ChREBP‐β Exacerbates Renal Tubular Disorders Caused by Fructose via ATF4

Ting Fang; Xinyu Yang; Xiaoqing Deng; Jingyi Wang; Ting Li; Hang Guo; Bei Sun; Liming Chen

doi:10.1096/fj.202600490R

. 2026 Apr 13;40(8):e71774. doi: 10.1096/fj.202600490R

ChREBP‐β Exacerbates Renal Tubular Disorders Caused by Fructose via ATF4

Ting Fang ^1,², Xinyu Yang ¹, Xiaoqing Deng ¹, Jingyi Wang ¹, Ting Li ¹, Hang Guo ¹, Bei Sun ^1,^✉, Liming Chen ^1,^✉

PMCID: PMC13075461 PMID: 41973151

ABSTRACT

Excessive fructose intake is strongly associated with metabolic diseases, with the carbohydrate response element‐binding protein (ChREBP) playing a key role in its metabolism, particularly in renal tubules. However, the role of its active form, ChREBP‐β, was previously unclear. In this study, ChREBP‐β overexpression and ChREBP knockout mouse models were utilized to investigate the effects of excessive fructose intake in vivo. In addition, primary renal tubular epithelial cells from mice and human kidney‐2 (HK2) cells were applied for further validation in vitro. We found that ChREBP‐β leads to increased transcription to mediate endoplasmic reticulum stress and mitochondrial dysfunction, which ultimately impairs renal function. Our findings underscore the critical role of ChREBP‐β in fructose‐related renal disorders.

Keywords: ChREBP‐β, endoplasmic reticulum stress, fructose, mitochondrial dysfunction, renal tubular disorder

Schematic diagram of the mechanism underlying fructose‐induced renal tubular injury via ChREBP‐β‐mediated disruption of endoplasmic reticulum and mitochondrial homeostasis. This diagram illustrates that fructose induces the transcriptional activation of ChREBP‐β, which subsequently triggers endoplasmic reticulum stress, disturbs inter‐organelle calcium homeostasis, impairs mitochondrial function, and ultimately results in renal tubular epithelial cell damage and renal tubular dysfunction.

graphic file with name FSB2-40-e71774-g004.jpg

Abbreviations

4‐PBA: 4‐phenylbutyric acid
ATF4: activating transcription factor 4
ChIP: Chromatin immunoprecipitation analysis
ChREBP: Carbohydrate response element‐binding protein
DNL: de novo lipogenesis
ER: endoplasmic reticulum
ETC: electronic transport chain
GO: Gene Ontology
GTT: glucose tolerance test
HDL: high‐density lipoprotein cholesterol
ITT: insulin tolerance test
LDL: low‐density lipoprotein cholesterol
mtDNA: mitochondrial DNA
NAG: N‐acetyl‐β‐D‐glucosaminidase
OCR: oxygen consumption rate
PLA: in situ proximity ligation assay
SREBP1c: sterol regulatory element‐binding protein 1
TC: total cholesterol
TM: tunicamycin
UPR: unfolded protein response
β2‐MG: β2‐microglobulin

1. Introduction

Fructose is a monosaccharide isomerized with glucose, which plants and honey bees have pathways to make. In recent years, high‐fructose corn syrup sweeteners have been increasingly used in foods such as bread and jelly [1, 2]. As fructose intake has increased for its sweetness [3], health problems, especially metabolic diseases [4], have followed [5]. The specific roles of fructose in driving lipogenesis and pathology in the current food environment have not been fully explained. The two major mechanisms by which fructose is thought to be more lipogenic and/or toxic than glucose discussed are (1) higher affinity of ketohexokinase for fructose than glucokinase for glucose [6], (2) bypass of PFK1, which can cause hepatocytes to deplete ATP [7]. A third mechanism was recently proposed by Tiwari et al.: fructose is more lipogenic than glucose because it has a greater ability to produce glycerol‐3‐phosphate (G3P), thereby activating ChREBP and thereby driving greater transcription of lipogenic genes (as well as other targets like FGF21) [8]. Note that this mechanism invoked genes such as GPD1, GPD1L, and GK in activation of ChREBP, a key regulator of fructose metabolism [9].

ChREBP has two subtypes: α, which is highly expressed, and β, which is expressed at very low levels and acts active [10]. Unlike ChREBP‐α, ChREBP‐β lacks a low‐glucose inhibitory domain, leading to its constitutive transcriptional activity, making it the functionally active isoform of the ChREBP family under physiological and pathological conditions [11]. Regarding the activating ligands of ChREBP‐β, several molecules have been proposed in existing studies. In addition to the G3P [8], metabolites such as glucose 6‐phosphate (G6P) [12] and fructose‐2,6‐bisphosphate (F2,6BP) [13] have also been reported to act as activating ligands participating in the regulation of ChREBP‐β activity. ChREBP‐β is involved in the regulation of systemic metabolic homeostasis, and its abnormal expression is closely related to insulin resistance and B‐cell dysfunction in type 2diabetes [14].

The kidneys are one of the main organs to fructose metabolism [15], and excessive fructose intake, especially over the long‐term, leads to adverse reactions in various tissues and organs including oxidative stress, chronic inflammation, endothelial dysfunction, and autophagy [16]. Although fructose is not broken down as strongly in the kidneys as in the liver, it should not be ignored. Excessive fructose intake and rapid breakdown create metabolic stress, depleting ATP, elevating blood uric acid levels, and causing renal damage [17, 18]. The proximal straight tubule is a key site to metabolite fructose by fructokinase [19, 20]. Fructose is also metabolized in the proximal convoluted tubule, where fructokinase and the inducible aldolase B are present [21, 22]. All kidney cells require ATP for function, but ATP production varies by cell type. Proximal tubules rely on oxidative phosphorylation for efficient ATP production, which is essential for the active transport of glucose and nutrients [17]. The high energy demands of proximal tubules make aerobic respiration their primary ATP source [23]. Mitochondrial dysfunction reduces ATP production, disrupts cellular functions and structures, and contributes to renal diseases [24]. Activating transcription factor 4 (ATF4) is a stress‐induced protein, which helps cells cope with stressors like hypoxia and amino acid scarcity [25, 26]. Emerging evidence suggests ATF4 also plays a role in energy homeostasis, as it is upregulated in response to amino acid or glucose deprivation in HepG2 cells, indicating its function as a nutrient sensor [27]. This suggests that ATF4 may be important for fructose homeostasis as well.

ChREBP is a key regulator of fructose metabolism in the liver and intestine [9], but the specific role of ChREBP‐β in renal injury is still unclear. In this study, we employed mouse models with ChREBP‐β overexpression and ChREBP knockout to explore the impact of excessive fructose consumption. ChREBP‐β contributes to renal tubular reabsorption disorders by affecting mitochondrial function. Mechanistically, levels of ATF4 were significantly increased in the mutant kidneys, leading to mitochondrial dysfunction, while the removal of ATF4 mitigated these effects. Therefore, we hypothesize that ChREBP‐β is essential for regulating fructose metabolism in the kidneys and managing fructose‐induced metabolic stress through the modulation of ATF4.

