tissue (BAT) of HFD-fed wild-type mice (Supplementary Fig. 2A). In addition, the mRNA expression of Ucps, PGC-1α, and deiodinase 2 in BAT and of Ucps in skeletal muscle was not altered by glucoraphanin supplementation in NC- and HFD-fed wild-type mice (Supplementary Fig. 2B and C). In BAT, HFD increased Ucp1 protein expression, but glucoraphanin did not alter the expression in wild-type or Nrf2^−/− mice (Fig. 4A). Brown-like adipocytes expressing Ucp1, also known as beige cells, exist in various WAT depots and can contribute to thermogenesis (26). Compared with NC, HFD significantly decreased Ucp1 protein levels in epididymal and inguinal WAT of both wild-type and Nrf2^−/− mice (Fig. 4A). Glucoraphanin supplementation restored HFD-induced reduction in Ucp1 protein levels in epididymal and inguinal WAT of wild-type mice but not in Nrf2^−/− mice. To examine whether the effects of glucoraphanin were fat cell autonomous and Nrf2 mediated, we tested the effects of sulforaphane, an active metabolite of glucoraphanin, on the expression of brown fat–selective genes in primary beige adipocytes obtained from inguinal WAT of wild-type and Nrf2^−/− mice. In wild-type beige adipocytes, treatment with sulforaphane induced the Nrf2 target gene NAD(P)H:quinone oxidoreductase 1 (Nqo1) (Fig. 4B) and antioxidant genes (Supplementary Fig. 3A). Concurrently, sulforaphane significantly increased the mRNA expression of brown fat–selective genes, including Ucp1, Prdm16, Cidea, and Elovl3 (Fig. 4B). In contrast, in Nrf2-deficient beige adipocytes, sulforaphane failed to activate Nrf2, as judged by unaltered mRNA expression of the target genes, and to promote the expression of brown fat–selective genes (Supplementary Fig. 3B and Fig. 4C). Of note, Nrf2-deficient beige adipocytes exhibited fewer differentiation levels associated with attenuated lipid accumulation (Supplementary Fig. 3C) and lower mRNA expression of fatty acid binding protein 4 (Supplementary Fig. 3C) and brown fat–selective genes compared with wild-type beige adipocytes (Fig. 4C).

The HFD caused hepatic steatosis and inflammation, eventually leading to steatohepatitis. As shown in Fig. 5A, the increase in liver weight caused by the 14-week HFD was alleviated by glucoraphanin supplementation. Glucoraphanin also attenuated HFD-induced hepatic steatosis (Fig. 5B). Additionally, compared with the HFD group, the lower levels of plasma ALT, plasma AST, liver triglycerides, and liver FFAs in the HFD-GR mice indicated that glucoraphanin alleviated HFD-induced liver damage (Fig. 5C and D). The reduction in hepatic steatosis was accompanied by the decreased expression of the following lipogenic genes: sterol regulatory element binding transcription factor 1c (Srebf1), fatty acid synthase (Fasn), and peroxisome proliferator–activated receptor γ (Pparγ) (Fig. 5E). Additionally, hepatic levels of malondialdehyde, a marker of lipid peroxidation, were increased by the HFD. Glucoraphanin attenuated lipid peroxidation (Fig. 5F) and decreased gene expression of the NADPH oxidase subunits gp91^phox, p22^phox, p47^phox, and p67^phox (Fig. 5E). The HFD led to a compensatory increase in the expression of genes involved in fatty acid β-oxidation (Ppara and Cpt1a) and antioxidative stress (Cat, Gpx1, and Sod1) in the liver. However, glucoraphanin did not further increase the expression of these genes in the liver of HFD-fed mice (Fig. 5E).

