Introduction

Sepsis-associated acute kidney injury (SA-AKI) is a common complication in critically ill patients and is linked to prolonged hospitalization, higher risk of chronic kidney disease (CKD), and increased mortality [1,2,3,4]. The 28th Acute Disease Quality Initiative (ADQI) consensus specifically defines SA-AKI as acute kidney injury (AKI) that fulfills the Kidney Disease: Improving Global Outcomes (KDIGO) criteria and occurs within 7 days of sepsis onset, in which sepsis is considered the primary and proximate cause of kidney dysfunction [5]. Despite its clinical severity, there are currently no effective targeted therapies; management remains largely supportive, focusing on infection control and renal replacement therapy when required [4,5,6]. The lack of specific therapy for SA-AKI highlights an urgent need to identify novel therapeutic approaches.

Recent studies highlight metabolic reprogramming as a central mechanism in SA-AKI [5]. Normally, proximal tubular epithelial cells (TECs) rely on mitochondrial fatty acid oxidation (FAO) as their main energy source [7, 8]. During sepsis, FAO is suppressed and glycolysis predominates, leading to mitochondrial dysfunction, lipid accumulation, and aggravated tubular injury [9, 10]. The buildup of fatty acids and lipid droplets in TECs exacerbates cellular injury and inflammation, contributing to tubular damage and fibrosis [10]. Preserving FAO has been shown to protect against kidney damage, suggesting that restoration of energy metabolism may offer therapeutic benefit [7, 9].

Fenofibrate, a clinically approved peroxisome proliferator-activated receptor-α (PPARα) agonist, enhances FAO and improves mitochondrial fatty acid utilization [7, 9]. Preclinical studies indicate that PPARα activation can increase adenosine triphosphate (ATP) generation while reducing toxic lipid byproducts and inflammation in kidney tissue [9, 11]. Fenofibrate, as a PPARα agonist, has demonstrated renoprotective effects in experimental models of SA-AKI – for instance, attenuating tissue damage in cecal ligation and puncture (CLP)-induced models [11, 12]. Based on evidence, we hypothesize that fenofibrate can bolster FAO and restore energy homeostasis in septic kidneys, thereby alleviating SA-AKI. This study aims to explore how fenofibrate influences metabolic reprogramming and kidney damage in SA-AKI, with the goal of uncovering a new pharmacological strategy for this serious condition.

Methods

Animal model

Male C57BL/6 mice (6–8 weeks old, 20–25 g) were acquired from Charles River Laboratories and housed in specific pathogen-free (SPF) conditions with a 12-hour light/dark cycle. Following a one-week acclimation period, the mice were randomly assigned to three groups (n = 6 per group): control group (C group), lipopolysaccharide (LPS)-induced SA-AKI group (L group), and fenofibrate treatment group (FF group). Mice in the FF group received a standard chow diet supplemented with 0.2% fenofibrate (w/w) (Synergy Bio, China), equivalent to approximately 200 mg/kg/day, for seven consecutive days. The C and L groups were fed a standard chow diet without fenofibrate. In the L and FF groups, SA-AKI was established by administering an intraperitoneal dose of LPS (10 mg/kg; Sigma-Aldrich, USA) mixed with phosphate-buffered saline (PBS; VivaCell, China). The C group received an equal volume of PBS intraperitoneally. All animal procedures were approved by the Institutional Animal Care and Use Committee (Approval No. SYDW2025–052) and conducted following ethical guidelines for laboratory animal use.

Sample collection

Twenty-four hours after LPS administration, all mice were anesthetized by intraperitoneal injection of tribromoethanol (brand name Avertin, Sigma-Aldrich), prepared according to established protocols. A stock solution was made by dissolving 1 g of 2,2,2tribromoethanol in 1 ml of tertamyl alcohol (2methyl2butanol), then diluted to a working solution by adding 0.6 ml of this concentrate to 59.4 ml of sterile deionized water (yielding a 1.0% w/v solution). Each mouse (~20–25 g) received 0.4 ml intraperitoneally, equivalent to approximately 160–200 mg/kg of pure tribromoethanol, providing a surgical plane of anesthesia lasting about 20–30 minutes.

