Male A/J, C3H/HeJ, and C3H/HeOuJ mice were obtained from The Jackson Laboratory (Bar Harbor, ME, United States) and housed at 22 °C ± 2 °C on a 12-h light/dark cycle (lights on at 6:00 a.m. [ZT0], lights off at 6:00 p.m. [ZT12]). Unless otherwise specified, mice had ad libitum access to standard chow and water. All procedures conformed to the NIH Guide for the Care and Use of Laboratory Animals and were approved by the University of Maryland, Baltimore Institutional Animal Care and Use Committee (IACUC protocol #AUP-00001103). The study was designed and reported in accordance with ARRIVE guidelines.

To investigate the differential effects of simvastatin in fasting versus feeding states, mice were maintained on standard low-fat chow, under which they typically consume ∼80% of their daily food during the dark (active) phase (ZT12–ZT24) and very little during the light (rest) phase (ZT0–ZT12) (Mukherji et al., 2019; Turek et al., 2005). Two complementary dosing regimens were used: oral gavage during the light-phase fasting period (“fasting-phase” dosing) and dietary admixture during the dark-phase feeding period (“feeding-phase” dosing).

Six days before completion of the 5-week simvastatin or control treatment, glucose tolerance was assessed. For the fasting-phase cohort, A/J mice were fasted for 5 h after their daily oral gavage and then subjected to GTT at ZT09 (3:00 p.m.). For the feeding-phase cohort, A/J, C3H/HeJ, and C3H/HeOuJ mice were fasted for 5 h after removal of food, and GTT was performed at ZT05 (11:00 a.m.). Baseline fasting blood glucose was measured from tail-vein blood prior to glucose administration. Mice then received an intraperitoneal injection of glucose (2 g/kg body weight), and blood glucose concentrations were measured from tail-vein samples at 15, 60, 120, and 180 min using an AlphaTRAK glucometer (Zoetis, Parsippany, NJ). Glucose tolerance was quantified as the area under the curve (AUC) using trapezoidal integration (Solberg et al., 2006; Zhang et al., 2024).

Three days before completion of the 5-week simvastatin or control treatment, glycerol tolerance was evaluated in the same cohorts. For the fasting-phase cohort, A/J mice were fasted for 5 h after their daily oral gavage and tested at ZT09 (3:00 p.m.). For the feeding-phase cohort, A/J, C3H/HeJ, and C3H/HeOuJ mice were fasted for 5 h after removal of food, and testing was performed at ZT05 (11:00 a.m.). Baseline fasting blood glucose was measured from tail-vein blood, after which mice received an intraperitoneal injection of glycerol (1.5 g/kg body weight). Plasma glucose levels were measured from tail-vein blood at designated time points after injection to assess glycerol tolerance (Kim et al., 2017).

Plasma cholesterol fractions (HDL-C, total cholesterol [TC], LDL-C, unesterified cholesterol [UC]) and triglycerides were quantified using enzymatic kits (MilliporeSigma) according to the manufacturer’s protocols. Reference calibrators supplied with these kits were analytical grade with certified purity ≥95% according to the manufacturers’ certificates of analysis.

Liver lipids were extracted using a modification of the Bligh and Dyer method (Zhang et al., 2019). Approximately 100 mg of liver tissue was homogenized in 4 mL of chloroform/methanol (2:1) and incubated overnight at room temperature. After adding 800 µL of 0.9% saline, samples were centrifuged at 2,000 g for 10 min. The organic phase was collected, dried under vacuum, and dissolved in butanol/(Triton X-100/methanol [2:1]) (30:20). Triglycerides were quantified using a colorimetric assay (Sigma), and free fatty acids were measured using FFA kits (Wako Chemicals). All lipid standards used for calibration were analytical grade with manufacturer-certified purity ≥95%.