2. Materials and Methods

2.1. Mouse Experiments

This study received approval from the Animal Care and Use Committee of Chu Hisen‐I Memorial Hospital at Tianjin Medical University (Approval No. DXBYY‐IACUC‐2023010), adhering to internationally recognized standards for laboratory animal care. The mice were kept in a specific pathogen‐free facility, maintained at a controlled temperature of 20°C–26°C, with a 12‐h light/dark cycle, and were fed a normal chow diet. C57BL/6J mice also received a normal chow diet, unless specified otherwise, in which case they were given a high‐fructose diet (RD22031502, Readydietech Co. Ltd., Shenzhen, P. R. China) with 34% fructose in calories. Adeno‐associated virus (AAV)2/9 with shRNA (ChREBP) targeting the ksp‐cadherin promoter was injected via the tail vein to achieve specific depletion of the ChREBP‐β gene in mouse renal tubules. AAV2/9 with shRNA (NC) targeting the ksp‐cadherin promoter was used as a control. The mice were injected with AAV2/9 via the tail vein after being fed a normal diet or a high‐fructose diet for ten weeks, and samples were collected six weeks after injection. Male mice were used for the experiments. Fru feed formulation was provided in Table S3.

2.2. Genotyping

Mice were genotyped by PCR using DNA extracted from tail biopsies and the primers listed in Table S1. DNA extraction was performed using the One Step Mouse Genotyping Kit (PD101‐01, Vazyme) according to the manufacturer's instructions.

2.3. Glucose Tolerance Tests

Following 6 h of fasting, glucose tolerance was tested by intraperitoneal injection of glucose (2 g/kg of body weight) (CAS No. 50–99‐7, Sigma). Blood glucose was measured using a glucose glucometer by tail bleeding at 0, 15, 30, 60, and 120 min after injection.

2.4. Insulin Tolerance Tests

Following 6 h of fasting, insulin tolerance was tested by intraperitoneal injection of insulin (0.5 IU/kg of body weight) (Novo Nordisk). Blood glucose was measured using a glucose glucometer by tail bleeding at 0, 15, 30, 60, and 120 min after injection.

2.5. mRNA Expression Analysis

Total RNA was extracted from kidney tissue by the TRIzol method. The concentration and purity of total RNA were measured using spectrophotometry (Nano300; Allsheng) with the ratios of A260 to A280 > 1.8. cDNA was synthesized by First Strand cDNA Synthesis SuperMix (Code #AT301‐03, TRANS, China). Quantitative PCR was performed on cDNAs by QuantStudio 3 Applied Biosystems (Thermo Fisher Scientific) with SYBR Green and specific primers for the target gene. All of the reactions were conducted in 96‐well plates in a total volume of 20 μL. All of the primers were synthesized by Tsingke Biotechnology Company (Beijing, China). In every plate, 18 s was used as the internal control.

2.6. Western Blot Analysis

Proteins were extracted from cells or pieces of kidney samples in RIPA lysis buffer supplemented with PMSF and phosphatase inhibitors. Samples were clarified by centrifugation at 12000 g for 10 min at 4°C. Protein concentrations were determined by the BCA Protein Assay Kit (Thermo Fisher Scientific). Equal amounts of proteins were resolved by SDS–PAGE and transferred to a nitrocellulose membrane. All washes were carried out with 0.1% Tween in TBS. Membranes were blocked for 1 h at room temperature in 5% nonfat dry milk in TBS containing 0.1%. Primary antibody incubation in blocking solution was performed overnight at 4°C. After incubation with goat anti‐rabbit and goat anti‐mouse secondary antibodies, the membranes were imaged.

2.7. Histological Analyses

Pieces of the kidney fixed in 4% paraformaldehyde were embedded in paraffin and stained with hematoxylin and eosin. Immunohistochemical analysis of ATF4 and Grp78 was performed on kidney sections by overnight incubation at 4°C with rabbit polyclonal anti‐ATF4 and Grp78 antibodies (1:1000) and subsequent incubation at 25°C with HRP‐polymer conjugated anti‐rabbit IgG (1:1000; ab6721) for 2 h before visualization with 0.05% DAB.

2.8. Transmission Electron Microscopy Examination

Fresh kidney tissues (1 mm3) were harvested and immediately fixed with 2.5% glutaraldehyde and postfixed with 1% osmium tetroxide. After washing, the samples were dehydrated, embedded, and polymerized. Ultrathin sections were cut and stained with uranyl acetate and lead citrate. The structure was finally visualized under a transmission electron microscope at 15000× magnification.

2.9. TUNEL Assay

A TUNEL assay kit (Beyotime Biotechnology, China) was used to detect kidney apoptosis. Briefly, kidney sections were dewaxed and then incubated with proteinase K, and cells were drilled with 0.3% Triton. Each section and cell were stained with TUNEL reaction mixture, including 5 μL of TdT and 45 μL of fluorescein‐labeled dUTP solution. After counterstaining cell nuclei with DAPI, TUNEL‐positive fluorescent images were captured, and the proportion of apoptosis was counted.

2.10. Measurement of Mitochondrial Oxygen Consumption

The analysis of mitochondrial respiration was conducted using an Oxygraph‐2 k (OROBOROS, Austria) with different inhibitors present.

2.11. In Situ Proximity Ligation Assay (PLA)

PLA was performed on paraffin‐embedded kidney sections and cells. The process involved blocking and incubation with primary antibodies, followed by treatment with PLA probes and amplification solutions. Staining with hematoxylin or DAPI was done before imaging with light (Olympus, Japan) or confocal microscopy (Carl Zeiss, Germany) at 400× magnification.

2.12. Isolation of Primary Renal Tubular Epithelial Cells

Kidneys from ChREBP‐β‐KI and WT mice (4‐ to 6‐week‐old males) were collected after euthanization and minced into pieces of approximately 1 mm3. These pieces were digested with 10 mL HBSS containing 0.1% collagenase IV for 30 min at 37°C with gentle stirring, and the supernatants were sieved through a 100 μm nylon mesh and a 40 μm nylon mesh. After centrifugation for 5 min at 800 g, the pellet was resuspended in DMEM/F12 1:1 supplemented with 10% FBS and 1% penicillin/streptomycin at 37°C and 5% CO₂. Cells were used between days 7 and 10 of culture.

2.13. Cell Culture

Human kidney‐2 (HK‐2) cells were cultured in DMEM/F12 (Gibco) supplemented with 10% FBS and 1% penicillin/streptomycin. HEK293 cells were cultured in DMEM (Gibco) supplemented with 10% FBS and 1% penicillin/streptomycin. All cells were cultured at 37°C and 5% CO₂ and treated with trypsin every 3 days for subculturing.

2.14. Ca²⁺ Measurements

Human kidney‐2 (HK2) cells were treated with 2 μM Fura‐2 AM (ab120873, Abcam) and 1 μM Rhod2‐AM (ab142780, Abcam) in a Pluronic‐F27 solution for 15 min at 37°C, followed by a wash. Time‐lapse fluorescence imaging of calcium levels was conducted at 20‐s intervals. To trigger Ca²⁺ release from ER, 1 μM ATP (11 140 965 001, Roche) was applied. Calcium levels were measured as relative fluorescence (F/F0) compared to baseline.

