Urotensin II Receptor Deletion Mitigates Kidney Injury and Renal Lipid Remodeling in Mice Fed a Long‐Term High‐Fat Diet

ABSTRACT

Urotensin II (UII) signaling through its Gq‐coupled receptor (UT) has been implicated in metabolic and vascular dysfunction; however, its contribution to diabetic kidney disease (DKD) and renal lipid remodeling remains incompletely defined. We examined kidney injury and lipidomic alterations in wild‐type (WT) and UT knockout (UT KO) mice fed a long‐term high‐fat diet (HFD). Male WT and UT KO mice were fed HFD (60% kcal fat) or low‐fat diet (LFD; 10% kcal fat) for 33 weeks. Despite comparable HFD‐induced obesity, hyperglycemia, and hyperinsulinemia, UT deletion significantly attenuated albuminuria, glomerular and tubular fibrosis, histopathologic injury scores, glomerular basement membrane thickening, and podocyte foot process effacement. UT deficiency also reduced systemic and renal markers of endoplasmic reticulum (ER) stress. Untargeted renal lipidomics profiling (1320 lipid species) revealed no genotype‐dependent differences under LFD conditions. In contrast, HFD altered 52 lipid species in WT kidneys, prominently affecting phospholipids, glycosphingolipids, ceramides, sphingomyelins, and triglycerides. UT deletion mitigated dysregulation of 41 of these lipid species, indicating that UT signaling promotes maladaptive renal lipid remodeling in DKD. UT expression was confirmed in native and cultured mouse podocytes. In vitro, UII stimulated ER Ca²⁺ release, induced ER stress, and disrupted the actin cytoskeleton in podocytes; these effects were prevented by UT antagonism or inhibition of transient receptor potential canonical 6 channels. Collectively, these findings identify UII–UT signaling as a key mediator of lipid dysregulation and global kidney injury affecting the glomerular filtration barrier, tubules, and interstitium under HFD, supporting UT antagonism as a potential renometabolic therapeutic strategy.

Urotensin II receptor deletion mitigates high‐fat diet‐induced kidney injury and renal lipid remodeling. Male wild‐type (WT) and urotensin II receptor–knockout (UT KO) mice were fed a high‐fat diet (HFD; 60% kcal from fat) for 33 weeks. In cultured mouse podocytes, urotensin II (UII)–UT signaling enhanced transient receptor potential canonical 6 (TRPC6)–dependent intracellular Ca²⁺ signaling and promoted endoplasmic reticulum (ER) stress, whereas pharmacological inhibition of UT or TRPC6 attenuated these cellular responses. In vivo, global UT deletion reduced HFD‐induced renal ER stress, glomerular and podocyte injury, albuminuria, kidney injury biomarkers, and maladaptive renal lipid remodeling. Together, these findings support a contribution of UII–UT signaling to kidney injury and renal lipid remodeling during long‐term HFD feeding. Ca²⁺, calcium; DKD, diabetic kidney disease; ER, endoplasmic reticulum; HFD, high‐fat diet; TRPC6, transient receptor potential canonical 6; UII, urotensin II; UT, urotensin II receptor; UT KO, urotensin II receptor knockout; WT, wild type. (Created with BioRender.com).

1. Introduction

Urotensin II (UII) is a cyclic peptide hormone first identified in the caudal neurosecretory system of fish [1]. Subsequently, UII expression has been demonstrated in mammals, including humans [2, 3], across multiple organ systems, ranging from the central nervous system to the cardiovascular and renal systems [2, 3]. UII acts via a G‐protein–coupled receptor, the urotensin II receptor (UT), which is also widely expressed in the nervous, cardiovascular, and renal systems [2, 3, 4]. Functionally, UII is best known as a potent vasoconstrictor [2, 5, 6], although endothelium‐dependent vasorelaxant effects have also been reported [7, 8]. Dysregulation of the UII/UT system has been implicated in several cardiovascular and renal disorders, including hypertension, heart failure, and chronic kidney disease [3, 4, 9, 10].

Over the past two decades, converging clinical and experimental evidence has linked UII signaling to diabetes mellitus (DM) and its renal complications. UII and UT expression are increased in the kidneys of humans and animal models with DM [7, 8], and patients with DM exhibit elevated plasma UII concentrations and urinary UII excretion [11]. Genetic association studies have identified single‐nucleotide polymorphisms in the UII gene locus that correlate with DM risk [12, 13]. In diabetic rodents, elevated UII levels promote renal fibrosis and functional decline, and pharmacological UT inhibition reduces albuminuria in patients with diabetic kidney disease (DKD) [7, 14]. UII‐induced endoplasmic reticulum (ER) stress has also been documented in renal tubules of diabetic mice [15].

We recently reported that global UT knockout (UT KO) in mice protects against streptozotocin (STZ)–induced hyperglycemia and kidney injury, a model of Type 1 DM [16]. Independent studies have shown that targeting the UT system, either genetically or pharmacologically, attenuates atherosclerosis and metabolic disturbances in multiple rodent models [17, 18, 19, 20, 21], underscoring UT as a key modulator of cardiometabolic disease. However, whether UT deletion confers protection against DKD secondary to Type 2 DM (T2DM), which is typically driven by obesity, insulin resistance, and lipotoxicity, remains unclear.

Podocyte injury is an essential component of DKD pathogenesis, leading to disruption of the glomerular filtration barrier, foot process effacement, and albuminuria [22, 23, 24]. Perturbations in intracellular Ca²⁺ homeostasis, ER stress, and actin cytoskeletal remodeling are critical mediators of podocyte dysfunction [24, 25, 26, 27]. Transient receptor potential canonical 6 (TRPC6) channels are Ca²⁺‐permeable, mechanically and receptor‐operated channels enriched in podocytes, where gain‐of‐function mutations or heightened activity cause proteinuric kidney disease and focal segmental glomerulosclerosis [25, 28, 29, 30, 31, 32, 33]. UT is a Gq‐coupled receptor that signals via phospholipase C, a pathway well‐positioned to engage TRPC6‐dependent Ca²⁺ influx [34, 35]. Yet, whether UT activation in podocytes functionally couples with TRPC6 to drive Ca²⁺ dysregulation, ER stress, and cytoskeletal injury in DKD remains unestablished.

In parallel, aberrant renal lipid accumulation and remodeling are increasingly recognized as pathogenic drivers of DKD. Lipotoxicity in glomerular and tubular compartments promotes mitochondrial dysfunction, oxidative stress, inflammation, and fibrosis [22, 23, 27]. However, the relationship between UII signaling and the renal lipidome remains poorly defined. Consequently, it is unclear whether UT signaling merely reflects metabolic stress or actively promotes maladaptive lipid remodeling in the diabetic kidney.

In the present study, we addressed these gaps using a high‐fat diet (HFD) model of T2DM‐associated DKD in global UT KO mice, complemented by mechanistic studies in cultured mouse podocytes. We tested the hypothesis that UT signaling promotes global kidney injury and maladaptive renal lipid remodeling in T2DM DKD. We show that UT deletion mitigates HFD‐induced albuminuria, glomerular and tubular structural injury, ER stress, and kidney injury biomarkers, and limits HFD‐driven changes in the renal lipidome. At the cellular level, we identify UT expression in native and cultured mouse podocytes and demonstrate that UII activates a TRPC6‐dependent Ca²⁺ influx pathway that triggers ER stress and actin cytoskeletal fragmentation, as visualized by super‐resolution microscopy. These findings define a previously underappreciated UT–TRPC6– Ca²⁺ signaling axis and support UT as a therapeutic target for limiting DKD progression and associated renal lipid remodeling in T2DM.

2. Results

2.1. UT Deletion Reverses HFD‐Induced Albuminuria

Our initial protocol of 21 weeks of high‐fat diet (HFD) feeding did not result in significant fasting hyperglycemia in the mice, despite significant weight gain during the same period (Figure 1A,B). Mice fed an HFD for 33 weeks exhibited significant increases in body weight and plasma glucose levels compared with LFD–fed mice (Figure 1A,B). HFD feeding also resulted in markedly elevated plasma insulin concentrations, consistent with the development of peripheral insulin resistance (Figure 1C). Deletion of the urotensin II receptor (UT KO) did not alter HFD‐induced weight gain at 21 and 33 weeks, and hyperglycemia and hyperinsulinemia at 33 weeks (Figure 1A–C). In contrast, UT KO mice fed an HFD exhibited significantly reduced urinary albumin excretion compared with WT HFD mice (Figure 1D), indicating a protective effect of UT deletion on kidney function independent of systemic metabolic control. Plasma glucagon levels were not significantly affected by HFD feeding in either genotype (Figure 1E). Consistent with prior reports in diabetic patients and experimental models [7, 8, 10], renal UII and UT mRNA expression were significantly increased in kidneys from HFD‐fed mice (Figure 1F,G).