Hepatic 24(S)-hydroxycholesterol (24S-HC) levels were measured using a 24(S)-hydroxycholesterol ELISA kit (Abcam, #ab204530). Approximately 200 mg of liver tissue was homogenized in 3 mL of 95% ethanol. The homogenate was centrifuged, pellets were resuspended in 1 mL ethanol/dichloromethane (1:1, v/v), and the mixture was centrifuged again. Supernatants from both steps were combined and evaporated to dryness using a rotary evaporator. Dried extracts were reconstituted in 16 μL 95% ethanol and 484 μL assay buffer, then diluted as needed in assay buffer before ELISA. The 24S-HC reference standard supplied with the kit has a purity ≥95% per the manufacturer’s specifications and was used to generate the calibration curve.

Huh-7 human hepatoma cells, originally provided by Dr. Liqing Yu, were maintained at 37 °C in a humidified incubator with 5% CO₂. Cells were grown in Dulbecco’s Modified Eagle Medium (DMEM) supplemented with 10% fetal bovine serum (FBS), 100 U/mL penicillin, and 100 μg/mL streptomycin and were used for experiments at subconfluence.

For fasting-mimetic conditions (Figures 3A,B,E; Supplementary Figure S3), cells were washed twice with 1× PBS, incubated overnight in serum-free DMEM, and then treated for 24 h with 5 µM simvastatin alone or in combination with one or more of the following: 50 µM oleic acid (MilliporeSigma; C18:1, #O1383, purity >99% by GC); 10 µM GW6471 (MedChemExpress; #HY-15372, purity 99%), a selective PPARα antagonist; 500 µM leucine (MedChemExpress; #HY-N0486, purity 98%), which suppresses autophagy initiation via mTORC1 activation; or 50 µM leupeptin (MedChemExpress; #HY-18234A, purity 99.39%), a cysteine/serine/threonine protease inhibitor that blocks autophagic flux (Chen et al., 2021; Desideri et al., 2023; Sun et al., 2021).

For serum-replete conditions (Figures 4A,C), cells were maintained in DMEM containing 10% FBS and treated for 8 h with 50 µM oleic acid, 250 µM leucine, or 10 nM insulin, with or without 10 µM simvastatin. After treatment, total RNA was isolated using TRIzol reagent according to the manufacturer’s instructions.

Liver tissues were snap-frozen in liquid nitrogen at the time of dissection, stored at −80 °C, then cut into ∼50-mg pieces on dry ice and homogenized in TRIzol reagent (Invitrogen) using a motorized tissue homogenizer. Total RNA was extracted and purified according to the manufacturer’s instructions, and 1–3 µg of RNA were used for cDNA synthesis with the iScript™ Advanced cDNA Synthesis Kit (Bio-Rad). Quantitative real-time PCR was performed with iTaq™ Universal SYBR® Green Supermix (Bio-Rad) on a StepOnePlus (Applied Biosystems) or CFX96 (Bio-Rad) instrument.

Target transcripts included genes involved in cholesterol and HDL metabolism, autophagy, and SREBP/LXR/PPARα signaling (e.g., Apoa1, Apoa2, Pltp, Srebf2, Sqle, Lxrα, Ppara, and others as listed in Supplementary Table S1). For mouse tissues, expression levels were normalized to the housekeeping genes B2m and Hprt; for human Huh-7 cells, normalization was performed to B2M and HPRT1. Relative mRNA abundance was calculated using the ΔΔCt method.

Statistical analyses were performed using Prism (GraphPad Software, San Diego, CA). Two-way ANOVA was applied for experiments involving multiple groups, followed, when appropriate, by pairwise Student’s t-tests when significant interactions were detected (p < 0.05). Exact n values for each group are provided in the figure legends. Data are presented as mean ± standard error of the mean (SEM), and significance levels from pairwise comparisons are indicated in the figures.