2.15. Flow Cytometry

An Annexin V‐FITC/PI Apoptosis Detection Kit (BestBio, China) was used to assess the apoptosis of primary renal tubular epithelial cells according to the manufacturer's instructions. Cells were collected and resuspended in binding buffer. After incubation with Annexin V‐FITC for 15 min and PI for 5 min at 4°C in the dark, the samples were analyzed by a flow cytometer (Thermo Fisher, America).

2.16. FITC‐BSA Staining

Primary renal tubular epithelial cells and HK2 cells were cultured in 24‐well plates and treated with 50 μg/mL FITC‐BSA (SF063, Solarbio, Beijing) for 3 h in the dark. Subsequently, the cells were rinsed twice with PBS. Cells in 24‐well plates were directly observed using a fluorescence microscope (Olympus, Tokyo, Japan).

2.17. Dual Luciferase Assay

The human ChREBP‐β/Mlx expression plasmid and the human ATF4 promoter plasmid (GeneCopoeia) were co‐transfected into HEK293T cells with the help of Transfection Reagent (Invigentech). Luciferase activity was measured 48 h post‐transfection using a dual‐luciferase assay kit (GeneCopoeia).

2.18. Chromatin Immunoprecipitation (ChIP) Analysis

Cells were crosslinked with 1% formaldehyde for 10 min at room temperature, and the reaction stopped using 0.5 M glycine. After digesting with micrococcal nuclease (#10011, Cell Signaling Technology) and sonication, the chromatin was incubated with an anti‐ChREBP antibody or normal rabbit IgG. This was followed by incubation with ChIP‐Grade Protein G agarose beads. The purified chromatin DNA was analyzed by PCR using gene promoter‐specific primers, with sequences provided in Table S2.

2.19. Statistical Analysis

Statistical analysis was performed using GraphPad Prism 8.0 software. Data are presented as means ± SD. Statistical significance was set at a p value of less than 0.05. Comparisons between two groups were conducted using unpaired Student's t‐tests, while comparisons among multiple groups utilized one‐way analysis of variance (ANOVA) followed by a post hoc Tukey's test.

3. Results

3.1. High Fructose Induces Insulin Resistance and Renal Tubular Injury

C57BL/6J mice were fed either a normal chow diet (Chow) or a high‐fructose diet (34% Fru) with 34% fructose in calories for 16 weeks. Mice that received 34% Fru had significantly greater body weight gain after 1 week than the mice that received Chow (Figure 1A), and high blood glucose levels were detected after 2 weeks (Figure 1B). A glucose or insulin tolerance test (GTT/ITT) revealed 34% Fru‐fed mice had low glucose clearance capacity and insulin sensitivity compared with Chow‐fed mice (Figure 1C–F). The 34% Fru‐fed mice had larger body sizes and kidney weights than the control mice (Figure S1A,B). Serum total cholesterol (TC), low‐density lipoprotein cholesterol (LDL‐C), and high‐density lipoprotein cholesterol (HDL‐C) were assessed. The results revealed higher TC, LDL‐C, and HDL‐C levels in mice that received 34% Fru than in the Chow‐fed mice (Figure S1C–E). Twenty‐four‐hour urine results revealed that urine volume and urine protein were increased in 34% Fru‐fed mice compared with Chow‐fed mice (Figure 1G). Markers of renal tubular injury (NAG and β2‐MG) were detected with an ELISA (Figure 1H–J). H&E staining revealed obvious architectural changes, such as glomerular enlargement and tubular atrophy, in the mice that received the 34% Fru diet, whereas the kidneys of control mice did not show significant abnormalities (Figures 1K and S1F). Transmission electron microscopy revealed that the continuity of the brush border was disrupted in the 34% Fru‐fed group, and the surface area of the microvilli was significantly decreased (Figure 1L).

Effects of fructose exposure on the kidneys. (A,B) Weight and blood glucose of mice after different treatments for 16 weeks. (C,D) GTT, (E,F) ITT of mice. (G–J) Urine volume, urinary β2‐MG, NAG, and protein of mice after different treatments for 16 weeks. (K) Representative images of HE‐stained kidney sections. Scale bars = 100 μm. (L) Representative TEM images showing the brush border area in the kidneys of mice. Scale bars = 1 μm. (M) mRNA expression of ChREBP‐β was measured by RT–PCR. n = 6; all data are expressed as mean ± SD. *p < 0.05.

3.2. Generation of Renal Tubular‐Specific ChREBP‐β‐Overexpressing Mice

The mRNA expression of ChREBP‐β was measured via real‐time PCR, and expression increased significantly in the 34% Fru‐fed group (Figure 1M). To explore the in vivo function of renal tubular ChREBP‐β, we generate renal tubular‐specific ChREBP‐β‐overexpressing (ChREBP‐β‐KI) mice by crossing Rosa‐ChREBP‐β knock‐in mice with Cdh16‐Cre mice (Figure 2A,B). Overexpression efficiency was confirmed at the mRNA level via RT–PCR (Figure 2C). A GTT revealed a lower glucose clearance capacity in the ChREBP‐β‐KI mice than in the control mice (Figure 2D,E). H&E staining and TUNEL staining revealed the same pathological changes in the ChREBP‐β‐KI mice (Figures 2F and S1G,H). Transmission electron microscopy revealed that the continuity of the brush border was disrupted in the ChREBP‐β‐KI group, and the surface area of the microvilli was significantly decreased compared with that in the control group (Figure 2G).

Effects of ChREBP‐β renal tubular‐specific overexpression on the kidney of mice. (A) Schematic demonstration. (B) Representative demonstration for PCR‐based genotyping. (C) Real time PCR analysis for ChREBP‐β overexpression efficiency in the kidney. (D,E) GTT of mice and the area under the curve statistics. (F) Representative images of HE and Tunel‐stained kidney sections. Scale bars = 100 μm. (G) Representative TEM images showing the brush border area in the kidneys of mice. Scale bars = 1 μm. n = 6; all data are expressed as mean ± SD. *p < 0.05.

3.3. High Fructose or Renal Tubular‐Specific ChREBP‐β Overexpression Induces Mitochondrial Dysfunction

Transcriptomic and proteomic data were compared between the kidneys of ChREBP‐β‐KI mice and control mice. Gene Ontology (GO) enrichment analysis of the transcriptomics revealed that transmembrane transport and the brush border were the main biological processes and cellular components. Molecular function was associated with oxygen supply (Figure S2A–C). The proteomics results revealed that the energy supply for the tricarboxylic acid cycle was reduced and that mitochondrial function was impaired (Figure S2D,E). The oxygen consumption rate (OCR) in homogenized mouse kidneys was lower in the 34% Fru‐fed and ChREBP‐β‐KI mice than in Chow‐fed and control mice, as determined by O2k detection (Figure 3A). Transmission electron microscopy revealed greater mitochondrial size in the kidneys of 34% Fru‐fed and ChREBP‐β‐KI mice than in the kidneys of control mice; in addition, mitochondrial vacuolization with crista loss and expansion in the endoplasmic reticulum (ER) were observed (Figure 3B). As mitochondrial size increased, mitochondrial dynamics (fission and fusion) were detected via Western blotting. The results revealed increased fusion in 34% Fru‐fed mice (Figure S3A). Reactive oxygen species (ROS) staining of kidneys revealed increased oxidative stress in 34% Fru‐fed and ChREBP‐β‐KI mice (Figures 3C and S3B). More importantly, MitoSOX staining also revealed increased production of mitochondrial ROS (Figures 3D and S3C). To investigate the mechanism behind impaired mitochondrial function in ChREBP‐β‐KI mice, we analyzed the expression of relevant genes in the kidneys. We found that mitochondrial gene expression was lower in the ChREBP‐β‐KI mice compared to the control mice (Figure 3E). Mitochondrial respiration relies on the activity of the electron transport chain (ETC), composed of five protein complexes (C‐I to C‐V). We assessed whether long‐term fructose exposure or ChREBP‐β overexpression altered the composition of the ETC. Notably, there was a reduction in several respiratory chain components, including CIII‐UQCRC2, CV‐ATP5A, CII‐SDHB, and CI‐NDUFB8, in the kidneys of ChREBP‐β‐KI mice compared to controls, with the C‐II subunit SDHB showing the most significant decrease, while CIV‐MTCO1 remained unchanged (Figure 3F,G). These findings indicate that ChREBP‐β overexpression may impair mitochondrial respiration and contribute to mitochondrial stress.