FIGURE 1.

High‐fat diet caused significant increases in (A) body weight, (B) fasting blood glucose, and (C) plasma insulin levels in both WT and UT KO. Fasting blood glucose levels at 21 weeks were not significantly elevated by HFD despite a significant increase observed in body weight. (D) Normalized urine albumin excretion was significantly elevated in only WT HFD but not KO HFD, suggesting a probable protective effect of UT KO. (E) Glucagon levels were unaltered. (mean ± SEM; n = 5; *p < 0.05, **p < 0.01, ***p < 0.001, One‐way or two‐way ANOVA with Tukey's multiple comparisons test). A high‐fat diet increased (F) UII and (G) UT mRNA expression in kidney tissue. (mean ± SEM; n = 5; *p < 0.05, Mann Whitney test).

2.2. UT Deletion Protects Against HFD‐Induced Kidney Fibrosis and Histopathologic Injury

Immunohistochemical analysis of kidney sections from WT mice fed an HFD revealed markedly increased collagen IV and fibronectin staining compared with LFD–fed mice, indicating enhanced renal fibrosis (Figure 2A–F). This fibrotic response was largely absent in KO mice subjected to HFD. Increased collagen IV and fibronectin immunoreactivity was observed in both glomerular and tubular compartments in WT HFD kidneys. Consistent with these findings, histopathologic evaluation of PAS–stained kidney sections showed a significantly higher combined renal injury score in WT HFD mice, whereas UT deletion markedly mitigated HFD‐induced injury (Figure 2G,H). WT HFD kidneys exhibited prominent glomerular tuft expansion, mesangial matrix accumulation, and proximal tubular epithelial cytoplasmic vacuolization. In contrast, UT KO HFD kidneys preserved glomerular architecture, with minimal mesangial expansion and reduced tubular injury.

FIGURE 2.

UT KO alleviates high‐fat diet‐induced interstitial fibrosis, and glomerular and tubular injury. A high‐fat diet significantly increased total kidney and intraglomerular collagen IV and fibronectin content. (A), (B) and (C) show an increase in collagen IV staining (brown) in WT HFD mice, which UT KO reversed. (D), (E) and (F) show increased fibronectin stain intensity in WT HFD, which was reversed in KO HFD. (G) Pathological scoring of Periodic Acid–Schiff (PAS) stained kidney sections showed significantly higher combined injury score in WT HFD, with injury mitigation in UT KO. Each panel shows glomeruli on the left and tubules on the right. WT HFD images exhibit moderate distention of the glomerular interstitium by a PAS‐positive material (yellow arrow—indicating glomerulonephritis/membranoproliferative changes) and well‐demarcated cytoplasmic vacuolation (white arrow), mainly of the epithelial cells of the proximal tubules. Glomerular distension with mesangial expansion was notably absent in KO HFD. (H) Combined injury scores (comprising inflammatory cell infiltrates, tubular casts, epithelial vacuolization, and mesangial expansion; n = 5; One‐way ANOVA with Tukey's multiple comparisons test) *p < 0.05, **p < 0.01, ***p < 0.001.

2.3. UT Deletion Preserves Glomerular Filtration Barrier Integrity in HFD‐Fed Mice

Given the marked albuminuria observed in WT HFD mice and its attenuation in UT KO HFD mice, we next examined the ultrastructural integrity of the glomerular filtration barrier by transmission electron microscopy (TEM) of the kidney cortex. TEM analysis revealed pronounced podocyte foot process effacement in WT HFD mice, evidenced by a significant increase in foot process width (Figure 3A,B). In addition, glomeruli from WT HFD mice displayed marked glomerular basement membrane thickening (Figure 3A,C), a recognized early ultrastructural hallmark of diabetic nephropathy [36]. Both podocyte foot process effacement and basement membrane thickening were significantly attenuated in UT KO mice fed an HFD, indicating preservation of glomerular filtration barrier architecture. These findings demonstrate that UII–UT signaling adversely affects key components of the filtration barrier, particularly podocytes, and that UT deletion confers structural protection against HFD‐induced glomerular injury.

FIGURE 3.

Transmission electron microscopic imaging of kidney glomeruli. (A) Representative images of mouse glomeruli showing glomerular basement membrane (GBM) and podocyte foot processes. Yellow arrowheads indicate areas of significant podocyte foot process effacement. White arrowheads show examples of normal foot processes. Yellow parallel lines indicate thickening of the GBM, white parallel lines show normal GBM. Scale bars: 2 μM. WT HFD showed (B) podocyte foot process effacement, and (C) thickening of GBM, which were mitigated by UT KO. Foot process width was calculated using the formula: FPW $=\frac{\mathrm{\pi }.}{4.}$ $\frac{\text{Total length of}\mathrm{GBM}}{\text{number of foot processes}}$, where the correction factor π/4 accounts for the random orientation of glomeruli in tissue sections (n = 22 glomerular sections from 5 mice; One‐way ANOVA with Tukey's multiple comparisons test; *p < 0.05, **p < 0.01, ****p < 0.0001).

2.4. UT Is Expressed in Podocytes

Although previously known sites of UT expression in the kidneys, such as mesangial cells [37, 38, 39], tubular cells, and the interstitium [8, 10] are sufficient to explain the adverse effects observed in the glomerular matrix and kidney parenchyma, the involvement of podocyte foot processes and the GBM suggests a potential role for UT in podocytes. UT expression in podocytes has not been previously established. Therefore, to determine whether UT is expressed in podocytes in situ, we performed immunofluorescence staining of mouse glomerular explants with antibodies against UT, together with established podocyte markers, nephrin and podocin. As shown in Figure 4A,B, UT immunoreactivity robustly overlapped spatially with both nephrin‐ and podocin‐positive cells within the glomerulus, indicating UT expression in podocytes. To validate these findings and exclude potential confounding effects of neighboring glomerular cell types, we next examined UT expression in a cultured mouse podocyte cell line. Nested PCR analysis demonstrated the presence of UT mRNA in podocytes, yielding a product of the expected size, while appropriate negative controls showed no amplification (Figure 4C). Consistent with these transcriptional data, immunofluorescence analysis of cultured podocytes revealed UT protein expression, which localized predominantly to the plasma membrane and cytoplasmic compartments (Figure 4D). UT staining was observed in cells exhibiting intact actin cytoskeletal organization, further supporting functional receptor expression in podocytes (Figure 4D). Together, these complementary in situ and in vitro approaches establish that podocytes express UT, providing a structural basis for direct UII–UT signaling within the glomerular filtration barrier and supporting a podocyte‐intrinsic role for UT in diabetic kidney disease.

FIGURE 4.

Ut is expressed by podocytes in glomerular explants and in culture. Colocalization of (A) UT (red) and nephrin (green), and (B) UT (red) and podocin (green) in glomerular explants. (C) A nested PCR gel blot image of an immortalized podocyte cell line shows UT expression. WT‐1 is a podocyte marker. (D) Immunofluorescence images of cultured podocytes show UT (green) and F‐Actin (red).

2.5. UII Triggers Cellular ER Stress and Ca²⁺ Release

Having established that UT is expressed in podocytes and that UT deletion protects against HFD‐induced glomerular barrier dysfunction, we next investigated the cellular mechanisms by which UII promotes podocyte injury. Immunofluorescence staining of kidney sections demonstrated markedly increased expression of ER stress markers GRP78/BiP and C/EBP homologous protein (CHOP) in WT mice fed an HFD (Figure 5A–E). Elevated ER stress signals were evident throughout the renal cortex, including both glomerular and tubular compartments. In contrast, UT KO mice subjected to HFD exhibited reduced GRP78/BiP and CHOP staining, indicating attenuation of HFD‐induced renal ER stress. These observations were further corroborated by quantitative RT‐PCR analysis of ER stress–related gene GRP94 in renal tissue (Figure 5F), as well as by reduced circulating levels of the ER stress–associated protein CRELD2 in UT KO HFD mice (Figure 5G).

FIGURE 5.