In response to the HFD, liver-resident macrophages (Kupffer cells) increase the production of proinflammatory cytokines that promote insulin resistance and NAFLD in mice (27). In particular, chemokine (C-C motif) ligand 2 (Ccl2) promotes the recruitment of chemokine (C-C motif) receptor 2 (Ccr2)–positive monocytic lineages of myeloid cells into the liver (28). These recruited cells produce a large amount of proinflammatory mediators and activate a lipogenic program (28). Here, we found a prominent induction of tumor necrosis factor-α (Tnf-α), Ccl2, and Ccr2 in the liver of HFD-fed mice, which was markedly reduced in glucoraphanin-treated mice (Fig. 6A). Glucoraphanin significantly suppressed HFD-induced inflammatory pathways, such as c-Jun N-terminal kinase (JNK) and extracellular signal–regulated kinase (Erk) (Fig. 6B). Glucoraphanin tended to decrease levels of p-NF-κB p65 (Ser⁵³⁶) in HFD-fed mice, although this decrease was not statistically significant (Fig. 6B). Of note, in the liver of Nrf2^−/− mice, glucoraphanin failed to suppress HFD-induced inflammatory signal pathways (Supplementary Fig. 4). In addition, glucoraphanin significantly decreased the HFD-induced hepatic expression of macrophage markers, including F4/80, Cd11b, and Cd68 (Fig. 6C). Tissue macrophages are phenotypically heterogeneous and have been characterized according to their activation/polarization state as M1-like proinflammatory macrophages or M2-like anti-inflammatory macrophages (29). Consistent with the decreased expression of macrophage markers, glucoraphanin prevented macrophage (F4/80⁺CD11b⁺ cell) accumulation in the liver of HFD-fed mice (Fig. 6D). Additionally, glucoraphanin decreased the number of M1-like liver macrophages expressing surface markers (F4/80⁺CD11b⁺CD11c⁺CD206⁻) (Fig. 6E). In contrast, glucoraphanin increased the number of M2-like liver macrophages (F4/80⁺CD11b⁺CD11c⁻CD206⁺), resulting in a predominantly M2-like macrophage population (Fig. 6E). Moreover, glucoraphanin decreased the mRNA expression of Tnf-α and NADPH oxidase in the epididymal WAT of HFD-fed mice (Supplementary Fig. 5A). Although the HFD-induced expression of macrophage markers and macrophage accumulation in epididymal WAT were not altered by glucoraphanin (Supplementary Fig. 5B and C), the number of M1-like macrophages was significantly decreased in the epididymal WAT of HFD-GR mice (Supplementary Fig. 5D).

Gut microbiota–derived LPS induces chronic inflammation that eventually leads to insulin resistance in obesity, termed metabolic endotoxemia (1,2). On the basis of our observation that glucoraphanin alleviates inflammation in the liver and epididymal WAT of HFD-fed mice, we subsequently investigated the effects of glucoraphanin on metabolic endotoxemia and gut microbiota. In accordance with previous studies (1,2), the HFD induced a twofold increase in circulatory LPS levels, which was reduced by glucoraphanin supplementation (Fig. 7A). Furthermore, plasma and hepatic levels of the LPS marker LBP were significantly elevated by the HFD and reduced by glucoraphanin supplementation (Fig. 7B). A principal component analysis distinguished cecal microbial communities based on diet and treatment, revealing that the metagenomes of HFD-fed mice formed a cluster distinct from that formed by NC-fed mice (Fig. 7C). However, samples from HFD-GR mice formed a cluster that was indistinguishable from that of NC or NC-GR mice (Fig. 7C). Of note, consistent with previous reports (30,31), further analysis at the phylum level demonstrated that the proportion of gram-negative Proteobacteria was significantly elevated in the gut microbiomes of HFD-fed mice, which was suppressed by glucoraphanin supplementation (Fig. 7D and E). The increase in the relative abundance of Proteobacteria in HFD-fed mice is mostly explained by an increase in the relative abundance of bacteria from the family Desulfovibrionaceae (Fig. 7F), key producers of endotoxins in animal models of obesity (30). In fact, the relative abundance of Desulfovibrionaceae was positively correlated with plasma LPS levels (Fig. 7G). Furthermore, plasma LPS levels were significantly and positively correlated with the hepatic mRNA levels of Tnf-α, gp91^phox, and F4/80 (Fig. 7H). Similarly, the liver expression of other marker genes was significantly and positively correlated with plasma LPS levels and with one another (Supplementary Table 4).

In the current study, we demonstrated that glucoraphanin, a stable precursor of the Nrf2 inducer sulforaphane, mitigated HFD-induced weight gain, insulin resistance, hepatic steatosis, oxidative stress, and chronic inflammation in mice. The weight-reducing and insulin-sensitizing effects of glucoraphanin were abolished in Nrf2^−/− mice. Additionally, glucoraphanin lowered plasma LPS levels in HFD-fed mice and decreased the relative abundance of Desulfovibrionaceae. At the molecular level, glucoraphanin increased Ucp1 protein expression in WAT depots while suppressing the hepatic mRNA expression of genes involved in lipogenesis, NADPH oxidase, and inflammatory cytokines. The data suggest that in diet-induced obese mice, glucoraphanin restores energy expenditure and limits gut-derived metabolic endotoxemia, thereby preventing hepatic steatosis, insulin resistance, and chronic inflammation.