Under standard chow conditions, mice consume approximately 80% of their food during the dark (active) feeding phase (ZT12–ZT24) and minimally during the light (rest) fasting phase (ZT0–ZT12) (Mukherji et al., 2019; Turek et al., 2005). Capitalizing on this natural diurnal rhythm, we investigated whether fasting versus feeding states would differentially modulate simvastatin’s lipid and glucose regulatory effects in A/J mice over 5 weeks. We used two dosing strategies: simvastatin incorporated into chow (0.1 g/kg food weight; formulation D11060903i, Research Diets), ensuring drug ingestion during the active feeding phase, and oral gavage of an equivalent dose (16 mg/kg body weight) at 10:00 a.m. (ZT04), coinciding with the light-phase fasting period (Figures 1A,B). A typical 25-g mouse consumes approximately 3–5 g of food daily, corresponding to about 16 mg/kg body weight, which aligns with the orally gavaged dose. This approach preserved normal feeding rhythms and allowed a direct comparison of simvastatin’s effects when co-administered with food versus during fasting. Mice were housed on a 12-h light/dark cycle (lights on at 6:00 a.m. [ZT0] and off at 6:00 p.m. [ZT12]) with ad libitum access to food and water. The 16 mg/kg dose was chosen to approximate an 80 mg human equivalent daily dose (Zhang et al., 2024), maintaining translational relevance.

After 5 weeks, we collected plasma and tissues at two time points to capture the drug’s effects in distinct metabolic states: the fasting-state group (ZT09, 3:00 p.m.), 5 hours after the last oral gavage, and the feeding-state group (ZT05, 11:00 a.m.), 5 hours after the active eating period. Mice receiving simvastatin by gavage during fasting showed a modest increase in HDL-C (p = 0.07), with no significant changes in total cholesterol (TC), unesterified cholesterol (UC), or triglycerides (TG) (Figure 1C). Low-density lipoprotein cholesterol (LDL-C) also remained unchanged, consistent with the characteristically low LDL-C in rodents (which lack cholesteryl ester transfer protein, CETP) (Steenbergen et al., 2010). Notably, fasting-phase simvastatin markedly impaired glucose metabolism, as evidenced by higher glucose area under the curve (AUC) values in glucose tolerance tests (GTT) in 6 days before completion of the 5-week simvastatin or control treatment (Figure 1C). In contrast, co-administration of simvastatin with food significantly lowered HDL-C levels and improved glucose homeostasis, indicated by reduced fasting blood glucose and enhanced glucose tolerance (Figure 1D).

To determine whether these time-of-day–dependent effects are restricted to a single sampling point or reflect broader alterations in circadian lipid homeostasis, we next profiled 24-h plasma lipid dynamics in a separate cohort. Serial sampling across the light period (0:00, 6:00, 12:00, and 18:00 h) revealed robust nyctohemeral variation in HDL-C, and confirmed that simvastatin given in the fasting state versus with food produced consistently opposite effects on HDL-C throughout the daytime interval (06:00–18:00) (Supplementary Figure S1). Fasting-phase simvastatin shifted HDL-C upward relative to controls, whereas feeding-phase simvastatin shifted HDL-C downward across these time points, reinforcing the concept that dosing in the fasted versus fed state drives divergent HDL responses over the entire active phase rather than at a single clock time. Together, these data indicate that administering simvastatin during fasting versus feeding exerts opposing effects on HDL and glucose metabolism.

We also assessed whether these interventions altered the hepatic molecular clock. Hepatic mRNA levels of two core clock genes, Bmal1(Arntl) and Per2, were measured in all four primary groups (fasting-vehicle, fasting-simvastatin, feeding-vehicle, feeding-simvastatin) (Supplementary Figure S2). Bmal1 and Per2 displayed the expected anti-phasic pattern in relation to the light–dark cycle (Greco et al., 2021), but their expression did not differ substantially between vehicle and simvastatin within a given nutritional state. These findings suggest that our dosing protocols did not markedly re-phase the hepatic clock and support the conclusion that feeding status—rather than disruption of intrinsic circadian machinery—is the dominant driver of the divergent metabolic responses to simvastatin.