Under sterile conditions, peripheral blood was collected via retro-orbital venous puncture. Immediately following blood collection, mice were euthanized by cervical dislocation while still under deep anesthesia, as confirmed by the absence of reflex responses (e.g., toe-pinch). This procedure complies with the American Veterinary Medical Association (AVMA) guidelines, which recommend a humane physical method following chemical anesthesia to ensure rapid and painless death. Blood samples were left at room temperature for 30 minutes to clot, then centrifuged at 3000 rpm for 10 minutes at 4 °C to separate the serum for further biochemical testing. The right kidney was quickly frozen in liquid nitrogen and kept at −80 °C for molecular and biochemical analysis, while the left kidney was preserved in 10% neutral-buffered formalin for histopathological examination.

Tissue sectioning and staining procedures

Following collection, kidney tissues were preserved in 10% neutral-buffered formalin, dehydrated using a graded ethanol series, embedded in paraffin, and cut into 4 μm thick sections. Hematoxylin and eosin (H&E) staining was performed to evaluate renal histomorphology. For periodic acid–Schiff (PAS) staining, sections were treated with Schiff reagent for 10 minutes, washed under running water for 5 minutes, counterstained with Mayer’s hematoxylin for 3 minutes, and then rinsed again for 5 minutes. The stained sections were then dehydrated, cleared with xylene, and mounted with coverslips for microscopic observation.

To evaluate tubular injury, histological features such as epithelial simplification, brush border loss, apical blebbing, epithelial cell detachment, tubular dilation, and cast formation were assessed. For each specimen, ten randomly selected, non-overlapping cortical or outer medullary fields were examined at × 400 magnification. Two renal pathologists independently and blindly graded the samples using a validated semi-quantitative scale: 0, normal tubules without visible damage; 1, ≤10% injured; 2, 11–25% injured; 3, 26–75% injured; 4, > 75% injured. Scores from each field were then used for intergroup comparisons.

To evaluate lipid accumulation in kidney tissues, Oil Red O staining was performed on frozen kidney sections. Cryosections (10 μm) were fixed in 10% neutral-buffered formalin for 10 minutes and washed with distilled water. The sections were then incubated with freshly prepared Oil Red O solution for 10 minutes, briefly differentiated in 60% isopropanol, and rinsed with distilled water again. Counterstaining was done using Harris hematoxylin for 1–2 minutes, followed by a thorough rinse with tap water. The sections were mounted with an aqueous medium and examined under a light microscope.

Assessment of renal function and inflammation

Renal function was assessed by quantifying serum creatinine (Scr) and blood urea nitrogen (BUN) levels with commercial colorimetric assay kits (Nanjing Jiancheng Bioengineering Institute, China), according to the manufacturer’s guidelines.

Scr concentrations were determined using a creatininase–creatinase–sarcosine oxidase–peroxidase coupled enzymatic method. Serum samples (6 μL) were added to enzyme reagent A and incubated at 37 °C for 5 minutes. Enzyme reagent B was introduced, and the mixture was incubated at 37 °C for 5 minutes. Absorbance was measured at 546 nm with a microplate reader, and creatinine concentrations were determined by calculating the absorbance change using a calibration standard of 442 μmol/L.

BUN was measured using the urease–phenol–hypochlorite method. In brief, serum samples were incubated with urease-containing buffer at 37 °C for 10 minutes to hydrolyze urea into ammonium ions. Following the addition of phenol chromogenic reagent and alkaline hypochlorite, the mixture was incubated at 37 °C for 10 minutes. The developed blue color was measured at 640 nm, and BUN levels were determined by referencing a 10 mmol/L standard solution.