ChREBP‐β renal tubular‐specific overexpression impaired mitochondrial function in the kidney of mice. (A) Fresh kidneys were subjected to analysis of oxygen consumption in the presence of specific inhibitors. After stabilization of basal respiration, rotenone was added to obtain nonmitochondrial oxygen consumption, followed by titration of UCCP to maximum oxygen flux. Finally, the ATP‐synthase inhibitor oligomycin was added to obtain a measure of LEAK respiration. (B) Representative TEM images of mitochondria in the kidneys of mice. Scale bars = 1 μm. (C) Representative images of ROS‐stained kidney sections. Scale bars = 100 μm. (D) Representative images of MitoSOX‐stained kidney sections. Scale bars = 50 μm. (E) mRNA levels of mitochondrial respiration‐related genes. (F) Western blot analysis of the kidney mitochondrial respiratory complex. (G) Quantitative analysis of the indicated protein levels in (F). n = 6; all data are expressed as the mean ± SD. *p < 0.05.

3.4. High Fructose‐ Or Renal Tubular‐Specific ChREBP‐β Overexpression Induces Kidney ER Stress and Calcium Homeostasis Disorders

Transmission electron microscopy analysis revealed expansion in the ER (Figures 3B and 4A), and we detected the degree of accumulation of unfolded proteins via TPE‐MI probes [28]. The results showed increased fluorescence intensity in the kidneys of 34% Fru‐fed and ChREBP‐β‐KI mice, which indicated the accumulation of more unfolded proteins (Figures 4B and S3D). Moreover, the levels of expression of markers of ER stress and one of the classic pathways of the unfolded protein response (UPR), Grp78, p‐PERK/PERK, p‐ elf2α/elf2α, ATF4, and CHOP, were detected. Consistently, expression was greater in the kidneys of 34% Fru‐fed and ChREBP‐β‐KI mice (Figure 4C,D). The same changes occurred in the livers of 34% Fru‐fed mice (Figure S3E). The other two classic pathways demonstrated no differences (Figure S3A). Immunohistochemical results for the marker proteins Grp78 and ATF4 also suggested that the UPR was enhanced (Figures 4E and S3F,G). Since close contact occurs between mitochondria and the ER (e.g., via Ca²⁺ transport), VDAC1 and IP3R1 expression levels were determined. VDAC1 was found to be enriched in the mitochondrial fraction, while IP3R1 was predominantly located in the ER. The expression levels were similar between the control and ChREBP‐β‐KI mice (Figures 4F and S3H). Additionally, in situ PLA was conducted to examine the interaction between VDAC1 and IP3R1 and to visualize calcium transport. Interestingly, lower levels of VDAC1/IP3R1 complexes were detected in the kidneys of 34% Fru‐fed and ChREBP‐β‐KI mice (Figures 4G and S3I). Furthermore, the calcium concentration was measured via a confocal live‐cell workstation, with mitochondrial calcium concentration ([Ca²⁺]_m) assessed via Rhod‐2 AM, and cytosolic ([Ca²⁺]_c) using Fura‐2 AM. Both cytosolic and mitochondrial calcium levels were monitored after ATP‐induced calcium release from the ER. Compared with the controls (Movie S1), mitochondrial Ca²⁺ uptake was less stimulated by fructose, and the cytosolic Ca²⁺ concentration was greater (Figures 4H and S3J, Movie S2).

ER stress, apoptosis, and calcium homeostasis disorder were exacerbated in 34% Fru and ChREBP‐β‐KI kidney. (A) Representative TEM images of the endoplasmic reticulum area in the kidneys of mice. Scale bars = 1 μm. (B) Representative images of TPE‐MI‐stained kidney sections. Scale bars = 50 μm. (C) Western blot bands and quantitative analysis of Grp78, p‐PERK, PERK, p‐eIF2α, eIF2α, ATF4, CHOP, Bax, and Bcl2 in kidney lysates. (D) Western blot bands and quantitative analysis of Grp78, p‐PERK, PERK, p‐eIF2α, eIF2α, ATF4, CHOP, Bax, and Bcl2 in kidney lysates. (E) Immunohistochemical staining of Grp78 and ATF4 in kidney sections. Scale bars = 100 μm. (F) Western blot bands of IP3R1 and VDAC1 in kidney lysates. (G) Representative in situ PLA images in the kidneys. Scale bars = 100 μm. (H) Cells were analyzed for calcium content in response to ATP stimulation. After loading the mitochondrial and cytoplasmic calcium probes for stabilization, calcium flow was stimulated using ATP, and representative images were acquired by prolonged shooting. n = 6; all data are expressed as the mean ± SD. *p < 0.05.

3.5. ChREBP Knockout Relieves Fructose‐Mediated Renal Tubular Injury

To verify the role of ChREBP‐β in fructose‐induced tubular injury, AAV2/9 with shRNA (ChREBP) with the ksp‐cadherin promoter was injected via the tail vein to achieve specific depletion of the ChREBP‐β gene in mouse renal tubules. The knockdown efficiency was confirmed at the mRNA level with RT–PCR (Figure 5A). GTTs were performed on the mice at the endpoint of the experiment. The 34% Fru‐fed‐AAV‐ChREBP mice had greater glucose clearance than the 34% Fru‐fed‐AAV‐NC mice did (Figure 5B,C). These findings suggest that fructose‐induced impairment of glucose tolerance can be alleviated after renal tubular ChREBP knockout. Mouse urine was collected for an ELISA, which showed that the albumin‐to‐creatinine ratio (ACR) and the renal tubular injury indicators NAG and β2‐MG were elevated in fructose‐fed mice compared with Chow‐fed‐AAV‐NC mice but decreased in ChREBP‐knockdown mice compared with 34% Fru‐fed‐AAV‐NC mice (Figure 5D–F). H&E staining revealed that structural changes such as glomerular enlargement and tubular atrophy that occurred in the kidneys of 34% Fru‐fed‐AAV‐NC mice were not significantly shown in the kidneys of 34% Fru‐fed‐AAV‐ChREBP mice (Figures 5G and S3K,L). ROS staining and MitoSOX staining revealed enhanced oxidative stress and enhanced production of mitochondrial ROS in the kidneys of 34% Fru‐fed‐AAV‐NC mice, but these effects were alleviated after ChREBP knockdown in the renal tubules of 34% Fru‐fed‐AAV‐ChREBP mice (Figures 5H,I and S3M,N). TPE‐MI staining revealed that the fructose‐induced accumulation of unfolded proteins in the renal tubular ER was also alleviated by ChREBP knockdown (Figures 5J and S3O). Compared with Chow‐fed‐AAV‐NC mice, the ETC composition was reduced, and classic UPR pathways (including ATF4 and apoptosis) were enhanced; however, after ChREBP knockdown, these effects were alleviated in the kidneys of 34% Fru‐fed‐AAV‐ChREBP mice compared with 34% Fru‐fed‐AAV‐NC mice (Figure 5K,L). The above data show that ChREBP‐β plays a dominant role in fructose‐induced renal tubular ER stress and mitochondrial dysfunction.