To determine whether the opposing effects of simvastatin on circulating HDL-C reflect coordinated changes in HDL biogenesis, we prospectively quantified hepatic expression of genes involved in HDL synthesis (apolipoprotein A1 [Apoa1], Apoa2, Apom, and ATP-binding cassette subfamily A member 1 [Abca1]), HDL maturation (lecithin–cholesterol acyltransferase [Lcat] and phospholipid transfer protein [Pltp]), and HDL clearance (scavenger receptor class B type 1 [Scarb1], encoding SR-B1) (Paththinige et al., 2017). When simvastatin was administered during fasting, Pltp expression increased significantly and Apoa1 showed a modest increase (p < 0.1), whereas Apoa2, Apom, Abca1, Lcat, and Scarb1 remained unchanged (Figure 2A). By contrast, simvastatin administration during feeding suppressed both Apoa1 and Apoa2. Thus, our a priori analysis of HDL pathway genes confirms that fasting- and feeding-phase simvastatin differentially remodel the hepatic transcriptional program governing HDL synthesis and remodeling, consistent with the divergent HDL-C phenotypes (Figures 1C,D, 2A).

Because Apoa1, Apoa2, and Pltp are regulated by PPARα ligands such as fibrates and because statins can activate PPARα to induce APOA1 in human cells, we hypothesized that nutritional state would dictate how simvastatin engages PPARα to control these HDL-related genes (Bouly et al., 2001; Han et al., 2006; Martin et al., 2001). Consistent with this model, fasting-phase simvastatin significantly upregulated hepatic PPARα and its canonical targets, including carnitine palmitoyltransferase 1A (Cpt1a), carnitine palmitoyltransferase 2 (Cpt2), acyl-CoA synthetase long-chain family member 1 (Acsl1), acyl-CoA dehydrogenase long chain (Acadl), and acyl-CoA oxidase 1 (Acox1). In contrast, simvastatin co-administered with food reduced the expression of several key PPARα targets, including Cpt1a, acyl-CoA dehydrogenase medium chain (Acadm), and acyl-CoA synthetase long-chain family member 5 (Acsl5) (Figure 2B). Together with the Apoa1, Apoa2, and Pltp data (Figure 2A), these findings indicate that fasting and feeding states critically shape simvastatin-mediated PPARα activation and, in turn, transcriptional regulation of HDL synthesis genes.

Beyond lipid handling, PPARα plays a major role in glucose homeostasis, particularly by promoting glycerol-fueled gluconeogenesis (Patsouris et al., 2004). To evaluate how nutritional state modifies this axis, we examined PPARα-responsive genes in the glycerol pathway. Fasting-phase simvastatin significantly enhanced hepatic expression of glycerol-3-phosphate dehydrogenase 1 (Gpd1), glycerol kinase (Gk), and aquaporin 9 (Aqp9), whereas feeding-phase simvastatin decreased their expression (Figure 2C). In contrast, expression of core gluconeogenic enzymes phosphoenolpyruvate carboxykinase 1 (Pck1) and glucose-6-phosphatase catalytic subunit (G6pc) was unchanged by simvastatin in either nutritional state (Figure 2D). Thus, simvastatin dosing during fasting versus feeding differentially modulates PPARα-dependent glycerol utilization without directly altering the gluconeogenic program, providing a mechanistic basis for the distinct glycemic responses observed in vivo (Figures 1C,D).

To further test whether PPARα activation is required for simvastatin’s effects on HDL-related genes and glycerol metabolism, we treated Huh-7 hepatoma cells with simvastatin in the presence or absence of the selective PPARα antagonist GW6471 (Sun et al., 2021). Simvastatin alone robustly increased APOA1 and PLTP mRNA, as well as the glycerol pathway gene GK, whereas co-treatment with GW6471 largely abolished these responses; GW6471 alone had minimal effects (Supplementary Figure S3). These pharmacologic data support a causal role for PPARα in mediating simvastatin-induced upregulation of HDL synthesis genes and glycerol-handling enzymes in hepatocytes, reinforcing the in vivo conclusion that PPARα links nutritional state to simvastatin’s effects on HDL-C and glucose homeostasis.