Quantitative real-time polymerase chain reaction (qPCR) was used to measure the expression levels of messenger ribonucleic acid (mRNA) for renal injury markers, including kidney injury molecule-1 (KIM-1) and neutrophil gelatinase-associated lipocalin (NGAL), as well as pro-inflammatory cytokines such as interleukin-1β (IL-1β), interleukin-6 (IL-6), and tumor necrosis factor-α (TNF-α). Total RNA was isolated from kidney tissues with RNA-Solv Reagent (Omega Bio-tek, USA) according to the manufacturer’s protocol. Reverse transcription was carried out using 2 μg of total RNA and the TransScript First-Strand cDNA Synthesis SuperMix (Roche). qPCR was conducted using the TransStart Top Green qPCR SuperMix (TransGen Biotech, China). Gene expression was adjusted to glyceraldehyde-3-phosphate dehydrogenase (GAPDH) and determined via the comparative Ct method (2^–ΔΔCt). The primer sequences for amplification are provided in Table 1 (RuiBiotech, China).

Table 1 Primers that were used in this study

Immunohistochemistry

For immunohistochemical detection of KIM-1 and NGAL, paraffin-embedded renal sections (4 μm) were deparaffinized in xylene, rehydrated through graded ethanol, and subjected to heat-induced antigen retrieval in citrate buffer (pH 6.0) at ~ 95 °C for 15 min. Endogenous peroxidase was blocked with 3% hydrogen peroxide for 10 min, followed by 5% normal goat serum for 30 min at room temperature. Sections were incubated overnight at 4 °C with primary antibodies against KIM-1 [1: 200] and NGAL [1: 300], then treated with biotin-conjugated secondary antibodies and a streptavidin–horseradish peroxidase complex (30 min each). Visualization was achieved with diaminobenzidine (DAB), and sections were counterstained with Mayer’s hematoxylin, dehydrated, cleared, and mounted for light microscopy. Negative controls without primary antibodies were included.

For semi-quantitative analysis, three non-overlapping sections per group were selected, with two random cortical fields (×200) analyzed per section (six images per group, n = 6). Positive staining areas were quantified using ImageJ by thresholding the brown DAB signal and expressed as a percentage of the total field area.

Energy metabolism & lipid accumulation

Renal energy metabolism and lipid deposition were analyzed by assessing ATP levels, triglycerides (TG), glycerol, and non-esterified fatty acids (NEFA) using commercial colorimetric or luminescent assay kits (Nanjing Jiancheng Bioengineering Institute, China), as per the manufacturer’s guidelines.

ATP concentrations were measured using a firefly luciferase-based chemiluminescence assay kit. Renal tissues were lysed in buffer (5–10 µL/mg tissue), centrifuged at 12,000 ×g for 5 minutes at 4 °C, and the supernatants were analyzed in white 96-well plates. Luminescence was detected with a luminometer. ATP levels were adjusted for total protein and expressed as relative values, with results shown as fold changes in the L and FF groups compared to the C group.

TG levels were quantified using the glycerol-3-phosphate oxidase – phenol aminophenazone technique. Kidney tissues were homogenized in 0.9% saline (1:9, w/v), centrifuged, and 2.5 µL of supernatant was mixed with 250 µL of reaction reagent in a 96-well plate. After incubating at 37 °C for 10 minutes, absorbance was recorded at 500 nm. Results were normalized to protein concentration and reported as mmol/g.

Glycerol levels were determined using a colorimetric assay kit, which quantifies glycerol via enzymatic conversion to hydrogen peroxide and chromogenic dye. Each sample (10 µL) was combined with 190 µL of working solution in 96-well plates, and absorbance was recorded at 505 nm. Glycerol concentrations were determined using a standard curve and adjusted for total protein.

NEFA concentrations were determined using a two-step enzymatic procedure. Samples (4 µL) were incubated with reagent 1 followed by reagent 2 at 37 °C, and the optical density was recorded at 546 nm before and after the second incubation. NEFA levels were derived from ΔA values and expressed as mmol/gprot.

Mitochondrial function & FAO capacity

Citrate synthase activity, a surrogate marker of mitochondrial content and function, was quantified with a commercial colorimetric assay kit (Nanjing Jiancheng Bioengineering Institute, China). Renal tissues were processed in extraction buffer at a ratio of 1:9 (g/mL), centrifuged at 10,000 rpm for 10 minutes at 4 °C, and the supernatant was collected. The assay was performed in 96-well plates: reaction mixtures containing buffer, substrate, and chromogenic agent were pre-incubated at 37 °C for 3–5 minutes before adding 10 μL of sample. Absorbance was measured at 412 nm both at baseline (0 min) and after 15 min incubation. Citrate synthase activity was calculated using the change in absorbance over time and normalized to protein concentration.