Fructose‐induced kidney injury reversed by ChREBP knockout. (A) Real time PCR analysis for ChREBP‐β knockout efficiency in the kidney. (B,C) GTT of mice and area under the curve statistics. (D–F) Urine ACR, urinary β2‐MG, and NAG of mice after different treatments. (G) Representative images of HE‐stained kidney sections. Scale bars = 100 μm. (H) Representative images of ROS‐stained kidney sections. Scale bars = 100 μm. (I) Representative images of MitoSOX‐stained kidney sections. Scale bars = 50 μm. (J) Representative images of TPE‐MI‐stained kidney sections. Scale bars = 50 μm. (K,L) Western blot and quantitative analysis of the kidney mitochondrial respiratory complex and Grp78, p‐PERK, PERK, p‐eIF2α, eIF2α, ATF4, CHOP, Bax, and Bcl2. n = 6; all data are expressed as mean ± SD. *p < 0.05.

3.6. ChREBP‐β Induces Renal Tubular ER Stress, Mitochondrial Dysfunction, and Apoptosis

To understand the mechanism by which renal tubular‐specific ChREBP‐β overexpression causes the above phenomenon, we isolated and cultured primary renal tubular epithelial cells from mice. The isolation and culture process and identification results are shown in Figure S4A,B. The reduced reabsorption of the cells from ChREBP‐β‐KI mice was evident from the diminished fluorescence intensity of the FITC‐BSA staining compared with that of the control (Figures 6A and S4C). The ratios of total NAD⁺ to total NADH and the number of mitochondrial DNA (mtDNA) copies were both lower in the cells of the ChREBP‐β‐KI mice (Figure 6B,C). ETC composition was reduced, and the classic UPR pathways, which include ATF4 and apoptosis, were enhanced in cells from ChREBP‐β‐KI mice (Figure 6D–G). The flow cytometry results also revealed that more apoptotic cells appeared in the cells of the ChREBP‐β‐KI mice (Figure 6H). The above results were consistent with in vivo results. To alleviate ER stress, we treated cells from ChREBP‐β‐KI mice with 1 mM 4‐phenylbutyric acid (4‐PBA) for 24 h. Cells from control mice treated with tunicamycin (TM) 0.5 μg/mL for 24 h were used as a positive control. Western blot analyses demonstrated that 1 mM 4‐PBA alleviated the UPR and mitochondrial stress. In addition, mitochondrial respiration (ETC composition) improved after intervention (Figure 6I,J), and ROS staining showed that oxidative stress was alleviated (Figures 6K and S4D). VDAC1/IP3R1 complex formation increased, as shown by in situ PLA, after intervention (Figures 6L and S4E). These results suggest that ER stress plays a leading role in the process of progression.

Relieving endoplasmic reticulum stress improves mitochondrial function and apoptosis in the renal tubular epithelial cells. (A) Representative images of FITC‐BSA staining cells. Scale bars = 50 μm. (B) Measurement of mtDNA copy number normalized to the nuclear gene. (C) Measurement of the NAD⁺/NADH ratio in cells. (D) Western blot analysis of the mitochondrial respiratory complex. (E) Quantitative analysis of the indicated protein levels in (D). (F) Western blot bands of Grp78, ATF4, CHOP, IP3R1, VDAC1, Bax, and Bcl2. (G) Quantitative analysis of the indicated protein levels in (F). (H) Representative images of flow cytometry in renal tubular epithelial cells. (I) Western blot bands of p‐eIF2α, eIF2α, ATF4, CHOP, Bax, Bcl2, and the mitochondrial respiratory complex in renal tubular epithelial cells after different interventions. (J) Quantitative analysis of the indicated protein levels in I. (K) Representative images of ROS staining in renal tubular epithelial cells. Scale bars = 50 μm. (L) Representative in situ PLA images in renal tubular epithelial cells after different interventions. Scale bars = 50 μm. n = 3; all data are expressed as the mean ± SD. *p < 0.05.

3.7. ChREBP Knockout Relieves Fructose‐Mediated ER Stress to Improve Mitochondrial Dysfunction in HK2 Cells

In addition to our research with mice, we verified our findings in the human HK2 cell line. ChREBP‐β and MLX plasmids were cotransfected into HK2 cells, and HK2 cells transfected with the control plasmid were used as a negative control. Western blot analyses revealed that the ETC composition was reduced and that classic UPR pathways, including ATF4 and apoptosis, were enhanced (Figure 7A–D). Compared with that of the control, reduced reabsorption by the cells was evident from the diminished fluorescence intensity of FITC‐BSA staining (Figures 7E and S4F). To define the role of ChREBP in the process of progression in vitro, we transfected HK2 cells treated with 5 mM fructose for 72 h with si‐ChREBP. We found that ChREBP knockdown alleviated the fructose‐mediated UPR and mitochondrial stress. In addition, mitochondrial respiration (ETC composition) improved after ChREBP knockdown (Figure 7F–I). More importantly, the fructose‐induced decrease in reabsorption capacity was reversed after ChREBP knockout (Figures 7J and S4G). To alleviate ER stress, we treated cells transfected with ChREBP‐β and MLX plasmids with 0.5 mM 4‐PBA for 24 h. Cells transfected with the control plasmid and treated with 0.5 μg/mL TM for 24 h were used as positive controls. Western blot analyses demonstrated that 1 mM 4‐PBA alleviated the UPR and mitochondrial stress. In addition, mitochondrial respiration (ETC composition) improved after intervention (Figure 7K,L). The results were consistent with those of the primary cells. The optimal concentrations of fructose, 4‐PBA, and TM were explored via RT–PCR and Western blotting (Figure S5). These results suggest that relieving ER stress improves mitochondrial function.

ChREBP‐β regulated endoplasmic reticulum stress, mitochondrial function, and reabsorption capacity in HK2 cells. (A) Western blot analysis of the mitochondrial respiratory complex. (B) Quantitative analysis of the indicated protein levels in A. (C) Western blot bands of Grp78, p‐eIF2α, eIF2α, ATF4, CHOP, Bax, and Bcl2. (D) Quantitative analysis of the indicated protein levels in (C). (E) Representative images of FITC‐BSA staining. Scale bars = 50 μm. (F) Western blot analysis of the mitochondrial respiratory complex. (G) Quantitative analysis of the indicated protein levels in (F). (H) Western blot bands of Grp78, p‐PERK, PERK, p‐eIF2α, eIF2α, ATF4, CHOP, Bax, and Bcl2. (I) Quantitative analysis of the indicated protein levels in (H). (J) Representative images of FITC‐BSA staining. Scale bars = 50 μm. (K) Western blot bands of Grp78, p‐eIF2α, eIF2α, ATF4, CHOP, Bax, Bcl2, and the mitochondrial respiratory complex. (L) Quantitative analysis of the indicated protein levels in (K). n = 3; all data are expressed as the mean ± SD. *p < 0.05.