We next asked how simvastatin activates PPARα under fasting-like conditions. Serum-starving Huh-7 cells overnight and then exposing them to increasing doses of simvastatin led to a dose-dependent induction of PLTP, APOA1, and GPD1 (Figure 3A), mirroring the fasting-phase effects observed in vivo (Figure 2A). Although free fatty acids (FFAs) generated by fasting-induced adipose lipolysis are known PPARα ligands (Bideyan et al., 2021), treatment of Huh-7 cells with oleic acid alone did not increase PLTP, APOA1, or GPD1 expression, suggesting that FFAs require additional signals to fully engage PPARα. Notably, when oleic acid was combined with simvastatin, APOA1 expression was markedly higher than with simvastatin alone (Figure 3B), indicating that simvastatin can potentiate FFA-mediated activation of the PPARα–HDL gene program.

Autophagy has emerged as a key facilitator of PPARα signaling by degrading nuclear corepressors such as NCoR1 and HDAC3 (Iershov et al., 2019; Saito et al., 2019), and SREBP-2 transactivation can itself trigger autophagy (Seo et al., 2011). In liver from fasting mice, simvastatin markedly increased SREBP-2 expression and its downstream cholesterol synthesis targets by ∼1–3-fold (Figure 3C), coincident with upregulation of key autophagy-related genes, including Atg7, Atg12, and Becn1 (Figure 3D). These findings suggest that simvastatin activates a SREBP-2–dependent autophagy program under fasting conditions.

To directly test whether autophagic flux is required for simvastatin’s transcriptional effects, serum-starved Huh-7 cells were treated with simvastatin together with pharmacologic inhibitors of autophagy. Leucine, which activates mTORC1 and suppresses autophagy (Chen et al., 2021), completely abolished simvastatin-induced APOA1 expression (Figure 3E, left). Leupeptin, a cysteine/serine/threonine protease inhibitor that blocks lysosomal degradation and causes accumulation of autolysosomes (Desideri et al., 2023; Yang et al., 2013), significantly attenuated—but did not fully abolish—the simvastatin-mediated increase in APOA1 mRNA (Figure 3E, right). Thus, inhibition of either the initiation phase of autophagy (via mTORC1 activation) or the degradative phase (via lysosomal blockade) markedly blunts simvastatin-induced APOA1 upregulation, supporting a requirement for intact autophagic flux. Collectively, these data support a model in which simvastatin triggers SREBP-2–dependent autophagy during fasting, thereby enhancing FFA-mediated PPARα activation and ultimately modulating HDL levels and hepatic glucose metabolism in vivo.

Co-administration of simvastatin with food suppressed PPARα activation, contrasting with the enhanced activation observed when simvastatin was given during fasting (Figures 2B,C). To identify feeding-associated factors that could antagonize PPARα, we first tested common nutrients and hormones in Huh-7 cells under serum-replete conditions (10% FBS). Neither leucine nor insulin altered simvastatin-induced APOA1 expression (Figure 4A), and neither oleic acid nor high glucose affected simvastatin’s transcriptional effects (data not shown). These observations suggest that typical postprandial nutrient and insulin signals are insufficient to explain the attenuation of PPARα activation by feeding.

Recent work instead implicates postprandial absorption of gut microbiota–derived LPS in transient elevations of circulating LPS, a phenomenon termed metabolic endotoxemia (Cani et al., 2007; Mohammad and Thiemermann, 2020). LPS can be incorporated into intestinally secreted lipoproteins such as chylomicrons and thereby enter the systemic circulation (Clemente-Postigo et al., 2012). We therefore measured plasma LPS levels during fasting (ZT03–09) and feeding (ZT15–21) and found that, irrespective of simvastatin treatment, circulating LPS peaked during feeding and was significantly lower during fasting (Figure 4B). To test whether LPS directly modulates simvastatin’s actions, we co-treated Huh-7 cells with simvastatin and LPS under serum-replete conditions. LPS co-treatment abolished simvastatin-induced upregulation of PLTP, APOA1, and GK (Figure 4C). These findings indicate that microbiota-derived LPS, which rises with feeding, can antagonize simvastatin-driven PPARα activation and HDL/glycerol gene expression, providing a mechanistic explanation for the diminished PPARα signaling observed when simvastatin is administered in the fed state (Figures 2A–C).