FAO capacity was assessed using a colorimetric assay kit (Elabscience, China). Kidney tissue was lysed in 0.9% saline (1:9, g/mL), and the supernatant was collected after centrifugation at 10,000 ×g for 15 minutes at 4 °C. The assay was conducted in 96-well plates following the manufacturer’s instructions: 50 μL of the sample was mixed with substrate, cofactor, and chromogenic reagents, then incubated at 37 °C for 30 minutes. Absorbance was recorded at 450 nm using a microplate reader. FAO capacity was determined from a standard curve and adjusted for total protein content.

qPCR for FAO-related genes

To evaluate the transcriptional control of FAO, qPCR was conducted to quantify mRNA levels of key genes involved in FAO, such as PPARα, peroxisome proliferator-activated receptor gamma coactivator 1-alpha (PGC1α), carnitine palmitoyltransferase 1A (CPT1A), carnitine palmitoyltransferase 2 (CPT2), and acyl-CoA oxidase 1 (ACOX1). The experimental procedures for RNA extraction, reverse transcription, and qPCR were the same as described above. Primer sequences are listed in Table 1.

Western blotting

Western blotting was conducted to assess the levels of PPARα, AMP-activated protein kinase (AMPK), phosphorylated AMPK (P-AMPK), PGC1α, CPT1A, CPT2, and ACOX1 expression. Kidney tissues were lysed, and total protein was extracted using standard lysis buffer. Equal protein quantities were separated using 10% SDS-PAGE and transferred to polyvinylidene fluoride membranes. The membranes were blocked with 5% non-fat milk and incubated overnight at 4 °C with primary antibodies against the target proteins and the internal control β-actin (beta-actin) (all from Proteintech Group, Wuhan, China). After incubation with appropriate horseradish peroxidase-conjugated secondary antibodies, protein bands were detected using an enhanced chemiluminescence system. Band intensities were analyzed using densitometry with ImageJ software (National Institutes of Health, Bethesda, MD, USA) and normalized to β-actin.

Immunofluorescence

Immunofluorescence (IF) staining was used to detect AMPK, P-AMPK, PGC1α, PPARα, CPT1A, CPT2, and ACOX1. Paraffin sections were deparaffinized, subjected to antigen retrieval and permeabilization, then washed with PBS (3 × 5 min). After blocking with 5–10% normal goat serum or BSA for 1 h at room temperature, sections were incubated overnight at 4 °C with primary antibodies diluted in blocking buffer. The next day, sections were washed, incubated with fluorophore-conjugated secondary antibodies for 1 h at room temperature in the dark, counterstained with 4′,6-diamidino-2-phenylindole (DAPI), mounted with antifade medium, and imaged using a fluorescence microscope.

For semi-quantitative analysis, three non-overlapping sections per group were used, with two random cortical fields (×400) analyzed per section (six images per group). Mean fluorescence intensity was measured in tubular regions of interest using ImageJ and normalized to DAPI-positive nuclei.

Statistical analyses

Statistical analyses were conducted using GraphPad Prism (version 10.0; GraphPad Software, La Jolla, CA, USA). Data are expressed as mean ± standard error of the mean (SEM). Comparisons between two groups were performed using unpaired Student’s t-tests, and one-way analysis of variance (ANOVA) was used for comparisons among more than two groups.

Results

Fenofibrate alleviates LPS-induced renal histopathological injury

Histological staining confirmed the renoprotective effects of fenofibrate in SA-AKI (Fig. 1A). H&E staining from L group revealed extensive tubular injury, including epithelial cell swelling, cast formation, brush border loss, and prominent interstitial inflammatory infiltration. In contrast, fenofibrate-treated mice exhibited markedly reduced structural damage, with preserved tubular morphology and minimal inflammatory infiltration. PAS staining further showed disrupted tubular basement membranes and loss of brush border integrity in the L group, which were partially restored in the FF group. Semi-quantitative analysis of PAS-stained sections further confirmed the severity of tubular injury, with significantly higher tubular injury scores in the L group compared to the C group, while FF treatment significantly reduced the injury score (Fig. 1C).