3.8. ChREBP‐β Bind to ATF4 Promoter and Increase Its Transcription

The results presented above demonstrate a connection between ChREBP‐β and ER stress. We knocked out ChREBP in HEK293 cells and observed a significant reduction in ATF4 expression (Figure 8A,B). To further investigate the role of ATF4 in regulating ER stress in tubule cells exposed to fructose, we treated HK2 cells with 5 mM fructose for 72 h and transfected them with si‐ATF4. Our findings revealed that the knockdown of ATF4 diminished the fructose‐induced increases in proteins associated with the unfolded protein response (UPR) (Figure 8C,D). These results suggest that ATF4 plays a crucial role in this process.

ChREBP‐β binds to the ATF4 promoter and activates its transcriptional expression. (A) Western blot analysis of ChREBP and ATF4 in HEK293 cells. (B) Quantitative analysis of the indicated protein levels in (B). (C) Western blot bands of Grp78, p‐eIF2α, eIF2α, ATF4, and CHOP in HK2 cells. (D) Quantitative analysis of the indicated protein levels in (C). (E) Binding of ChREBP to promoters of different factors as predicted by the JASPAR database. (F) Relative luciferase activity of the ATF4 promoter in response to ChREBP‐β overexpression. (G) Values represent the relative increase in real‐time PCR signals for ChIP with anti‐ChREBP antibodies compared to those for ChIP with control IgG. n = 3; all data are expressed as the mean ± SD. *p < 0.05.

We evaluated the binding of ChREBP to the promoters of ATF4, Grp78, eIF2α, and CHOP using the JASPAR database. The score values indicated that ChREBP had the highest likelihood of binding to the ATF4 promoter (Figure 8E). Subsequently, we confirmed that ChREBP‐β could directly bind to the ATF4 promoter region. Overexpression of ChREBP‐β significantly increased the protein level of ATF4 (Figure 4D,E), confirming that ChREBP‐β activates ATF4 expression. Additionally, knockdown of ChREBP resulted in a significant decrease in ATF4 protein levels (Figure 7H). ChREBP‐β/Mlx markedly enhanced the transcriptional activity of the ATF4 promoter (Figure 8F). We then investigated the physical interactions between ChREBP‐β and the ATF4 promoter using a ChIP assay on HEK293 cells. The DNA from the immunoprecipitates was amplified via PCR with primers located in the promoter region near the putative binding site for ChREBP‐β. Our findings revealed that ChREBP‐β physically bound to this region (Figure 8G). These results confirmed that ChREBP‐β regulates ATF4 transcription and thereby impacts mitochondrial respiration.

4. Discussion

Fructose metabolism has been shown to occur in the liver [29, 30], but few studies of the kidneys have been conducted. ChREBP‐β is a key factor in fructose metabolism. In this study, we established an important role for ChREBP‐β in fructose metabolism in the kidneys. First, the accumulation of fructose led to renal tubular injury in mice, which was manifested by increases in urinary protein, NAG, and β2‐MG after long‐term exposure to a 34% fructose diet. Second, conditional overexpression of ChREBP‐β substantially impaired renal mitochondria, especially oxidative phosphorylation (mitochondrial respiration), and increased mitochondrial and ER stress, which was mediated by increased expression of ATF4. Third, ChREBP‐β ablation reversed the impairments in renal mitochondria and the ER that were induced by fructose due to a reduction in ATF4. Fourth, we determined that ATF4 was regulated by ChREBP‐β, which plays a previously unrecognized role in renal function.

The current study clarified the role of mitochondria and ER stress in renal injury. Mitochondria provide the energy and transmission electron microscopy revealed larger mitochondria in the kidneys after fructose intake and ChREBP‐β overexpression, which indicated that fructose and ChREBP‐β could result in mitochondrial dysfunction. These results also demonstrated an imbalance in mitochondrial dynamics (fission and fusion). The expression of MFN2 was increased, whereas that of DRP1 was not, which led to increased mitochondrial fusion. Samir Softic's work has provided valuable insights into the effects of fructose on mitochondrial dynamics, particularly fission and fusion processes, and suggests that fructose consumption may promote mitochondrial fission, a process in which mitochondria divide into smaller units [31]. This can lead to an increase in the number of fragmented mitochondria, which may affect cellular energy metabolism [32]. Although we found that fructose intake resulted in increased mitochondrial fusion, we did not investigate it further in this study. Mitochondria are critical for ATP synthesis through oxidative phosphorylation. Dysfunction can result from factors such as oxidative stress, ischemia, or toxins, which lead to reduced ATP levels. The OCR in homogenized mouse kidneys was lower in 34% Fru‐fed and ChREBP‐β‐KI mice than in Chow‐fed and control mice, as determined by O2k detection. A decrease in ATP can impair kidney function and contribute to cellular injury. Over time, mitochondrial dysfunction can lead to impaired mitochondrial biogenesis and increased mitophagy (the process of removing damaged mitochondria). This attrition can exacerbate energy deficits in renal cells and contribute to progressive renal disease [33]. In summary, mitochondrial dysfunction is closely linked to reduced ATP levels in renal cells and can lead to mitochondrial loss over time, further compromising kidney function. Transmission electron microscopy analysis revealed expansion in the endoplasmic reticulum. Fructose increases the UPR pathway, which includes PKR‐like kinase (PERK) and ATF4 [34]. PERK and ATF4 hyperactivation can increase the expression of CHOP, which promotes cell death [35, 36].

We established the role of ChREBP‐β in fructose‐induced ER and mitochondrial stress in the kidney. As the main transcription factor, ChREBP‐β responds to carbohydrate stimulation [37]. Our results demonstrated that ChREBP‐β governs ER and mitochondrial functions. Excessive fructose metabolic demand mediates ER stress via ChREBP‐β. In response to this pressure, the ER initiated the UPR. The knockdown of ChREBP clearly alleviated this metabolic pressure by decreasing the expression of UPR‐related proteins. Moreover, mitochondrial function, especially mitochondrial respiration, was impaired when the expression of ChREBP‐β increased and was rescued following ChREBP knockdown.

This study examines the interconnected roles of mitochondria and ER stress [38], which can lead to mitochondrial dysfunction through excess calcium influx [39], resulting in increased mitochondrial fragmentation and reduced membrane potential. Fructose feeding and specific experiments showed decreased VDAC1/IP3R1 complexes in kidneys and lower mitochondrial Ca²⁺ uptake, despite higher cytosolic Ca²⁺ levels. An ER stress inhibitor improved mitochondrial respiration, suggesting unresolved ER stress contributes to mitochondrial dysfunction and cell death [40, 41]. This process involves the release of pro‐apoptotic factors, triggering apoptosis via effector caspases and regulation by the BCL‐2 family of proteins [42]. Overall, ER stress significantly influences fructose‐related metabolic effects by exacerbating mitochondrial dysfunction.