Consistent with previous reports demonstrating the antiobesity effects of synthetic Nrf2 inducers (9–11), we show that the oral administration of glucoraphanin mitigates HFD-induced weight gain (Fig. 1A). The dose of glucoraphanin used in the current study (∼12 μmol/mouse/day) is similar to that used in other experiments that investigated its antitumor effects in mice (14,32,33). Here, we show that the effect of glucoraphanin on whole-body energy expenditure and the protein expression of Ucp1 in WAT were abolished in Nrf2^−/− mice (Figs. 3D and 4A). A study that used adipocyte-specific PRDM16-deficient mice indicated that adaptive thermogenesis in beige fat also contributes to systemic energy expenditure (26). The mutant mice in the aforementioned study, which exhibited markedly reduced Ucp1 mRNA expression in inguinal WAT and minimal effects on BAT, developed obesity and insulin resistance in response to an HFD. Thus, we believe that the increased energy expenditure in HFD-GR mice at least in part stems from an increase in beige fat, even though the expression of Ucps in BAT and skeletal muscle is not altered. Further analysis in Ucp1 knockout mice will elucidate the relative contribution of beige fat to the Nrf2-mediated metabolic effects elicited by glucoraphanin.

Several studies have indicated that Nrf2^−/− mice are partially protected from HFD-induced obesity and are associated with milder insulin resistance compared with wild-type counterparts (9,34,35). Recently, Schneider et al. (35) demonstrated mitigation of HFD-induced obesity in Nrf2^−/− mice, which was 25% less body weight than that of wild-type mice after 6 weeks of feeding. They also found that HFD-fed Nrf2^−/− mice exhibited a 20–30% increase in energy expenditure associated with an approximately threefold upregulation of Ucp1 protein expression in abdominal WAT. In the current study, Nrf2^−/− mice gained less weight after 6 weeks of HFD feeding than the HFD-fed wild-type mice (35.9 ± 0.9 vs. 39.0 ± 0.8 g, Nrf2^−/− wild type, respectively; P < 0.05) (Figs. 1A and 3A). The lower body mass of Nrf2^−/− mice raises the possibility that the antiobesity effect of glucoraphanin was completely phenocopied by Nrf2 gene deficiency. However, several observations suggest that Nrf2^−/− mice only partially phenocopy the effect of glucoraphanin on weight gain reduction. First, in the current study, the weight difference between Nrf2^−/− and wild-type mice was only 8%, which is much less than that in Schneider et al. Second, in Nrf2^−/− mice, HFD induced significant weight gain (Fig. 3A), glucose intolerance (Fig. 3H), and insulin resistance compared with NC, as judged by increased HOMA-IR (Supplementary Table 3). Third, metabolic rate and energy expenditure of Nrf2^−/− mice were comparable with those in wild-type mice (Figs. 1C–E and 3B–D). Finally, Ucp1 protein levels in both epididymal WAT and inguinal WAT of HFD-fed Nrf2^−/− mice were lower than in HFD-GR wild-type mice (Fig. 4A). Taken together, these findings suggest that Nrf2 gene deficiency is not sufficient to block HFD-induced obesity by increasing energy expenditure and Ucp1 expression in WAT depots and to mask the effect of glucoraphanin. However, we cannot fully exclude the possibility that the effects of glucoraphanin are mediated by Nrf2-independent mechanisms. Possible reasons for the discordance in metabolic phenotypes of Nrf2^−/− between Schneider et al. and the current study may be due to differences in knockout mouse lines and experimental conditions (e.g., age of mice at beginning of HFD feeding, composition of HFD, temperature in the metabolic chamber).

The current in vitro study of primary beige adipocytes revealed that sulforaphane promotes the expression of brown fat–selective genes (Fig. 4B). Of note, the concentration of sulforaphane used in cell culture (0.2–5 μmol/L) is comparable with that detected in mice fed NC-GR and HFD-GR (Supplementary Fig. 1B). Moreover, we determined that Nrf2 acts as a positive regulator of beige adipocyte differentiation (Fig. 4C and Supplementary Fig. 3C). The fewer differentiation levels in Nrf2-deficient beige adipocytes agree with previous reports demonstrating that Nrf2 induces white adipocyte differentiation through increasing the gene expression of Pparγ (34) and Cebpβ (36), common transcription factors that regulate the differentiation of brown, beige, and white adipocytes. Furthermore, Nrf2 has been reported to bind NF-E2–binding sites in the 5′ flanking region of human and rodent Ucp1 genes (37). However, we cannot exclude the possibilities that glucoraphanin affects sympathetic nervous activity or that hormonal factors regulate fat browning (38). In addition, mitochondrial reactive oxidative species facilitate Ucp1-dependent respiration in BAT and whole-body energy expenditure by promoting the sulfenylation of a specific cysteine residue (Cys253) in Ucp1 (39). The molecular mechanism by which Nrf2 regulates the expression and thermogenic activity of Ucp1 in beige adipocytes requires further investigation.