Fig. 1
figure 1

(A) Representative histological kidney sections stained with H&E, PAS, and oil red O from C, L, and FF mice (original magnification × 20). H&E and PAS staining revealed severe tubular injury and brush border loss in the L group, which were alleviated in the FF group. Oil red O staining demonstrated marked lipid accumulation in the L group, while fenofibrate treatment reduced lipid deposition in renal tissues. (B) Representative immunohistochemical staining for KIM-1 and NGAL in renal tissues (×200). Strong brown DAB signals were observed in the L group, indicating significant tubular epithelial injury, whereas FF treatment substantially reduced the expression of both markers. (C) semi-quantitative analysis of immunohistochemistry and tubular injury. Semi-quantitative analysis of KIM-1 and NGAL positive staining areas was performed based on immunohistochemistry. Six images per group (three sections × two cortical fields) were analyzed using ImageJ, with the brown DAB-positive area expressed as a percentage of the total field area. FF treatment significantly decreased the positive staining area compared with the L group, confirming histological improvements. In addition, tubular injury scores were obtained from PAS-stained sections by two independent pathologists in a blinded manner, based on the percentage of injured tubules (0–4 scale). The L group showed significantly higher scores than the C group, whereas FF treatment markedly reduced tubular injury

Fenofibrate improves renal function and reduces inflammation

Biochemical and molecular analyses demonstrated that fenofibrate attenuated LPS-induced renal dysfunction and inflammation (Fig. 2). Relative to the C group, the L group exhibited considerable increases in Scr and BUN levels, along with heightened expression of KIM-1 and NGAL. In addition, inflammatory cytokines, such as IL-1β, IL-6, and TNF-α, were markedly elevated. Fenofibrate treatment significantly reduced all of these markers compared to the L group, indicating that fenofibrateF mitigates both renal injury and the associated inflammatory response in SA-AKI. Consistent with qPCR findings, immunohistochemical staining revealed intense KIM-1 and NGAL signals in TECs of the L group, indicative of severe renal injury. In contrast, staining intensity was markedly reduced in the FF group, suggesting attenuated epithelial damage and further supporting the histological protective effect of FF (Fig. 1B–C).

Fig. 2
figure 2

Scr, BUN, kidney injury biomarkers (KIM-1 and NGAL), and pro-inflammatory cytokines (IL-1β, IL-6, and TNF-α) measured in the C, L, and FF groups. Mice in the L group exhibited significant renal dysfunction and inflammation compared to the control, while FF group markedly attenuated renal injury and reduced cytokine levels

Fenofibrate restores energy metabolism and improves lipid metabolic balance

In the L group, renal ATP levels were significantly decreased, while TG content was markedly elevated, reflecting impaired energy production and disrupted lipid utilization. Glycerol and NEFA levels were notably higher in the L group, indicating increased lipolysis but insufficient subsequent oxidation, reflecting metabolic limitations. FF group markedly reversed these changes: ATP production was restored, and tissue levels of TG, glycerol, and NEFA were significantly reduced, suggesting recovery of mitochondrial energy output and improved lipid handling (Fig. 3). These biochemical findings were corroborated by Oil Red O staining, which revealed extensive lipid droplet accumulation in the L group and substantial reduction following fenofibrate administration, indicating attenuated lipid overload and enhanced fatty acid clearance in renal tissues (Fig. 1).