HFD‐induced ER stress is attenuated by UT knockout in the mouse kidney, including the glomeruli. (A) Immunofluorescence imaging of kidney sections showed a significant increase in total kidney and glomerular staining of ER stress markers, GRP78/BiP (B, C) and CHOP (D, E) in WT HFD, the effects reversed by UT KO (n = 15 data points from 5 mice; One‐way ANOVA with Tukey's multiple comparisons test). A high‐fat diet (F) increased kidney tissue mRNA expression of ER stress marker GRP94 (n = 5; Kruskal–Wallis test) and (G) elevated plasma levels of the soluble ER stress marker CRELD2 (n = 5; One‐way ANOVA with Tukey's multiple comparisons test). These effects were alleviated by UT KO. (H) UII (5 μM) increased [Ca²⁺]_I, and (I) depletion of the ER Ca²⁺ store with 10 μM thapsigargin prevented the action of UII on subsequent exposure. (J) UII‐induced ER Ca²⁺ release blocked by thapsigargin (mean ± SEM; n = 6–7, unpaired t‐test); *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001.

To determine whether alterations in intracellular Ca²⁺ homeostasis drive UII‐induced ER stress, we performed ratiometric Ca²⁺ imaging in cultured mouse podocytes. Acute application of UII elicited a rapid increase in intracellular Ca²⁺ levels in cells superfused with Ca²⁺‐free solution containing EGTA (Figure 5H), indicating mobilization of Ca²⁺ from intracellular stores. Reintroduction of extracellular Ca²⁺ in the presence of the L‐type Ca²⁺ channel blocker nimodipine resulted in a robust Ca²⁺ influx, consistent with activation of store‐operated Ca²⁺ entry (SOCE). Importantly, pretreatment with thapsigargin, an inhibitor of the ER Ca²⁺‐ATPase, abolished the UII‐induced Ca²⁺ signal, confirming that ER Ca²⁺ release is required for this response (Figure 5I,J). Collectively, these data demonstrate that UII–UT signaling induces ER stress in podocytes by triggering ER Ca²⁺ release and subsequent activation of SOCE, providing a mechanistic link between UII signaling, Ca²⁺ dysregulation, and podocyte injury.

2.6. TRPC6 but Not TRPC5 Mediates UII‐Induced Intracellular Ca²⁺ Elevation and Cation Currents in Podocytes

Previous studies, including our own, have demonstrated that transient receptor potential (TRP) channels are critical mediators of intracellular Ca²⁺ signaling in renal cells [40, 41, 42, 43, 44]. Among these channels, TRPC5 and TRPC6 are expressed in podocytes and have been implicated in the pathogenesis of glomerular disease [29, 30, 31, 32, 33]. We therefore tested whether TRPC5 and/or TRPC6 mediate UII‐induced Ca²⁺ signaling in podocytes. To this end, we used the selective TRPC5 inhibitor AC1903 [45] and the TRPC6 inhibitor SAR7334 [46]. Ratiometric Ca²⁺ imaging revealed that UII‐induced increase in intracellular Ca²⁺ was markedly attenuated by TRPC6 inhibition with SAR7334, whereas blockade of TRPC5 with AC1903 had no significant effect on UII‐induced Ca²⁺ influx (Figure 6A,B). These findings indicate that TRPC6, but not TRPC5, is required for UII‐mediated Ca²⁺ entry in podocytes.

FIGURE 6.

UII stimulates TRPC6‐mediated SOCE and cation currents in podocytes. (A) UII‐induced SOCE in cultured mouse podocytes was significantly reduced by TRPC6 inhibition (SAR 7443), but not by TRPC5 inhibition (AC 1903). (B) Plot of Δ ratios depicting the SOCE‐associated increase in [Ca²⁺]_i with UII, TRPC5, and TRPC6 inhibition. (mean ± SEM; n = 6–8; One‐way ANOVA with Tukey's multiple comparisons test). Whole‐cell patch‐clamp recordings of cation currents from cultured mouse podocytes show (C) lack of inhibition of UII‐induced cation currents by TRPC5 inhibitor (20 μM AC 1903). (D) TRPC6 inhibition (200 nM SAR 7443) significantly reduced UII‐induced cation currents. (mean ± SEM; n = 7–8; One‐way repeated measures ANOVA with Tukey's multiple comparisons test); *p < 0.05, **p < 0.01, ***p < 0.001.

To directly assess the contribution of TRPC channels to UII‐induced ionic conductance, we performed whole‐cell patch‐clamp recordings in cultured mouse podocytes. TRPC‐mediated cation currents were evoked using a voltage ramp protocol from −100 to +100 mV (960 ms) (Figure 6C,D). UII markedly enhanced both inward and outward cation currents relative to control conditions (Figure 6C,D). These UII‐induced currents were significantly inhibited by SAR7334 (Figure 6D), but were unaffected by AC1903 (Figure 6C), further confirming the functional involvement of TRPC6 rather than TRPC5. Collectively, these data demonstrate that UII selectively activates a UT–TRPC6 signaling axis in podocytes, driving intracellular Ca²⁺ elevation and cation currents, while TRPC5 does not contribute to these responses.

2.7. Inhibition of UT and TRPC6 Reverses UII‐Induced ER Stress in Podocytes

To directly assess whether UII‐induced ER stress in podocytes is mediated through UT and downstream TRPC6 signaling, cultured mouse podocytes were exposed to UII for 72 h in the presence or absence of selective UT or TRPC6 inhibitors. Immunofluorescence analysis demonstrated that prolonged UII exposure markedly increased the expression of ER stress markers CHOP, a nuclear marker of the maladaptive unfolded protein response, and GRP78/BiP, a cytoplasmic ER chaperone (Figure 7A–C). UII treatment led to robust nuclear accumulation of CHOP and increased cytoplasmic GRP78/BiP immunoreactivity relative to control cells, indicating activation of ER stress pathways. Pharmacological inhibition of UT with GSK1562590 or TRPC6 with SAR7334 significantly attenuated UII‐induced increases in both CHOP and GRP78/BiP signal intensity, restoring ER stress marker expression toward control levels (Figure 7A–C). These findings indicate that UII‐mediated ER stress in podocytes requires UT activation and downstream TRPC6‐dependent signaling. To further corroborate these cellular findings, we quantified soluble ER stress–associated proteins released into the culture medium. Consistent with the immunofluorescence data, UII treatment significantly increased extracellular levels of CRELD2 and DNAJB11, two established markers of ER stress and activation of the unfolded protein response (Figure 7D,E). Inhibition of either UT or TRPC6 significantly reduced the secretion of both CRELD2 and DNAJB11, confirming effective suppression of UII‐induced ER stress at the molecular level (Figure 7D,E). Together, these results demonstrate that UII induces ER stress in podocytes via a UT–TRPC6 signaling pathway and that pharmacological targeting of either UT or TRPC6 effectively reverses this pathological response, providing mechanistic support for UT–TRPC6–Ca²⁺ signaling as a driver of podocyte injury.

FIGURE 7.

UII‐induced increase in podocyte ER stress is reversed by UT and TRPC6 inhibition. (A) UII (5 μM) increased ER stress in cultured mouse podocytes as indicated by the increase in fluorescence of GRP78 (Green) and CHOP (Red). Inhibition of UT (GSK1562590) and TRPC6 (SAR 7443) reversed this effect. Quantification of normalized fluorescent densities of (B) GRP78 (BiP) and (C) CHOP. (mean ± SEM; n = 25–26; One‐way ANOVA with Tukey's multiple comparisons test). UII treatment of cultured podocytes significantly elevated the concentrations of soluble ER stress markers (D) CRELD2 and (E) DNAJB11 in podocyte culture media. These were reversed by inhibition of UT (GSK) and TRPC6 (SAR) (mean ± SEM; n = 6–10; One‐way ANOVA with Tukey's multiple comparisons test); *p < 0.05, **p < 0.01, ****p < 0.0001.

2.8. UII Disrupts Podocyte Actin Cytoskeletal Organization via UT–TRPC6 Signaling

Dynamic regulation of the podocyte actin cytoskeleton is essential for maintaining podocyte morphology, motility, and the integrity of the glomerular filtration barrier [29, 47]. Dysregulation of intracellular Ca²⁺ signaling, particularly via TRPC channels, has been shown to profoundly influence actin dynamics and podocyte structure [29, 47]. Based on our findings that UII activates UT–TRPC6–dependent Ca²⁺ signaling, we next examined whether UII disrupts actin cytoskeletal organization in podocytes. Super‐resolution confocal microscopy was used to visualize F‐actin architecture in cultured mouse podocytes, followed by quantitative analysis of filament length distribution using an open‐source image analysis platform. Under control conditions, podocytes exhibited long, well‐organized actin stress fibers aligned along the longitudinal axis of the cell (Figure 8A). In contrast, UII treatment resulted in pronounced actin cytoskeletal disorganization, characterized by fragmentation of stress fibers and a significant shift toward shorter actin filament lengths (Figure 8A–D). Quantitative analysis confirmed a reduction in long filaments and a corresponding increase in short filament populations in UII‐treated podocytes (Figure 8B–D). Pharmacological inhibition of UT with GSK1562590 or of TRPC6 with SAR7334 markedly attenuated UII‐induced actin fragmentation and restored the filament length distribution to control levels (Figure 8A–D). These protective effects were not observed with TRPC5 inhibition in prior experiments, consistent with a selective role for TRPC6 in mediating UII‐induced cytoskeletal remodeling. Collectively, these findings demonstrate that UII disrupts podocyte actin cytoskeletal organization through a UT–TRPC6–dependent signaling pathway. This cytoskeletal remodeling offers a possible explanation for how UII‐induced Ca²⁺ dysregulation and ER stress may contribute to glomerular filtration barrier dysfunction.