Fructose has significant effects on metabolism, particularly in the context of diabetes and kidney disease [4]. Fructose is metabolized primarily in the liver, where it can cause increased fat production [43] and insulin resistance [44]. High fructose intake can worsen insulin sensitivity and make blood sugar more difficult to control for individuals with diabetes [45]. Diabetes can lead to the type of kidney damage known as diabetic kidney disease (DKD). High fructose consumption may exacerbate this condition. Fructose can promote inflammation and oxidative stress, both of which are harmful to kidney function [46]. Fructose metabolism produces uric acid [47], which can lead to hyperuricemia and is associated with kidney damage [48]. Furthermore, DKD involves complex cellular processes, including ER stress and mitochondrial dysfunction [49]. Elevated blood sugar levels can lead to increased protein synthesis, which overwhelms the ER. ER stress activates inflammatory pathways and triggers programmed cell death in renal cells [50]. Our results revealed that fructose exposure leads to ER stress and mitochondrial dysfunction, which may aggravate DKD. Individuals with DKD should limit fructose intake.

Sodium–glucose cotransporter 2 (SGLT2) inhibitors are used primarily to manage type 2 diabetes [51]. SGLT2 is a transporter protein located in the proximal convoluted tubule of the nephron [52]. The key pharmacological function of SGLT2 inhibitors is to block this transporter protein, which reduces glucose reabsorption and promotes glucose excretion in the urine [53]. By preventing glucose reabsorption, SGLT2 inhibitors lower blood glucose levels, which benefits patients with hyperglycemia. In this study, we found that fructose overintake resulted in impaired renal tubular reabsorption, which appeared to be similar to the effect of an SGLT2 inhibitor reducing glucose reabsorption. However, the fructose‐induced reabsorption impairment was not specific to glucose. The focus of this study was on protein reabsorption. Fructose may diminish the efficacy of SGLT2 inhibitors for the disruption of insulin sensitivity. Fructose intake tends to increase body weight and adiposity, and SGLT2 inhibitors typically promote weight loss [54], which is beneficial for individuals consuming high‐fructose diets. While SGLT2 inhibitors can effectively control blood glucose levels, it is also critical to consider dietary factors such as fructose intake.

Author Contributions

Ting Fang: Conceptualization and methodology. Xinyu Yang: Methodology and software. Xiaoqing Deng: Validation. Jingyi Wang: Writing – original draft. Ting Li: Writing – original draft. Hang Guo: Writing – original draft. Bei Sun: Writing – Reviewing and editing. Liming Chen: Writing – reviewing and editing and funding acquisition.

Funding

This work was supported by MOST | National Natural Science Foundation of China (NSFC) (Grants 82370842 and 82305007), Tianjin Key Medical Discipline Construction Project (Grant TJYXZDXK‐3‐007B), Research Project of the Affiliated Hospital of Zhejiang Chinese University (Grant 2025FSYYZQ02), the Natural Science Foundation of Tianjin (Grant 23JCZDJC00640).

Ethics Statement

This study received approval from the Animal Care and Use Committee of Chu Hisen‐I Memorial Hospital at Tianjin Medical University (Approval No. DXBYY‐IACUC‐2023010).

Conflicts of Interest

The authors declare no conflicts of interest.

Supporting information

Figure S1: Fructose exposure led to increased kidney weight and cholesterol. (A) Representative images of the whole body of mice in different groups. (B) Quantitative analysis of kidney weight, n = 5. (C‐E) Serum TC, LDL‐C and HDL‐C of mice in the fasted state in different groups. (F) Quantitative analysis of the HE‐stained kidney sections in Figure 1K. (G‐H) Quantitative analysis of the HE and TUNEL‐stained kidney sections in Figure 2F. Data are presented as the mean ± SD, n = 6 per group.

Figure S2: Transcriptomics and proteomics showed mitochondrial dysfunction and brush border injury in the kidneys of ChREBP‐β‐KI mice. (A–C) GO enrichment analysis of transcriptomics. (D) Subcellular organelle analysis of proteomics. (E) Heatmap analysis of mitochondria‐related proteins in proteomics.

Figure S3: Mitochondrial dynamics were imbalanced in the kidney and different in the liver. (A) Western blot analysis of ATF6, XBP1, P62, MFN2, DRP1^Ser616, DRP1^Ser637 and t‐DRP1 in kidney lysates. (B) Quantitative analysis of the ROS‐stained kidney sections in Figure 3C. (C) Quantitative analysis of the MitoSOX‐stained kidney sections in Figure 3D. (D) Quantitative analysis of the TPE‐MI‐stained kidney sections in Figure 4B. (E) Western blot analysis of CHREBP, Grp78, p‐eIF2α, eIF2α, ATF4, CHOP, Bax and Bcl2 in liver lysates. (F‐G) Quantitative analysis of immunohistochemical results in Figure 4E. (H) Quantitative analysis of the indicated protein levels in Figure 4F. (I) Quantitative analysis of the PLA‐stained kidney sections in Figure 4G. (J) Quantitative analysis of the Ca²⁺ levels in Figure 4H. (K‐L) Quantitative analysis of the HE ‐stained kidney sections in Figure 5G. (M‐O) Quantitative analysis of the ROS, MitoSOX and TPE‐MI‐stained kidney sections in Figure 5H–J.

Figure S4: Isolation, culture and identification of primary renal tubular epithelial cells. (A) Representative images of primary renal tubular epithelial cells on different days. (B) Immunofluorescence staining of CK18 and AQP1 in primary renal tubular epithelial cells. The scale bar represents 50 μm. (C) Quantitative analysis of the FITC‐BSA‐stained cells in Figure 6A. (D) Quantitative analysis of the ROS‐stained cells in Figure 6K. (E) Quantitative analysis of the PLA‐stained cells in Figure 6L. (F‐G) Quantitative analysis of the FITC‐BSA‐stained cells in Figure 7E,J.

Figure S5: Selection of different intervention concentrations for different cells. (A) Western blot bands and quantitative analysis of Grp78 in the primary renal tubular epithelial cells of ChREBP‐β‐KI kidneys stimulated by different concentrations of 4‐PBA. (B) Western blot bands and quantitative analysis of Grp78 in the primary renal tubular epithelial cells of the control kidney stimulated by different concentrations of TM. (C) mRNA levels of ChREBP‐β in HK2 cells stimulated by different concentrations of fructose. (D) mRNA levels of ChREBP‐β in HK2 cells stimulated by different treatment. (E) Western blot bands and quantitative analysis of ChREBP in HK2 cells with ChREBP knockdown. (F) Western blot bands and quantitative analysis of Grp78 in HK2 cells stimulated with 5 mM fructose and different concentrations of 4‐PBA. (G) Western blot bands and quantitative analysis of Grp78 in HK2 cells stimulated by different concentrations of TM.

Table S1: Primer sequences.

Table S2: Antibodies information.

FSB2-40-e71774-s001.docx^{(20.6MB, docx)}

Movie S1: Normal cells were analyzed for calcium content in response to ATP stimulation. After loading mitochondrial and cytoplasmic calcium probes for stabilization, calcium flow was stimulated using ATP and representative images were obtained by video capture.