Fig. 3
figure 3

Evaluation of renal energy metabolism, lipid metabolic balance, and FAO capacity in the C, L, and FF groups. Renal ATP levels were significantly reduced in the L group and restored following fenofibrate treatment. TG, glycerol, and NEFA were markedly elevated in the L group, reflecting impaired lipid utilization and excessive lipolysis, while these parameters were significantly decreased in the FF group. Citrate synthase activity, a marker of mitochondrial content and function, was reduced in the L group and recovered after fenofibrate administration. Similarly, FAO capacity was suppressed in the L group but significantly enhanced in the FF group, indicating restoration of mitochondrial oxidative function

Fenofibrate enhances FAO and mitochondrial function

Markers of mitochondrial function and FAO were assessed to further explore fenofibrate’s mechanism of action. Citrate synthase activity, a surrogate marker of mitochondrial content and functional capacity, was significantly reduced in the L group but restored following fenofibrate intervention. Similarly, FAO capacity was markedly suppressed in the L group but significantly improved with fenofibrate treatment, indicating reactivation of renal FAO pathways and amelioration of metabolic dysfunction during SA-AKI (Fig. 3).

Fenofibrate enhances mRNA expression of FAO-related genes

Compared to the C group, the L group showed a significant reduction in the mRNA expression of all five genes (PPARα, PGC1α, CPT1A, CPT2, and ACOX1), consistent with transcriptional repression of mitochondrial lipid metabolism during SA-AKI. Fenofibrate treatment markedly reversed these changes: mRNA levels of PPARα and PGC1α, two key transcriptional regulators of FAO, were significantly upregulated in the FF group. Similarly, expression of the FAO enzymes CPT1A, CPT2, and ACOX1 was also restored to near-control levels (Fig. 4). These findings suggest that fenofibrate activates the FAO program at the transcriptional level.

Fig. 4
figure 4

mRNA expression of fatty FAO-related genes in kidney tissues from C, L, and FF groups. The L group exhibited significantly reduced expression of PPARα, PGC1α, CPT1A, CPT2, and ACOX1, indicating suppression of mitochondrial FAO pathways during SA-AKI. Fenofibrate administration restored the transcription of these genes, with marked upregulation of PPARα and PGC1α, as well as recovery of CPT1A, CPT2, and ACOX1 expression, suggesting reactivation of the renal FAO program at the transcriptional level

Fenofibrate restores the AMPK–PGC1α–PPARα signaling axis and upregulates FAO-related enzymes

In comparison to the C group, mice in the L group exhibited a marked reduction in the ratio of P-AMPK to total AMPK, suggesting diminished AMPK activation during septic stress. This was accompanied by downregulation of PGC1α and PPARα. In parallel, the protein levels of three major FAO enzymes—CPT1A, CPT2, and ACOX1—were also significantly decreased in the L group, consistent with impaired FAO flux in the kidneys during sepsis. Importantly, the FF group significantly reinstated the expression of all the proteins mentioned above. A significant increase in the ratio of P-AMPK to total AMPK was observed, suggesting the reactivation of AMPK signaling. Upregulation of PGC1α and PPARα further suggested enhanced transcriptional control of FAO programs. Similarly, the FF group showed a notable elevation in CPT1A, CPT2, and ACOX1 expression compared to the L group, indicating restoration of the FAO-related enzymatic system (Fig. 5). IF analysis further supported these results, showing that FF restored P-AMPK, PGC1α, PPARα, CPT1A, CPT2, and ACOX1 expression and localization in renal TECs, which were markedly reduced in the L group (Fig. 6). These spatial observations visually confirm KD-mediated reactivation of the AMPK–PGC1α–PPARα axis and its downstream FAO components in kidney tissue.

Fig. 5
figure 5

Protein expression of the AMPK–PGC1α–PPARα signaling pathway and FAO-related enzymes in kidney tissues from the C, L, and FF groups. Representative Western blot bands and quantitative analysis of P-AMPK, total AMPK, PGC1α, PPARα, CPT1A, CPT2, and ACOX1 are shown. The L group exhibited significantly decreased P-AMPK/AMPK ratio, PGC1α and PPARα levels, and expression of CPT1A, CPT2, and ACOX1, indicating suppression of AMPK signaling and impaired FAO. Fenofibrate treatment markedly restored the P-AMPK/AMPK ratio and upregulated PGC1α and PPARα, along with recovery of CPT1A, CPT2, and ACOX1 levels, suggesting reactivation of the FAO regulatory network