FIGURE 8.

Super‐resolution microscopy of cultured mouse podocytes shows significant fragmentation of Actin filaments with UII, an effect reversed by inhibition of UT and TRPC6. (A) Representative images of podocyte Actin filaments (Green) in control (DMSO), UII (5 μM), GSK (1 μM) + UII, and SAR (200 nM) + UII. (B–D) Activation of UT caused a significant increase in the fragmentation of Actin filaments as indicated by a decrease in the percentage of long filaments (151–200 and 101–150 a.u.) and an increase in short filaments (51–100 a.u.; mean ± SEM; n = 10–13; One‐way ANOVA with Tukey's multiple comparisons test; *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001).

2.9. UT Deletion Attenuates HFD‐Induced Elevation of Kidney Injury Biomarkers

Having demonstrated that UII–UT signaling induces podocyte ER stress, Ca²⁺ dysregulation, cytoskeletal disruption, and structural injury to the glomerular filtration barrier, we next evaluated whether UT deletion mitigates kidney injury at the systemic, urinary, and tissue levels in the setting of HFD‐induced diabetes. To this end, we quantified established kidney injury biomarkers in plasma, urine, and renal tissue sections from WT and UT KO mice fed LFD or HFD diets. Plasma concentrations of the kidney injury biomarkers neutrophil gelatinase–associated lipocalin (NGAL) and cystatin C were significantly elevated in WT mice fed an HFD compared with LFD–fed mice (Figure 9A,B). In contrast, UT KO mice fed an HFD exhibited markedly lower plasma NGAL and cystatin C levels, approaching those observed in LFD–fed animals (Figure 9A,B). Similarly, urinary excretion of the tubular injury markers retinol‐binding protein 4 (RBP4) and α1‐microglobulin was significantly increased in WT HFD mice. In contrast, these increases were attenuated in UT KO HFD mice (Figure 9C,D), indicating preservation of tubular function. Consistent with these systemic and urinary findings, immunofluorescence analysis of kidney sections revealed robust upregulation of the cell cycle arrest–associated kidney injury biomarkers tissue inhibitor of metalloproteinase 2 (TIMP2) and insulin‐like growth factor–binding protein 7 (IGFBP7) in WT HFD kidneys (Figure 9E–I). Increased TIMP2 and IGFBP7 expressions were observed both globally throughout the renal cortex and within glomerular structures (Figure 9E). Notably, UT deletion significantly reduced HFD‐induced increases in TIMP2 and IGFBP7 immunoreactivity at both the whole‐kidney and glomerular levels (Figure 9E–I). These findings demonstrate that UT deletion protects against chronic HFD‐induced kidney injury, as evidenced by reduced circulating and urinary injury biomarkers and diminished tissue‐level injury responses, further implicating UII–UT signaling as a potential contributor to this process.

FIGURE 9.

UT KO mitigated HFD‐induced increase in kidney injury biomarkers. Plasma neutrophil gelatinase‐associated lipocalin (A) and cystatin C (B), and urinary retinol‐binding protein 4 (C) and α1‐macroglobulin (D) were elevated in WT HFD, but not in KO HFD mice (n = 5; One‐way ANOVA with Tukey's multiple comparisons test). (E) Immunofluorescence imaging of kidney slices shows increased expression of kidney injury biomarkers in WT HFD. Both total kidney and glomerular staining for tissue inhibitor of metalloproteinase 2 (F, G) and insulin‐like growth factor‐binding protein 7 (H, I) were significantly increased in WT HFD. This effect was reversed by UT KO (n = 15–25 data points from 5 mice; One‐way ANOVA with Tukey's multiple comparisons test). *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001.

2.10. UT Deletion Mitigates HFD‐Induced Renal Lipid Remodeling

To determine whether UT deletion attenuates lipid metabolic disturbances associated with T2DM pathogenesis [48, 49, 50], we performed comprehensive untargeted lipidomic profiling of kidney tissues. As expected, HFD feeding profoundly altered the renal lipidomic landscape in WT mice (Figure 10, WT HFD). Among the 1320 lipid species quantified, none differed significantly between WT LFD and KO LFD kidneys (the lowest p value = 0.3288 across all comparisons), indicating that UT deletion alone does not perturb basal renal lipid homeostasis. In contrast, HFD feeding in WT mice resulted in differential regulation of 52 lipid species relative to WT LFD controls, with 21 lipids significantly increased and 31 significantly decreased. These changes encompassed multiple lipid classes implicated in diabetic kidney injury, including phospholipids (phosphatidylcholines and phosphatidylethanolamines), sphingolipids (ceramides, sphingomyelins, and glycosphingolipids), and neutral lipids (triglycerides). Several of the HFD‐induced lipid alterations were consistent with lipotoxic stress signatures linked to membrane remodeling, mitochondrial dysfunction, and ER stress in DKD. Notably, UT deletion mitigated dysregulation of 41 of the 52 HFD‐responsive lipid species. Specifically, 16 lipid species that were increased and 25 lipid species that were decreased in WT HFD kidneys were partially or fully normalized in UT KO HFD mice. The lipid species rescued by UT deletion were enriched in ceramides and glycosphingolipids, lipid classes known to promote cellular stress signaling and apoptosis [51, 52, 53, 54], as well as in phospholipids critical for membrane integrity and organelle function [55, 56, 57]. In contrast, 11 lipid species (5 increased and 6 decreased) were similarly altered in both WT HFD and UT KO HFD groups, suggesting that these lipid changes reflect diet‐driven metabolic stress independent of UT signaling.

FIGURE 10.

UT deletion attenuated HFD‐induced alterations in the renal lipid profile. Volcano plots show differentially regulated lipid species in kidney samples from KO LFD, WT HFD, and KO HFD mice relative to the WT LFD group. No lipid species were significantly altered in the KO LFD group (FDR > 0.05). In contrast, 52 lipid species were differentially regulated in the WT HFD group at FDR < 0.05, whereas only 11 of these species remained significantly altered in the KO HFD group. The Venn diagram summarizes the number of unique and shared significantly altered lipid species among the three groups relative to the WT LFD group. Of the 52 lipid species altered in WT HFD mice, 41 were unique to the WT HFD group and 11 were shared with the KO HFD group. No significantly altered lipid species were unique to the KO LFD or KO HFD groups, and no shared lipid species involved the KO LFD group. These findings indicate that UT deletion attenuated the HFD‐induced differential regulation of 41 of the 52 lipid species identified in WT mice.

These findings indicate that UT signaling contributes to maladaptive renal lipid remodeling in the diabetic kidney, particularly within lipid classes linked to lipotoxicity, membrane dysfunction, and ER stress. UT deletion not only reduced the magnitude of HFD‐induced lipid disturbances but also preserved a more physiological renal lipid composition under metabolic stress. A detailed list of differentially regulated lipid species and associated statistical parameters is provided in Table 1, and representative volcano plots and a Venn diagram summarizing the global lipidomic dataset, including the 41 lipid species mitigated by UT deletion, are shown in Figure 10.

TABLE 1.

Renal tissue lipidomics in HFD‐fed mice.