To investigate how feeding-driven LPS might modulate simvastatin’s regulation of PPARα activity—which governs fasting glucose and HDL metabolism—we reanalyzed transcriptomic data from differentiated HepaRG (dHepaRG) cells treated with LPS for 6 hours (GEO: GSE230325) (Ehle et al., 2024). dHepaRG cells retain key hepatic features, including hepatocyte-like morphology and the expression of metabolic enzymes, nuclear receptors, and drug transporters (Ehle et al., 2024). Our analysis covered both protein-coding and non-coding RNAs, with 43,892 genes detected in total; a volcano plot revealed significant transcriptional changes (Figure 5A).

Gene Set Enrichment Analysis (GSEA) of LPS-upregulated genes showed enrichment in pathways related to infection, cytokine-receptor interactions, and Toll-like receptor signaling—specifically involving the LPS receptor TLR4 (Figure 5B). By contrast, LPS-downregulated genes were enriched in pathways tied to bile acid metabolism, xenobiotic and fatty acid metabolism, peroxisomal functions, and estrogen response (Chen et al., 2013; Kuleshov et al., 2016; Mootha et al., 2003; Subramanian et al., 2005; Xie et al., 2021). We focused on two salient changes: the reduction in bile acid metabolism-related genes and the induction of TLR signaling, aligning with previous reports that LPS suppresses bile acid synthesis and that statins also influence bile acid metabolism (Khovidhunkit et al., 2004; Schonewille et al., 2016). Additionally, LPS-activated TLR4 impacts lipid metabolism, including bile acid pathways (Castrillo et al., 2003).

In the liver, cholesterol is converted to bile acids through two major routes. In the classic (neutral) pathway, cholesterol 7α-hydroxylase (CYP7A1) catalyzes the rate-limiting step to generate 7α-hydroxycholesterol (7α-HC), which is further processed by 3β-hydroxy-Δ⁵-C₂₇-steroid oxidoreductase (HSD3B7) to form cholic acid and chenodeoxycholic acid (CDCA) (Pandak and Kakiyama, 2019). In the acidic pathway, sterol 27-hydroxylase (CYP27A1) produces (25R)26-hydroxycholesterol (27-HC), which—together with other oxysterols—is further hydroxylated by oxysterol 7α-hydroxylase (CYP7B1) en route to CDCA (Pandak and Kakiyama, 2019). Cholesterol 24(S)-hydroxylase (CYP46A1) generates 24(S)-hydroxycholesterol (24S-HC), best characterized in neurons but also expressed in liver, where it exerts regulatory effects on lipid metabolism (Si et al., 2020). In LPS-treated dHepaRG cells, expression of CYP7A1, CYP46A1, and CYP39A1 was markedly reduced, with a more modest decline in CYP27A1 (p = 0.07), whereas CYP7B1 was induced more than fivefold (Figure 5C). These data suggest that LPS not only suppresses the classic pathway but also reshapes the acidic pathway by downregulating oxysterol-producing enzymes (CYP27A1, CYP46A1) while upregulating the oxysterol-metabolizing enzyme CYP7B1, thereby favoring depletion of regulatory oxysterols such as 27-HC.

We next examined how bile acid and oxysterol metabolism are altered in vivo when simvastatin is administered during the feeding phase, a context in which circulating LPS is elevated (Figure 4B). Relative to fasting, feeding alone decreased hepatic expression of the oxysterol-producing enzymes Cyp7a1, Cyp27a1, and Cyp46a1 (Figure 5D). Feeding-phase simvastatin further increased the expression of the oxysterol-catabolizing enzymes Hsd3b7 and Cyp7b1 (Figure 5D). Although Cyp39a1 was unchanged, feeding-phase simvastatin further suppressed Cyp46a1, in contrast to the upregulation observed under fasting-phase simvastatin treatment (Figure 5D), indicating that feeding-related signals can override simvastatin-induced Cyp46a1 expression.