Download video file^{(5.3MB, avi)}

Movie S2: Fructose‐intervened cells were analyzed for calcium content in response to ATP stimulation. After loading mitochondrial and cytoplasmic calcium probes for stabilization, calcium flow was stimulated using ATP and representative images were obtained by video capture.

Download video file^{(12MB, avi)}

Contributor Information

Bei Sun, Email: sun_peipei220@hotmail.com.

Liming Chen, Email: xfx22081@vip.163.com.

Data Availability Statement

The datasets used and/or analyzed during the current study are available from the corresponding author on reasonable request.

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44. Stanhope K. L., Schwarz J. M., Keim N. L., et al., “Consuming Fructose‐Sweetened, Not Glucose‐Sweetened, Beverages Increases Visceral Adiposity and Lipids and Decreases Insulin Sensitivity in Overweight/Obese Humans,” Journal of Clinical Investigation 119, no. 5 (2009): 1322–1334, 10.1172/JCI37385. [DOI] [PMC free article] [PubMed] [Google Scholar]
45. Softic S., Gupta M. K., Wang G.‐X., et al., “Divergent Effects of Glucose and Fructose on Hepatic Lipogenesis and Insulin Signaling,” Journal of Clinical Investigation 127, no. 11 (2017): 4059–4074, 10.1172/JCI94585. [DOI] [PMC free article] [PubMed] [Google Scholar]
46. Mohamed B., Ghareib S. A., Alsemeh A. E., and El‐Sayed S. S., “Telmisartan Ameliorates Nephropathy and Restores the Hippo Pathway in Rats With Metabolic Syndrome,” European Journal of Pharmacology 973 (2024): 176605, 10.1016/j.ejphar.2024.176605. [DOI] [PubMed] [Google Scholar]
47. Feng D., Wang X., Song J., et al., “Association of Uric Acid and Fructose Levels in Polycystic Ovary Syndrome,” Human Reproduction 39 (2024): 2575–2586, 10.1093/humrep/deae219. [DOI] [PubMed] [Google Scholar]
48. Liu M., Shen J., Chen X., Dawuti T., and Xiao H., “Evaluating Renal Injury Characteristics in Different Rat Models of Hyperuricemia and Elucidating Pathological Molecular Mechanisms via Serum Metabolomics,” Frontiers in Pharmacology 15 (2024): 1433991, 10.3389/fphar.2024.1433991. [DOI] [PMC free article] [PubMed] [Google Scholar]
49. Liu Y., Qiao Y., Pan S., et al., “Broadening Horizons: The Contribution of Mitochondria‐Associated Endoplasmic Reticulum Membrane (MAM) Dysfunction in Diabetic Kidney Disease,” International Journal of Biological Sciences 19, no. 14 (2023): 4427–4441, 10.7150/ijbs.86608. [DOI] [PMC free article] [PubMed] [Google Scholar]
50. Xie Y., E J., Cai H., et al., “Reticulon‐1A Mediates Diabetic Kidney Disease Progression Through Endoplasmic Reticulum‐Mitochondrial Contacts in Tubular Epithelial Cells,” Kidney International 102, no. 2 (2022): 293–306, 10.1016/j.kint.2022.02.038. [DOI] [PMC free article] [PubMed] [Google Scholar]
51. Khunti K., Jabbour S., Cos X., et al., “Sodium‐Glucose Co‐Transporter‐2 Inhibitors in Patients With Type 2 Diabetes: Barriers and Solutions for Improving Uptake in Routine Clinical Practice,” Diabetes, Obesity & Metabolism 24, no. 7 (2022): 1187–1196, 10.1111/dom.14684. [DOI] [PMC free article] [PubMed] [Google Scholar]
52. Wright E. M. and Turk E., “The Sodium/Glucose Cotransport Family SLC5,” Pflügers Archiv/European Journal of Physiology 447, no. 5 (2004): 510–518. [DOI] [PubMed] [Google Scholar]
53. Zelniker T. A. and Braunwald E., “Mechanisms of Cardiorenal Effects of Sodium‐Glucose Cotransporter 2 Inhibitors: JACC State‐Of‐The‐Art Review,” Journal of the American College of Cardiology 75, no. 4 (2020): 422–434, 10.1016/j.jacc.2019.11.031. [DOI] [PubMed] [Google Scholar]
54. Osataphan S., Macchi C., Singhal G., et al., “SGLT2 Inhibition Reprograms Systemic Metabolism via FGF21‐Dependent and ‐Independent Mechanisms,” JCI Insight 4, no. 5 (2019): e123130, 10.1172/jci.insight.123130. [DOI] [PMC free article] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Table S1: Primer sequences.

Table S2: Antibodies information.

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Download video file^{(5.3MB, avi)}

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Data Availability Statement

The datasets used and/or analyzed during the current study are available from the corresponding author on reasonable request.

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PERMALINK

ChREBP‐β Exacerbates Renal Tubular Disorders Caused by Fructose via ATF4

Ting Fang

Xinyu Yang

Xiaoqing Deng

Jingyi Wang

Ting Li

Hang Guo

Bei Sun

Liming Chen

ABSTRACT

Abbreviations

1. Introduction

2. Materials and Methods

2.1. Mouse Experiments

2.2. Genotyping

2.3. Glucose Tolerance Tests

2.4. Insulin Tolerance Tests

2.5. mRNA Expression Analysis

2.6. Western Blot Analysis

2.7. Histological Analyses

2.8. Transmission Electron Microscopy Examination

2.9. TUNEL Assay

2.10. Measurement of Mitochondrial Oxygen Consumption

2.11. In Situ Proximity Ligation Assay (PLA)

2.12. Isolation of Primary Renal Tubular Epithelial Cells

2.13. Cell Culture

2.14. Ca2+ Measurements

2.15. Flow Cytometry

2.16. FITC‐BSA Staining

2.17. Dual Luciferase Assay

2.18. Chromatin Immunoprecipitation (ChIP) Analysis

2.19. Statistical Analysis

3. Results

3.1. High Fructose Induces Insulin Resistance and Renal Tubular Injury

FIGURE 1.

3.2. Generation of Renal Tubular‐Specific ChREBP‐β‐Overexpressing Mice

FIGURE 2.

3.3. High Fructose or Renal Tubular‐Specific ChREBP‐β Overexpression Induces Mitochondrial Dysfunction

FIGURE 3.

3.4. High Fructose‐ Or Renal Tubular‐Specific ChREBP‐β Overexpression Induces Kidney ER Stress and Calcium Homeostasis Disorders

FIGURE 4.

3.5. ChREBP Knockout Relieves Fructose‐Mediated Renal Tubular Injury

FIGURE 5.

3.6. ChREBP‐β Induces Renal Tubular ER Stress, Mitochondrial Dysfunction, and Apoptosis

FIGURE 6.

3.7. ChREBP Knockout Relieves Fructose‐Mediated ER Stress to Improve Mitochondrial Dysfunction in HK2 Cells

FIGURE 7.

3.8. ChREBP‐β Bind to ATF4 Promoter and Increase Its Transcription

FIGURE 8.

4. Discussion

Author Contributions

Funding

Ethics Statement

Conflicts of Interest

Supporting information

Contributor Information

Data Availability Statement

References

Associated Data

Supplementary Materials

Data Availability Statement

ACTIONS

PERMALINK

RESOURCES

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2.14. Ca²⁺ Measurements