#	Lipid constituent	WT HFD (log10FC)	KO HFD (log10FC)	WT HFD (adjusted p)	KO HFD (adjusted p)
Lipid species significantly altered by WT HFD and not KO HFD
1	Plasmanyl‐PC O‐34:0	0.446751	0.219974	0.006135	0.24221
2	Plasmanyl‐PC O‐34:2	0.459587	0.264029	0.006135	0.149621
3	Plasmenyl‐PE P‐36:0	0.497734	0.117996	0.008537	0.677554
4	Cer[NS] d34:1	0.872793	0.659132	0.014383	0.104379
5	LysoPG 20:2	0.250569	0.220271	0.014383	0.056258
6	Plasmenyl‐PC P‐34:0	0.913174	0.473039	0.018759	0.300466
7	LysoPI 20:2	1.10129	1.02344	0.021446	0.056258
8	GlcCer[NDS] d40:2	0.739232	0.647265	0.021446	0.076128
9	BMP 36:4	1.83248	1.63513	0.026018	0.077133
10	Cer[NDS] d36:2	0.263234	0.106698	0.028911	0.512822
11	HexCer[NS] d34:2	1.74816	1.82866	0.032667	0.052421
12	GlcCer[NDS] d34:1	0.337294	0.356379	0.032885	0.052421
13	GlcCer[NDS] d34:2	1.16964	1.12547	0.032885	0.077133
14	PC[OH] OH‐40:4	1.01817	0.318676	0.033872	0.664606
15	PE 38:6	1.50944	1.20985	0.036231	0.145676
16	Plasmanyl‐PE O‐36:2	0.215313	0.131239	0.041433	0.317408
17	FA 22:3	−0.42631	−0.28793	0.005501	0.081486
18	FA 24:3	−0.4323	−0.26536	0.006135	0.125532
19	PE 32:1	−0.29778	−0.18262	0.006135	0.125788
20	LysoPC 20:1	−0.32816	−0.23002	0.006135	0.08542
21	LysoPC 22:1	−0.29086	−0.21536	0.006135	0.076069
22	PC 33:6	−0.49587	−0.33501	0.014186	0.127754
23	PC 44:5	−0.28332	−0.1788	0.014383	0.163487
24	LysoPE 16:1	−0.29785	−0.20246	0.019109	0.148941
25	Plasmenyl‐PC P‐38:4	−0.63794	−0.50222	0.023502	0.114417
26	LysoPC 14:0	−0.28176	−0.16161	0.026242	0.270243
27	SM d32:1	−0.20034	−0.0973	0.026808	0.374884
28	Plasmenyl‐PE P‐34:4	−0.59457	−0.5551	0.026808	0.073717
29	PC 34:4	−0.2336	−0.16535	0.027386	0.162098
30	PC 36:6	−0.21652	−0.11837	0.032667	0.334606
31	PC 40:2	−0.22183	−0.20086	0.032885	0.098746
32	LysoPC 18:1	−0.25615	−0.14607	0.032885	0.317408
33	PC 32:2	−0.25419	−0.19238	0.033872	0.159647
34	SM d32:2	−0.19844	−0.1684	0.033872	0.124631
35	TG 42:0	−0.37067	−0.34842	0.035258	0.093751
36	SM d43:1	−1.79133	−0.0975	0.036231	0.98681
37	PC 32:1	−2.99239	−2.38252	0.036231	0.148941
38	PI 36:1	−0.235	−0.16196	0.036252	0.231179
39	AC 19:1	−0.41465	−0.37829	0.039698	0.114417
40	LysoPS 18:1	−0.2302	−0.1565	0.039839	0.245631
41	FA 19:3	−0.55501	−0.24218	0.045611	0.541268
Lipid species significantly altered by both WT HFD and KO HFD
1	DGDG 32:0	1.165850	1.246810	0.007347	0.027177
2	PG 32:0	0.425521	0.491244	0.029784	0.035919
3	HexCer[NS] d42:2	0.379457	0.505139	0.035258	0.027177
4	HexCer[NS] d42:3	0.573328	0.767668	0.039232	0.027177
5	GlcCer[NDS] d42:3	0.531609	0.654205	0.046365	0.040474
6	PC 38:2	−0.35493	−0.25942	0.002231	0.027177
7	PE 36:5	−0.42480	−0.30657	0.002231	0.035919
8	LysoPE 18:1	−0.47571	−0.33452	0.002231	0.042231
9	LysoPE 22:3	−0.56598	−0.49424	0.004619	0.027177
10	PE 34:3	−0.42588	−0.35734	0.005	0.035919
11	PC[OH] OH‐43:6	−0.68724	−0.78799	0.032667	0.035919

Note: Adjusted p values less than 0.05 are shown in bold.

3. Discussion

3.1. UT Activation Contributes to Diabetic Kidney Disease

In this study, we demonstrate that long‐term HFD feeding in mice induces obesity, hyperglycemia, hyperinsulinemia, and albuminuria, recapitulating key features of the T2DM phenotype. Deletion of the UT attenuated urinary albumin excretion without affecting HFD‐induced weight gain, hyperglycemia, or hyperinsulinemia. These findings indicate that UT deletion does not prevent the development of HFD‐induced diabetes per se but rather modifies downstream end‐organ injury in a tissue‐ and pathway‐specific manner. Collectively, our data suggest that systemic metabolic drivers of T2DM are mechanistically distinct from organ‐specific signaling pathways that determine renal vulnerability and the progression of diabetic kidney disease.

Our results extend prior work implicating the UII/UT system in DKD. Increased renal UII and UT expression have been reported in human diabetic nephropathy and in diabetic rodent models, and elevated circulating and urinary UII levels correlate with DKD severity and progression [8, 10, 11, 58, 59]. Genetic variants in the UII locus have been associated with T2DM risk [12, 13], and pharmacologic UT antagonism with palosuran reduced macroalbuminuria in a subset of patients with T2DM and nephropathy [14]. We now demonstrate that, in a diet‐induced T2DM model, UT deletion mitigates albuminuria, glomerular and tubular fibrosis, GBM thickening, and podocyte foot process effacement, in the setting of persistent obesity and hyperglycemia. Thus, UII–UT signaling appears not merely to be a biomarker of metabolic stress but also a causal effector of kidney injury in T2DM.

We further show that UII signaling contributes to HFD‐induced increases in collagen IV and fibronectin in both glomerular and tubular compartments. These findings are concordant with observations in STZ‐induced T1DM rats, in which diabetes increased renal UII, UT, TGF‐β₁, collagen IV, and fibronectin expression, and UII stimulation of proximal tubular cells enhanced TGF‐β₁ production in a UT‐dependent manner [7]. Together, these data support a model in which UII–UT signaling amplifies profibrotic pathways, likely including TGF‐β₁ and related mediators, in the diabetic kidney. Consistent with this view, we observed that HFD produced prominent glomerular and tubular injury, including membranoproliferative changes, tubular vacuolization, casts, and inflammatory infiltrates, and that UT KO markedly diminished these lesions. These results align with our recent work on STZ‐induced T1DM, in which UT KO reduced hyperglycemia and kidney injury, possibly through effects on pancreatic α‐cell function [16]. In contrast, in the present HFD model (T2DM), UT deletion confers renoprotection without improving systemic glycemia or hyperinsulinemia, suggesting that the kidney‐intrinsic effects of UT, including effects on podocytes, are sufficient to modify DKD progression even when systemic metabolic abnormalities persist. The differential effects of global UT KO on hyperglycemia in Type 1 and Type 2 DM models likely arise from the UII effects on pancreatic islet cells, which are the primary targets of STZ [16]. Collectively, these findings support UII–UT signaling as a shared pathogenic pathway in T1DM and T2DM models, although the relative contributions of endocrine pancreas versus renal UT signaling likely differ, with the islet cell effects of UII–UT being less pronounced or absent in T2DM.

3.2. UT as a Potential Pharmacological Target

The combined in vivo and in vitro findings presented here strongly support UT as a potential therapeutic target for DKD, particularly in T2DM driven by obesity and lipotoxicity. However, data from existing animal and clinical studies of UT antagonists are mixed. Palosuran, a small‐molecule UT antagonist with favorable safety and pharmacokinetic profiles in humans [60, 61], improved renal outcomes in models of ischemia–reperfusion injury [62] and STZ‐induced diabetes with uninephrectomy, where it enhanced survival, glycemic control, lipid profiles, and β‐cell preservation [63]. In contrast, palosuran did not alter first‐ and second‐phase insulin responses during a hyperglycemic clamp in T2DM patients [64], suggesting limited direct benefit on β‐cell secretory capacity in established disease.

Two clinical studies evaluated palosuran in hypertensive patients with T2DM and nephropathy [14, 65]. In one study, palosuran reduced macroalbuminuria in patients with mildly impaired renal function but not in those with more advanced impairment [14]. Another trial reported no significant overall effects on albuminuria, blood pressure, GFR, or renal plasma flow [65]. However, in both studies, the participants were taking either an ACE inhibitor or an angiotensin receptor blocker during the study period as treatment for hypertension. Hence, the interpretation of these trials is complicated by background RAAS blockade. Although palosuran functions as a UT antagonist, it has also been reported to activate somatostatin receptors [66]. In addition, its diminished affinity for UT in intact cells and vascular tissues suggests potential limitations in its utility as a selective UT antagonist [67]. Our data show that UT signaling drives renal lipotoxic remodeling and podocyte injury via Ca²⁺ dysregulation, ER stress, and cytoskeletal fragmentation, and suggest that highly selective UT antagonists may be most effective in earlier DKD stages characterized by active lipid remodeling and reversible podocyte injury, as well as in patient subsets with obesity and hyperlipidemia. Future studies will be required to define the optimal therapeutic window for UT inhibition, to determine whether more selective or biased UT ligands can preferentially suppress injurious Gq protein signaling while preserving potentially beneficial responses.