Fig. 6
figure 6

(A–G) Representative immunofluorescence images showing AMPK, P-AMPK, PGC1α, PPARα, CPT1A, CPT2, and ACOX1 expression and localization in renal tissues from the C, L, and FF groups (×400). Nuclei were counterstained with DAPI (blue). The L group displayed markedly reduced fluorescence for P-AMPK, PGC1α, PPARα, and key FAO-related enzymes (CPT1A, CPT2, ACOX1), reflecting suppressed AMPK activation and impaired FAO in SA-AKI. FF treatment restored these signals, predominantly in TECs. (H) semi-quantitative analysis of fluorescence intensity from six images per group (three sections × two cortical fields), normalized to DAPI-positive nuclei. The L group showed significant decreases in P-AMPK, PGC1α, PPARα, CPT1A, CPT2, and ACOX1 compared with controls, whereas FF intervention significantly increased their expression, confirming reactivation of the AMPK–PGC1α–PPARα axis and restoration of FAO-related proteins in kidney tissues

A schematic illustration summarizing the proposed mechanism and key findings is presented in the Fig. 7.

Fig. 7
figure 7

Schematic illustration of the proposed mechanism. LPS induces SA-AKI, characterized by tubular injury, inflammation, impaired FAO, mitochondrial dysfunction, and ATP depletion. Fenofibrate treatment restores FAO and mitochondrial function via activation of the AMPK–PGC1α–PPARα pathway, replenishes ATP levels, reduces lipid accumulation and inflammatory cytokine production, and ultimately alleviates renal injury

Discussion

This study demonstrated that fenofibrate markedly attenuates kidney injury in an LPS-induced SA-AKI model. Histological evaluation revealed substantial preservation of tubular architecture and reduced interstitial inflammation in fenofibrate-treated mice, in contrast to the severe tubular damage observed in untreated septic counterparts. Functionally, fenofibrate significantly improved renal performance, as evidenced by lower Scr and BUN levels, along with reduced renal expression of injury biomarkers KIM-1 and NGAL. In parallel, fenofibrate suppressed the production of pro-inflammatory cytokines. From a metabolic perspective, it restored renal ATP content and alleviated lipid accumulation—including triglycerides, glycerol, and NEFA—indicating correction of metabolic imbalance. Moreover, fenofibrate enhanced mitochondrial function, reflected by increased citrate synthase activity and improved FAO capacity. At the molecular level, fenofibrate upregulated FAO-regulatory genes and reactivated the AMPK–PGC1α–PPARα signaling pathway, as confirmed by Western blot and immunofluorescence analyses. These findings highlight fenofibrate’s dual ability to mitigate SA-AKI by restoring renal energy metabolism and suppressing inflammatory injury.

Our findings support growing evidence that metabolic reprogramming is central to SA-AKI pathophysiology [7, 8, 10]. Sepsis suppresses mitochondrial FAO in TECs through downregulation of PPARα and its transcriptional program, leading to decreased expression of FAO enzymes, impaired ATP production, and lipid accumulation [7, 8, 10]. This lipotoxicity drives mitochondrial damage, tubular dysfunction, and inflammation [9, 13]. Experimental studies show that PPARα deficiency exacerbates kidney injury in sepsis, while clinical transcriptomic analyses link repression of PPARα signaling to increased risk of severe AKI [9]. Thus, sepsis induces a maladaptive metabolic program in the kidney: the acute switch to glycolysis may initially be tolerogenic under inflammatory stress, but persistent suppression of FAO deprives tubular cells of efficient ATP generation and leads to lipid accumulation, tubular cell dysfunction, and injury [10]. This pathophysiological process is increasingly acknowledged as a key factor in the severity of SA-AKI and its progression to CKD [14].