Consistent with these in vivo findings, simvastatin markedly increased CYP46A1 expression in Huh-7 cells, an effect that was strongly attenuated by co-treatment with LPS (Figure 5E). In mouse liver, direct quantification of 24(S)-hydroxycholesterol (24S-HC) using an ELISA kit showed that simvastatin decreased 24S-HC levels when administered with food, whereas it increased 24S-HC under fasting conditions (Figure 5F). Taken together, these findings indicate that feeding-driven LPS signaling remodels hepatic oxysterol metabolism—suppressing oxysterol synthesis while enhancing oxysterol catabolism—such that simvastatin given in the fed state depletes hepatic oxysterol pools (Figure 5G). Because these oxysterols serve as endogenous ligands for LXR and thereby regulate Apoa1, Apoa2, and Pltp expression, this LPS–statin interaction provides an upstream mechanism by which nutritional state and metabolic endotoxemia shape simvastatin’s impact on HDL synthesis and PPARα-dependent glucose regulation.

Oxysterols—such as 24S-, 25-, and 27-hydroxycholesterol—serve as physiological ligands for liver X receptors (LXRα and LXRβ), activating them at concentrations typically found in vivo (Fu et al., 2001; Janowski et al., 1996; Lehmann et al., 1997). To determine whether feeding-phase simvastatin—which depletes hepatic oxysterols—alters LXR activity, we first measured the expression of canonical LXR targets that contribute to HDL biogenesis and cholesterol efflux. Notably, Abcg1, Abcg5, and Srebf1 (encoding SREBP-1c) were significantly repressed when simvastatin was given with food, but not during fasting (Figures 3C, 6A).

Although activation of Tlr3/Tlr4 by E. coli, influenza A, or ligands such as poly I:C (TLR3 agonist) and lipid A (TLR4 agonist) can block LXR signaling in macrophages via IRF-3 competition for the co-activator p300/CBP, we found that simvastatin co-administration with food significantly downregulated IRF-3 and NF-κB targets (Il15, Ppp2r1b, Crp, Saa1) (Figure 6B). These results suggest that the IRF-3 pathway is unlikely to mediate the synergistic inactivation of LXR by LPS and simvastatin in the liver, and instead point to oxysterol depletion as the primary mechanism.

Consistent with prior reports that statin exposure can reduce both membrane-bound and nuclear SREBP-1c in an LXR-dependent manner (DeBose-Boyd et al., 2001; Liang et al., 2002; Rong et al., 2017), simvastatin treatment during feeding—but not fasting—led to decreased SREBP-1c expression (Figure 3C). Reduced SREBP-1c transactivation was further evident in the feeding group, where SREBP-1c targets involved in lipogenesis (Elovl2, Elovl5), triglycerides synthesis (Gpat3, Agpat5), and phospholipid formation (Chpt1, Cept1) were suppressed (Figures 6C,D). Quantitatively, feeding-phase simvastatin did not alter hepatic free fatty acid levels but significantly lowered hepatic triglycerides content, consistent with diminished SREBP-1c activity (Figures 6E,F).

Because de novo lipogenesis—including phosphatidylcholine synthesis via choline/ethanolamine phosphotransferase 1 (Cept1)—is required for full PPARα activation (Guan et al., 2018; Zayed et al., 2021), our mechanistic data support a model in which feeding-driven LPS, together with simvastatin, depletes hepatic oxysterol pools and inactivates LXR. This, in turn, dampens SREBP-1c activity, reduces de novo lipogenesis (Figures 6C,D), and limits PPARα activation both during the feeding period and throughout the subsequent daytime fast when HDL-C and glucose were measured (Figure 1D; Supplementary Figure S1). Consequently, mice receiving simvastatin with food display persistently lower fasting blood glucose and HDL-C, in sharp contrast to the HDL-raising, PPARα-activating, and dysglycemic profile observed when simvastatin is administered during the fasting phase.