3.3. Lipidomics Profiles in DKD

Our untargeted lipidomic profiling demonstrates that UT deletion substantially attenuates HFD‐induced renal lipid remodeling. In WT mice, HFD altered 52 kidney lipid species across phospholipids (e.g., phosphatidylcholines, phosphatidylethanolamines), sphingolipids (ceramides, glycosphingolipids, sphingomyelins), and neutral lipids (triglycerides). Many of these lipid classes have been implicated in lipotoxic stress, mitochondrial dysfunction, and ER stress in DKD [48, 49, 50, 68, 69]. Notably, UT deletion normalized or partially mitigated 41 of these 52 HFD‐responsive lipids, including 16 that were increased and 25 that were decreased in WT HFD kidneys. The UT‐sensitive lipids were enriched in ceramides and glycosphingolipids, bioactive lipids linked to inflammatory signaling, insulin resistance, and apoptosis [51, 52, 53, 54], as well as in phospholipids essential for membrane integrity and organelle function [55, 56, 57]. In contrast, 11 lipids remained similarly altered in both WT HFD and UT KO HFD groups, suggesting that a subset of renal lipid changes reflects diet‐driven metabolic stress independent of UT signaling.

These findings are consistent with clinical and experimental lipidomics studies in DM and DKD. In obese women with T2DM, untargeted metabolomics revealed elevated ceramides, lysophosphatidylcholines, and multiple phosphatidylcholine and phosphatidylinositol species [70]. Increased ceramide content has been described in skeletal muscle and plasma of individuals with diabetes and insulin resistance [71, 72], and mitochondrial sphingomyelinase activation and lactosylceramide accumulation contribute to mitochondrial dysfunction in diabetic tissues [73, 74]. In the heart, insulin treatment can drive ceramide accumulation and cardiometabolic dysfunction [75].

In the context of DKD, specific phospholipid and sphingolipid species have been associated with the transition from uncomplicated DM to overt DKD and with CKD progression [68, 69, 76, 77]. LPC species such as LPC(16:0) and LPC(18:0) correlate with rapid kidney function decline and induce ER stress, mitochondrial impairment, and tubular lipotoxicity in experimental DKD [76]. Serum phospholipids, including PS(27:0)‐H and PS(30:2e)‐H, have been proposed as early DKD biomarkers [77]. Tissue‐specific analyses in diabetic mouse models show little overlap between plasma and kidney lipidomes, underscoring the need for direct renal lipid profiling [78].

Kidney‐focused studies have further demonstrated that lipid accumulation and remodeling drive glomerular hypertrophy, podocyte lipid deposition, mitochondrial injury, oxidative stress, and tubular damage [76, 79, 80]. In uninephrectomized HFD‐fed mice, cholesteryl ester 20:4 accumulation in podocytes has been linked to lipotoxicity, autophagy activation, and podocyte injury [79]. Our study adds to this body of work by providing evidence that genetic disruption of UT preserves a more physiologic renal lipid composition under chronic HFD stress. By mitigating ceramide and glycosphingolipid accumulation and stabilizing phospholipid composition, UT deletion likely reduces lipotoxic stress on podocytes and tubules, thereby dampening ER stress and structural injury.

Given the complexity of the renal lipidome and the heterogeneous findings across studies, it is premature to ascribe specific functional roles to each of the 41 UT‐sensitive lipids. Nonetheless, the consistent enrichment of these lipids in bioactive sphingolipids and membrane phospholipids strongly suggests that UT signaling intersects with pathways controlling renal lipid handling and organelle homeostasis. Integration of lipidomics with transcriptomics and metabolomics, and the use of cell–type–specific manipulations of UT signaling, will be the necessary next steps to define the precise molecular links between UII–UT, lipid metabolism, and DKD. To our knowledge, this is the first study to perform kidney tissue lipidomics in an HFD‐induced diabetes UT KO model. Our results reveal that UT deletion selectively normalizes a subset of HFD‐responsive lipids linked to lipotoxicity and ER stress, thereby providing a mechanistic bridge between UII–UT signaling, Ca²⁺ dysregulation, ER stress, and renal lipid remodeling in DKD.

3.4. Role of UT in Podocytes and Its Broader Impact on DKD

Because the role of UT in podocytes has not been explored previously, mechanistic studies were conducted in cultured mouse podocytes to confirm and further validate our findings. We demonstrate UT expression in podocytes in situ and in culture and show that UII triggers ER Ca²⁺ release, Ca²⁺ influx, and the induction of ER stress markers (GRP78/BiP, CHOP). Pharmacologic inhibition of UT or TRPC6, but not TRPC5, attenuates UII‐induced Ca²⁺ influx, reduces ER stress, and prevents actin filament fragmentation. Whole‐cell patch‐clamp recordings further demonstrate that podocytes exhibit UII‐induced cation currents which are sensitive to TRPC6 blockade and insensitive to TRPC5 inhibition, indicating that TRPC6 is the principal TRPC channel mediating UII‐induced Ca²⁺ entry in podocytes. These findings not only integrate and extend existing models of TRPC channel function in the podocyte cytoskeleton but also establish podocytes as a bona fide target of UII action. TRPC5 and TRPC6 have been described as exerting antagonistic effects on podocyte motility and actin dynamics, with TRPC5–Rac1 signaling favoring increased motility and loss of stress fibers, and TRPC6–RhoA signaling promoting stress fiber formation and reduced motility [29, 31]. Gain‐of‐function mutations and hyperactivation of TRPC6 cause focal segmental glomerulosclerosis and proteinuric kidney disease [32, 33, 81]. Our data add UII–UT to the repertoire of Gq‐coupled stimuli, alongside angiotensin II, endothelins, and thromboxanes, that activate TRPC6 and promote glomerular injury [82, 83]. Given that UT is a Gq‐coupled GPCR that signals via phospholipase C [5, 34, 35, 37, 84], it is plausible that UII–UT signaling generates diacylglycerol that directly activates TRPC6 [85].

Our Ca²⁺ imaging experiments show that UT activation elicits ER Ca²⁺ release followed by SOCE, and that the SOCE component is blunted by TRPC6 inhibition. These findings suggest that TRPC6 either contributes directly to SOCE or is functionally coupled to Orai1 and STIM1, as reported for TRPC3/6 complexes in other cell types [86, 87, 88, 89]. In the context of diabetes, in which intracellular Ca²⁺ homeostasis is chronically perturbed [90, 91], and TRPC6 hyperactivity elevates basal Ca²⁺ levels in kidney cells [92, 93, 94], UT–TRPC6 signaling may act as an amplifier of Ca²⁺ overload and downstream stress responses.

ER stress and Ca²⁺ homeostasis are intimately linked [95, 96, 97]. Depletion of ER Ca²⁺ stores and persistent cytosolic Ca²⁺ elevation activate the unfolded protein response, which, if unresolved, leads to apoptosis and fibrosis. Pharmacologic inhibition of ER stress has been shown to reduce albuminuria and glomerular injury in T2DM hypertensive rats [98]. We found that HFD increased ER stress markers in kidney tissue and plasma and that UT KO attenuated these changes in vivo. In vitro, UII increased ER stress markers and secretion of CRELD2 and DNAJB11 in podocytes, and these effects were reversed by UT or TRPC6 inhibition. These observations support a mechanistic link from UII–UT activation to TRPC6‐dependent Ca²⁺ signaling, ER stress, and podocyte structural injury.

Prior studies have reported variable renoprotective effects of TRPC6 deletion across diabetic models. In STZ‐treated Dahl salt‐sensitive rats, TRPC6 KO reduced nephrinuria, podocyte foot process effacement, and elevated basal intracellular Ca²⁺, but did not fully normalize hyperglycemia or albuminuria [92, 93]. In STZ‐treated Sprague–Dawley rats, TRPC6 inactivation did not ameliorate DKD [99]. In Akita mice, TRPC6 KO mitigated albuminuria and tubular injury at early time points, but not later, and hyperglycemia and fibrosis were largely unaffected [100]. These differences likely reflect species/strain variation, different diabetic stimuli (HFD vs. STZ vs. Ins2 mutation), and distinct durations and severities of hyperglycemia. In our long‐term HFD model on a C57BL/6J background with moderate but chronic hyperglycemia, UT deletion provided robust renoprotection over 33 weeks, suggesting that targeting a proximal Gq‐coupled receptor, such as UT, may provide broader benefit than targeting TRPC6 alone under some metabolic conditions.