Fenofibrate, a widely prescribed PPARα agonist, exerts pleiotropic effects on lipid metabolism and inflammation [15, 16]. By enhancing the transcription of FAO enzymes and fatty acid transport proteins, it promotes FAO activity and limits lipid accumulation in renal cells [17, 18]. In hyperlipidemic and diabetic models, fenofibrate alleviates renal lipotoxicity and activates the AMPK–PGC1α pathway to restore oxidative metabolism [19]. Fenofibrate’s anti-inflammatory actions are well documented in multiple organ systems [16, 20,21,22]. Its anti-inflammatory actions are mediated largely through suppression of nuclear factor kappa-light-chain-enhancer of activated B cells (NF-κB) signaling, thereby reducing cytokine production in endothelial, macrophage, and kidney cells [11, 21, 23, 24]. In fact, PPARα agonists blunt the cytokine surge in response to endotoxins; for example, PPARα^−/− mice exhibit exaggerated inflammatory responses to lipopolysaccharide, highlighting PPARα’s role in restraining inflammation [11]. Beyond PPARα activation, fenofibrate may also influence inflammatory cascades via lipid intermediates or off-target signaling effects [21]. In summary, fenofibrate corrects defective FAO while dampening inflammation—a dual mechanism particularly relevant to mitigating SA-AKI.

A key innovation of this work is the therapeutic targeting of metabolic reprogramming in SA-AKI—to our knowledge, this is the first study to demonstrate that restoring FAO in the kidney can effectively alleviate SA-AKI. Prior research had identified associations between PPARα-regulated metabolism and SA-AKI susceptibility, but our study provides direct evidence that augmenting FAO can causally protect against sepsis-induced organ damage [9]. This metabolic approach represents a novel strategy distinct from conventional hemodynamic or anti-inflammatory interventions for AKI. Fenofibrate’s dual action on energy metabolism and inflammation is especially advantageous in the context of sepsis, where mitochondrial dysfunction and excessive inflammation often coincide [25]. An important strength of our study is its translational potential: fenofibrate is an oral drug already approved for human use in dyslipidemia, which could facilitate rapid repurposing if efficacy in SA-AKI is confirmed. The present findings lay groundwork for exploring PPARα agonists as metabolic therapy in critical illness, and they complement emerging literature that metabolic modulation (e.g. enhancing PPARα pathways) may improve outcomes in SA-AKI. By highlighting a previously underappreciated facet of SA-AKI pathogenesis – the suppression of fatty acid metabolism – this study opens a new avenue for organ-protective therapies in sepsis that could be combined with standard antimicrobial and supportive care.

Several limitations of our study should be acknowledged. First, although our findings strongly implicate PPARα activation in fenofibrate’s mechanism, we did not perform loss-of-function experiments with PPARα-null mice or pharmacological antagonists. Global PPARα knockout animals develop profound metabolic fragility during sepsis, displaying exaggerated tubular necrosis, higher serum creatinine and markedly reduced survival, which would distort the SA-AKI phenotype and preclude a fair assessment of fenofibrate because the drug has no receptor to activate in these mice [26, 27]. Second, we deliberately avoided small-molecule PPARα inhibitors (e.g., GW6471, MK-886) because accumulating evidence shows they directly aggravate renal injury: GW6471 intensifies tubular damage, inflammation and apoptosis in ischemia-reperfusion and endotoxemic kidneys, while MK-886 reverses renoprotective PPARα signaling and exacerbates diabetic tubular injury in vivo and in vitro [28,29,30,31]. Introducing such nephrotoxic compounds would confound interpretation of fenofibrate’s benefits and raise ethical concerns. Third, fenofibrate has potential off-target effects—such as AMPK activation and direct NF-κB inhibition—that we did not dissect, so some benefit might derive from PPARα-independent pathways. Fourth, the work relied on a single murine SA-AKI model with prophylactic/early-stage dosing; human sepsis is more heterogeneous and often treated after AKI is established. Translational gaps also include species-specific pharmacokinetics and fenofibrate’s reversible Scr rise observed clinically. Finally, we did not track long-term outcomes; whether acute metabolic rescue translates into durable structural recovery remains unknown.

Conclusion

Our study demonstrates that fenofibrate significantly attenuates SA-AKI by restoring FAO and enhancing renal energy metabolism. Mechanistically, fenofibrate activates the AMPK–PGC1α–PPARα signaling pathway, which upregulates key enzymes involved in FAO, thus alleviating renal lipotoxicity and inflammation. By restoring FAO in septic kidneys, fenofibrate may offer a novel therapeutic approach for SA-AKI.