We observed that LPS suppresses hepatic expression of CYP7A1, CYP27A1, and CYP46A1, thereby priming the liver for further oxysterol depletion by simvastatin during feeding. Previous work has shown that LPS-mediated downregulation of hepatic cytochrome P450 (P450) mRNAs is entirely TLR4-dependent (Chaluvadi et al., 2009). To determine whether TLR4 is also essential for modulating simvastatin’s effects on HDL cholesterol and glucose homeostasis across feeding and fasting states, we used C3H/HeJ mice harboring a spontaneous Tlr4 mutation (Tlr4 ^Lps-d ), which renders them hyporesponsive to LPS (Kamath et al., 2003). These mice, along with wild-type C3H/HeOuJ controls, were fed simvastatin mixed in chow for 5 weeks.

In wild-type C3H/HeOuJ mice, simvastatin co-administration with food significantly improved glucose tolerance while modestly lowering HDL cholesterol (p = 0.08), mirroring the results in A/J mice (Figures 1B, 7A,B). In contrast, TLR4-deficient C3H/HeJ mice exhibited significantly elevated HDL cholesterol, total cholesterol, and free cholesterol after simvastatin treatment during feeding (Figure 7C). Moreover, the TLR4-mutant mice developed impaired glucose homeostasis under these conditions, as indicated by elevated fasting blood glucose and glucose intolerance (Figure 7D)—a phenotype that closely resembled A/J mice given simvastatin during fasting (Figure 1D). These findings highlight the critical role of TLR4 in mediating feeding-driven LPS modulation of simvastatin’s regulatory effects on HDL cholesterol and glucose metabolism. Without functional TLR4 signaling, the normal synergy between LPS and simvastatin is disrupted, shifting the metabolic outcomes toward elevated HDL and impaired glucose tolerance.

To elucidate how TLR4 modulates simvastatin’s metabolic effects, we compared hepatic expression of key bile acid/oxysterol-metabolizing enzymes between TLR4-deficient (C3H/HeJ) and wild-type (C3H/HeOuJ) mice by quantitative PCR. In wild-type mice, simvastatin treatment suppressed Cyp46a1 and upregulated Cyp7b1 (Figure 8A). In contrast, TLR4 deficiency elevated the basal expression of oxysterol-generating enzymes—including Cyp7a1, Cyp27a1, and Cyp46a1. Moreover, in TLR4-deficient livers, simvastatin further increased Cyp7a1 and Cyp46a1 expression while downregulating the oxysterol-metabolizing genes Hsd3b7 and Cyp39a1 (Figure 8A), suggesting that loss of TLR4 prevents simvastatin-induced depletion of hepatic oxysterols and instead favors their accumulation.

TLR4 deficiency also counteracted simvastatin-mediated inhibition of the LXR/SREBP-1c axis. This was evidenced by increased expression of Apoe, Gpat4, Cept1, and Cpt1a, which were otherwise reduced in simvastatin-treated wild-type mice (Figure 8B). These findings suggest that, in the absence of TLR4, simvastatin promotes LXR ligand synthesis and activation rather than suppressing it.

Analysis of the PPARα pathway further showed that simvastatin enhanced PPARα target genes relevant to HDL remodeling and lipid handling (Pltp, Gk, Cpt1a) in TLR4-deficient mice but downregulated these genes in wild-type controls (Figure 8C). To verify that PPARα contributes to dysglycemia in simvastatin-treated TLR4-deficient mice, we conducted glycerol tolerance tests 3 days before completion of the 5-week simvastatin or control treatment (Sumara et al., 2012). Glycerol administration led to higher circulating glucose levels in TLR4-deficient C3H/HeJ mice compared with wild-type C3H/HeOuJ mice, confirming that PPARα-driven, glycerol-fueled gluconeogenesis exacerbates glucose intolerance in TLR4-deficient mice given simvastatin (Figure 8D).

Collectively, these results demonstrate that TLR4 is essential for feeding-driven LPS regulation of simvastatin’s metabolic actions. In TLR4-deficient mice, simvastatin’s usual effects on HDL cholesterol and glucose metabolism are reversed because of disrupted SREBP-1c and PPARα pathways, underscoring the pivotal role of TLR4 in modulating simvastatin’s influence on hepatic lipid and glucose homeostasis.