Taken together, our data support a paradigm in which UT antagonism does not correct the primary metabolic abnormalities of diabetes (insulin deficiency, insulin resistance) but instead alleviates key aspects of DKD, including podocyte injury, tubular damage, albuminuria, and GBM injury by interrupting a maladaptive UT–Ca²⁺–ER stress pathway. These findings provide a mechanistic rationale for pharmacologic strategies targeting UT, as well as TRPC6, as a potential adjunctive therapy in DKD.

3.5. Limitations of This Study

This study used a global UT‐knockout model which cannot distinguish between kidney cell types responsible for a specific effect. Experiments reported here were limited to male mice and a single long‐term HFD model, warranting validation in female animals and other models of diabetic kidney injury. In addition, whole‐kidney lipidomics could not resolve the cellular origin or function of individual UT‐sensitive lipids. Future studies using cell‐specific UT deletion, cell‐specific lipidomics, genetic ablation of UT and TRPC6 in primary mouse and human podocytes, and selective UT antagonists should clarify the underlying mechanisms and therapeutic potential.

4. Conclusions

In summary, this study identifies UII–UT signaling as a central driver of podocyte injury, maladaptive renal lipid remodeling, and kidney damage in a high‐fat diet‐induced model of Type 2 diabetes. Genetic deletion of UT confers robust protection against DKD, preserving glomerular filtration barrier integrity, suppressing ER stress, and attenuating systemic, urinary, and tissue‐level markers of kidney injury, despite persistent systemic metabolic dysfunction. Pharmacologic inhibition of UII‐induced Ca²⁺ influx through TRPC6 attenuates ER stress and stabilizes the podocyte actin cytoskeleton. In parallel, UT deletion selectively mitigates lipotoxic remodeling of the renal lipidome, implicating UT signaling as a determinant of renal susceptibility to metabolic stress. Together, these findings define a UT–Ca²⁺–ER stress axis that mechanistically links UII signaling to podocyte dysfunction and highlight UT as a promising therapeutic target in obesity‐ and lipotoxicity‐driven DKD.

5. Materials and Methods

5.1. Animal Experiments

Wild‐type (WT) and urotensin II receptor knockout (UT KO; Uts2rtm1Djbe; Jackson Laboratories, stock no. 026012) mice were purchased and bred in‐house. Male mice aged 8–12 weeks were randomly assigned to receive either a high‐fat diet (HFD; 60% kcal fat; Research Diets, D12492) or a control low‐fat diet (LFD; 10% kcal fat; Research Diets, D12450B) for 33 weeks. Although our protocol at the onset was to feed mice with HFD for 21 weeks, this did not result in a significant increase in fasting blood glucose levels. Therefore, the duration of HFD feeding was increased to 33 weeks to allow for the development of hyperglycemia and diabetic kidney disease. The group fed with control LFD was also given an extended 33 weeks to control for any age‐related effects that might be observed. Randomization was performed at the time of diet assignment, and animals were housed under identical conditions throughout the study. At the conclusion of the feeding period, mice underwent necropsy to collect kidneys, plasma, and urine. All animal experiments were conducted at the University of Tennessee Health Science Center (UTHSC) and the University of Missouri (MU). Experimental protocols were approved by the Institutional Animal Care and Use Committees of UTHSC (protocol #16‐101.0) and MU (protocol #44428).

5.2. Cell Culture

SVI immortalized mouse podocytes (CLS Cell Lines Service GmbH, Germany) were cultured at 33°C as per the manufacturer's protocol and then transferred to 38°C for a minimum of 14 days for differentiation before experiments.

5.3. Intracellular Calcium Imaging

Mouse podocytes were cultured in 35 mm glass‐bottom culture dishes. On the day of the experiment, cells were loaded with a Ca²⁺‐sensitive dye Fura‐2‐acetoxymethyl ester (Fura‐2 AM; 10 μM) and 0.5% pluronic F‐127 for ∼1 h at room temperature in modified Krebs' buffer solution (MKBS: 134 mM NaCl, 6 mM KCl, 2.0 mM CaCl2, 2 mM MgCl2, 10 mM HEPES, and 10 mM glucose, pH 7.4). The cells were washed in MKBS and allowed to de‐esterify for 45 min; intracellular Ca²⁺ concentrations were then determined using a fluorescence photometry system (IonOptix Corp., Milton, MA, USA). Fura‐2 AM was excited at wavelengths of 340 and 380 nm, and background‐subtracted Fura‐2AM ratios were collected at 510 nm using a MyoCam‐S CCD digital camera. IonWizard software (IonOptix LLC) was used for acquisition and data analysis.

5.4. Patch‐Clamp Electrophysiology

Cation currents were recorded from cultured mouse podocytes in whole‐cell configuration. Patch pipettes were fabricated from borosilicate glass tubes using a P‐1000 Flaming/Brown micropipette puller (Sutter Instrument, Novato, CA, USA) and then fire‐polished using a microforge (Narishige, MF‐830) to achieve a resistance of 3–6 MΩ. Data were acquired using an Axopatch 200B patch‐clamp amplifier and pClamp 10 acquisition software (Molecular Devices, CA, USA) and digitized with an Axon Instruments Digidata 1550B analog‐to‐digital converter. Data were sampled at 10 kHz and filtered at 2 kHz (low‐pass Bessel filter). The membrane potential was held at 0 mV, and currents were recorded by applying 940‐ms voltage ramps from −100 to +100 mV. Cells were placed in a bath solution with the following composition: (in mM) NaCl 140, CsCl 5, MgCl2 2, CaCl2 2, HEPES 10, glucose 10, pH 7.4 (using 1 M NaOH). The pipette solution used had the following composition: (in mM) CsCl 140, MgCl2 2, HEPES 10, EGTA 2, glucose 10, Na‐ATP 3, pH 7.35 (adjusted with 1 M CsOH). All recordings were done with 70%–80% series resistance compensation.

5.5. Immunofluorescence

Cells cultured in coverslips were fixed with 4% formaldehyde (for 15 min) and permeabilized with 0.5% Triton X (for 15 min). The samples were incubated in 3% BSA to block nonspecific binding sites and incubated overnight at 4°C with primary antibodies. The next day, the samples were washed with PBS and incubated with secondary antibodies for 1 h at room temperature. Following the wash, the samples were mounted with ProLong Diamond Antifade Mountant with DAPI (Invitrogen) and sealed. Images were acquired using a Zeiss LSM 710 laser‐scanning confocal microscope or a Leica Microsystems Thunder imaging system.

Isolation of glomeruli from mouse kidneys was performed following a procedure described by Rush et al. [101]. Briefly, mouse kidneys were harvested and kept on ice. The kidneys were sliced longitudinally and passed through metal sieves with pore sizes of 190 μm and 104 μm. The tissues were mashed and sieved through 1% BSA in PBS. The flow‐through from the 190 μm and 104 μm sieves was then applied to a 40 μm nylon sieve. The glomeruli were collected from the top of the sieve. Glomeruli were then allowed to adhere to glass coverslips and processed for immunofluorescence imaging.

Formalin‐fixed, paraffin‐embedded kidney sections were deparaffinized in xylene and hydrated in decreasing concentrations of ethanol‐to‐water mixture. Antigen retrieval was performed in a steamer, with slides immersed in IHC antigen retrieval solution (Invitrogen, Cat. # 4955‐58, Life Technologies, Carlsbad, CA) at 90°C for 45 min. This was followed by permeabilization, antibody treatment, and coverslip mounting.

Super‐resolution confocal imaging was performed at the UTHSC Advanced Imaging Core using a ZEISS Elyra 7 Super‐Resolution Microscope (SRM Unit, Carl Zeiss Microscopy GmbH) equipped with a dual sCMOS camera. Actin filaments were imaged using the Lattice Structured Illumination Microscopy (SIM) method and processed with SIM Processing tools within the Zeiss Zen Black 3.0 software. Actin filaments were traced and their lengths measured (in arbitrary units) using FilamentSensor2.0, an open‐source software from Georg‐August‐University, Göttingen, Germany [102] (https://www.filament‐sensor.de/).