Glucoraphanin supplementation improved the systemic glucose tolerance and insulin sensitivity of HFD-fed mice. Although the molecular mechanism by which synthetic Nrf2 inducers enhance glucose uptake is unclear, AMPK activation may mediate this enhancement in mouse skeletal muscle and adipose tissue (9–11). The phosphorylation levels of AMPK (Thr¹⁷²) and acetyl-CoA carboxylase (Ser⁷⁹) in peripheral insulin target tissues were comparable between NC-GR and HFD-GR mice and vehicle-treated controls (Supplementary Fig. 6). These data suggest that AMPK activation is not necessary for glucoraphanin to exert its insulin-sensitizing effect on HFD-fed mice. Additional studies that use the hyperinsulinemic-euglycemic clamp technique are needed to determine which tissues contribute to the insulin-sensitizing effects of glucoraphanin.

The beneficial effects of glucoraphanin on hepatic lipid metabolism were not accompanied by AMPK activation or the increased expression of fatty acid β-oxidation genes (Fig. 5E). Instead, glucoraphanin mitigated HFD-induced oxidative stress and inflammation in the liver. In obesity, hepatic inflammation mediated by macrophage/monocyte-derived proinflammatory cytokines promotes lipogenesis through the inhibition of insulin signaling and SREBP activation (40,41). In fact, the depletion of Kupffer cells by clodronate liposomes ameliorates hepatic steatosis and insulin sensitivity in HFD-fed mice (27). Furthermore, Ccl2- or Ccr2-deficient mice are protected from diet-induced hepatic steatosis, even though they still become obese (42,43). Moreover, the specific ablation of M1-like macrophages restores insulin sensitivity in diet-induced obese mice (44), whereas the deletion of Pparδ, which promotes M2 activation, predisposes lean mice to developing insulin resistance (45). Therefore, decreased hepatic macrophage accumulation and M2-dominant polarization of hepatic and adipose macrophages account, at least in part, for the protection from hepatic steatosis and insulin resistance in HFD-GR mice.

One of the most important findings of this study is that glucoraphanin decreases the relative abundance of gram-negative Proteobacteria, particularly family Desulfovibrionaceae, while reducing circulatory LPS levels (Fig. 7). Studies have demonstrated a significant increase in Desulfovibrionaceae, potential endotoxin producers, in the gut microbiomes of both HFD-induced obese mice and obese human subjects compared with lean individuals (30,31,46). We cannot exclude the possibility that other microbiota-derived products, such as bile acids and short-chain fatty acids, also mediate the metabolic action of glucoraphanin. Whether the interaction between sulforaphane and gut microbiota is affected directly or indirectly by altered host physiology remains to be determined. However, several studies have suggested that sulforaphane can alter the gut microbiota directly because isothiocyanates (including sulforaphane) have been shown to exhibit antibacterial activity against Proteobacteria (47,48). This activity may proceed through redox disruption and enzyme denaturation reactions involving the isothiocyanate reactivity group, -N = C = S, the thiol group (-SH) of glutathione, and proteobacterial proteins (47,48). Additionally, sulforaphane exhibits antibacterial activity against Helicobacter pylori, a member of the phylum Proteobacteria (49). We are unaware of previous reports demonstrating that isothiocyanates inhibit the proliferation of Desulfovibrionaceae. The mechanistic underpinnings of this antibacterial activity require elucidation.

In conclusion, the results of this study indicate that glucoraphanin may be effective in preventing obesity and related metabolic disorders such as NAFLD and type 2 diabetes. Another clinical study demonstrated that supplementation with a dietary dose of glucoraphanin (69 μmol/day) for 2 months significantly decreased plasma liver enzymes, ALT, and AST, although body mass did not change (18). Long-term treatment with a higher dose of glucoraphanin (800 μmol/day), which can be safely administered without harmful adverse effects (17), may be required to achieve an antiobesity effect in humans.

Acknowledgments. The authors thank M. Nakayama and K. Hara (Kanazawa University) for technical assistance. The authors also thank Editage (www.editage.jp) for English-language editing.

Funding. This work was supported by Japan Society for the Promotion of Science KAKENHI grant numbers 15K00813 (to N.N.), 15K12698 (to T.O.), and 16H03035 (to T.O.).

Duality of Interest. No potential conflicts of interest relevant to this article were reported.

Author Contributions. N.N. collected data and wrote the manuscript. L.X., S.Ko., N.F., F.Z., Y.N., and M.N. collected data. Y.U., Y.A., and R.U. collected data and edited the manuscript. C.T., H.S., and S.Ka. contributed to the discussion and reviewed the manuscript. T.O. contributed to the discussion and reviewed and edited the manuscript. T.O. is the guarantor of this work and, as such, had full access to all the data in the study and takes responsibility for the integrity of the data and the accuracy of the data analysis.

Prior Presentation. Parts of this article were presented at the 76th Scientific Sessions of the American Diabetes Association, New Orleans, LA, 10–14 June 2016.