5.6. Antibodies

The following antibodies were used in immunofluorescence: anti‐BiP/GRP78 rabbit antibody (Cell signaling, Cat # C50B12), anti‐CHOP mouse antibody (Cell signaling, Cat # L63F7), anti‐IGFBP7 rabbit monoclonal antibody (Invitrogen, Cat # MA5‐29346), anti‐TIMP2 mouse monoclonal antibody (Abcam, Cat # Ab230511), anti‐nephrin mouse antibody (Biosynth, Cat # 20R‐NP002), anti‐podocin mouse antibody (Sigma, Cat # 4200810), anti‐UT rabbit antibody (Alomone labs, AER‐003), and ActinGreen 488 ReadyProbes reagent (Invitrogen, Cat # R37110). The following fluorescent‐labeled secondary antibodies were purchased from Biotium Inc., Fremont, CA: Highly cross‐adsorbed CF555 donkey anti‐rabbit IgG (# 20038), CF488 donkey anti‐rabbit IgG (# 20015), CF555 donkey anti‐mouse IgG (# 20037), and CF488 donkey anti‐mouse IgG (# 20014).

5.7. Polymerase Chain Reaction and RT‐PCR

Total RNA was extracted from kidney tissues or cultured cells by homogenizing in TRIzol Reagent (Cat. # R2050, Zymo Research) and then processed using a commercially available RNA preparation kit (Direct‐zol RNA MiniPrep Plus Kits, Zymo Research). The total RNA was then reverse‐transcribed using the High‐Capacity cDNA Reverse Transcription Kit (Applied Biosystems, Thermo Fisher Scientific, Waltham, MA).

PCR reactions were performed using an Applied Biosystems MiniAmp thermal cycler (Thermo Fisher Scientific, Waltham, MA), and the resultant products were run on a 1.5% agarose gel with GelRed nucleic acid stain to resolve the amplicons. The gel was then imaged using a Bio‐Rad ChemiDoc MP Imaging System (Bio‐Rad Laboratories Inc., Hercules, CA).

RT‐PCR reactions were performed in triplicate using an Applied Biosystems QuantStudio 5 system (Life Technologies, Carlsbad, CA). All genes of interest were normalized to β‐actin expression. Relative expression of gene transcripts was determined using 2^−ΔΔCt, where the Ct values of the gene of interest were first normalized to β‐actin and then to the ΔCt of the control LFD group.

Table 2 shows a list of oligonucleotide primer pairs used in this study.

TABLE 2.

Oligonucleotide primer sequences.

Gene	Sequence	Accession number	Product size (bp)
Urotensin 2 (UII)	Forward: GCCTGCTCTTCATAGGACTTC Reverse: CGTCTTCCTCAAGCACTGGAA	NM_011910.2	96
Urotensin 2 receptor (UT)	Forward: AGCATCTTCACCCTGACCAT Reverse: CAGCTTACGGTAACCCTTGG	NM_145440.1	96
UT	Forward: ATGGGGCCTTTGTGTGAGAG Reverse: AGGCCCCTTACCAACCAATG	NM_145440.1	378
UT (Nested)	Forward: TTGTGGAAAGTCCCCTCTTCT Reverse: GGGCTTACATCAAAGGTCAGG	NM_145440.1	214
WT‐1	Forward: GTTTTCTCGCTCAGACCAGC Reverse: AGTTTGCACACTTTCCTGCC	NM_144783.2	314
TRPC5	Forward: AACTCCCTCTACCTGGCAAC Reverse: GCTTCTGCAATCAGAGTCGG	NM_009428.3	113
TRPC6	Forward: TGTGGATATGCCAAGCTGGA Reverse: ACATCCAAAGGGCCAGGATT	BC068310.1	109
GRP94	Forward: AAGAATGAAGGAAAAACAGGACAAAA Reverse: CAAATGGAGAAGATTCCGCC	NM_011631.1	77
β‐Actin	Forward: CTTCCAGCAGATGTGGATCAG Reverse: AACGCAGCTCAGTAACAGTCC	NM_007393.5	99

5.8. Enzyme‐Linked Immunosorbent Assay (ELISA)

ELISAs of plasma, urine, and cell culture media were done using the following assay kits: mouse insulin (Cat. no: 90080, Crystal Chem, Elk Grove Village, IL), mouse glucagon (Cat. no: 81518, Crystal Chem, Elk Grove Village, IL), mouse albumin (Cat. no: E‐90AL, Immunology Consultants Laboratory Inc., Portland, OR), mouse CRELD2 (Cat. no: ELM‐CRELD2, RayBiotech Life Inc., Peachtree Corners, GA), mouse DNAJB11 (Cat. no: abx510480, Abbexa LLC, Houston, TX), mouse NGAL/lipocalin‐2 (Cat. no: ab199083, Abcam, Waltham, MA), mouse Cystatin C (Cat. no: 80738, Crystal Chem, Elk Grove Village, IL), mouse RBP4 (Cat. no: ab202404, Abcam, Waltham, MA), and mouse alpha 1‐microglobulin (Cat. no: MBS265422, MyBioSource Inc., San Diego, CA). All ELISAs were performed according to the manufacturer's protocols.

5.9. Transmission Electron Microscopy

Kidney tissue was collected and processed for transmission electron microscopy (TEM). Unless otherwise stated, all reagents were purchased from Electron Microscopy Sciences, and all specimen preparation was performed at the Electron Microscopy Core Facility, University of Missouri. The samples were fixed in 2% paraformaldehyde and 2% glutaraldehyde in 100 mM sodium cacodylate buffer, pH 7.35. Tissues were rinsed with 100 mM sodium cacodylate buffer, pH 7.35 (Sigma‐Aldrich, St. Louis, MO), and 130 mM sucrose. Secondary fixation was performed using 1% osmium tetroxide (Ted Pella Inc., Redding, California) in cacodylate buffer. Specimens were incubated at 4°C for 1 h, rinsed with cacodylate buffer, and then with distilled water. En bloc staining was performed with 1% aqueous uranyl acetate at 4°C overnight, followed by rinsing with distilled water. A graded dehydration series was performed using ethanol, followed by a transition into acetone. The dehydrated tissues were then infiltrated with EMbed 812 resin and polymerized at 60°C overnight. Semithin sections were collected at a thickness of 1 μm and stained with Toluidine blue to locate the region of interest. The block face was additionally trimmed for the area of interest, and sections were cut to a thickness of 75 nm using an ultramicrotome (Ultracut UCT, Leica Microsystems, Germany) and a diamond knife (Diatome, Hatfield, PA). Images were acquired using a JEOL JEM 1400 transmission electron microscope (JEOL, Peabody, MA) at 80 kV with a Gatan Rio CMOS camera (Gatan Inc., Pleasanton, CA).

5.10. Immunohistochemistry and Pathological Scoring

Tissue processing and immunohistochemistry were performed independently by iHisto (Salem, MA). Periodic acid–Schiff (PAS)—a board‐certified pathologist at iHisto evaluated stained kidney tissue sections. Histological alterations were graded for severity using a standardized scoring system: 0 = no significant change; 1 = minimal; 2 = mild; 3 = moderate; and 4 = severe. All assessments were conducted in accordance with the International Harmonization of Nomenclature and Diagnostic Criteria [103].

5.11. Lipidomics

Untargeted lipidomic profiling was performed by Metabolon (Morrisville, NC) using frozen kidney tissues collected at necropsy. Sample preparation, data acquisition, and compound identification were conducted by Metabolon using established platforms. Data processing and statistical analyses, including differential abundance testing and generation of volcano plots, were performed by Metabolon's data analysis team.

5.12. Data and Statistical Analysis

Data are expressed as mean ± standard error of the mean (SEM). GraphPad Prism (Sacramento, CA), Origin (OriginLab, Northampton, MA), NIH open‐source software ImageJ/Fiji, Image Lab v6 (Bio‐Rad Laboratories Inc., Hercules, CA), IonWizard software (IonOptix LLC, Westwood, MA), and pClamp 10 (Molecular Devices, San Jose, CA) were used for data analysis, data representation, and statistical analysis. Comparisons among groups were analyzed using one‐way ANOVA with Tukey's post hoc test. A p‐value less than 0.05 was considered significant.

Author Contributions

Praghalathan Kanthakumar: conceptualization, Methodology, Validation, Formal analysis, Investigation, Data Curation, Writing – Original Draft. Adebowale Adebiyi: conceptualization, Methodology, Validation, Writing – Review and Editing, Supervision, Funding acquisition.

Conflicts of Interest

The authors declare no conflicts of interest.

Data Availability Statement

Data will be made available on request.