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. 2024 Jun 19;300(8):107485. doi: 10.1016/j.jbc.2024.107485

WWC1 upregulation accelerates hyperuricemia by reduction in renal uric acid excretion through Hippo signaling pathway

Changshun Han 1,‡, Chengyong He 1,‡, Xiaoyan Ding 1, Zixuan Li 1, Tianyun Peng 1, Chensong Zhang 1, Haibing Chen 2, Zhenghong Zuo 1, Jiyi Huang 1,∗, Weiping Hu 1,∗
PMCID: PMC11301351  PMID: 38906255

Abstract

Hyperuricemia (HUA) is a metabolic disorder characterized by elevated serum uric acid (UA), primarily attributed to the hepatic overproduction and renal underexcretion of UA. Despite the elucidation of molecular pathways associated with this underexcretion, the etiology of HUA remains largely unknown. In our study, using by Uox knockout rats, HUA mouse, and cell line models, we discovered that the increased WWC1 levels were associated with decreased renal UA excretion. Additionally, using knockdown and overexpression approaches, we found that WWC1 inhibited UA excretion in renal tubular epithelial cells. Mechanistically, WWC1 activated the Hippo pathway, leading to phosphorylation and subsequent degradation of the downstream transcription factor YAP1, thereby impairing the ABCG2 and OAT3 expression through transcriptional regulation. Consequently, this reduction led to a decrease in UA excretion in renal tubular epithelial cells. In conclusion, our study has elucidated the role of upregulated WWC1 in renal tubular epithelial cells inhibiting the excretion of UA in the kidneys and causing HUA.

Keywords: hyperuricemia, renal tubule, UA excretion, Hippo signaling pathway, WWC1

Hyperuricemia (HUA) is a metabolic disorder characterized by high levels of uric acid (UA) in the serum. There has been a consistent upward trend in the global prevalence of this condition (1, 2). HUA has been recognized as a major global public health issue (3). The estimated occurrence of HUA among Chinese adults was 14.0% (4). Approximately 14.6% of the US population has HUA (5). HUA plays a pivotal role in the development and progression of gout, and it is also linked to increased risks of chronic kidney disease, end-stage kidney disease, cardiovascular events, and mortality (6, 7).

UA is the ultimate product of purine nucleotide metabolism in humans, and it is predominantly synthesized in the liver (8). In humans, the excretion of serum UA is mainly performed by the kidneys (two-thirds) and to a lesser extent by the gut (one-third) (9, 10). Upon entering the bloodstream, UA undergoes free filtration at the glomerulus, predominantly reabsorption and secretion in the S1-S2 segments of the proximal tubule, with potential reabsorption in the late proximal tubule (S3)—a topic that remains under debate (11, 12). This intricate process ultimately results in the reabsorption of approximately 90% of the filtered UA (13, 14).

HUA is a complex condition influenced by both genetic and environmental factors (15). While the exact cause of HUA remains uncertain in numerous cases, it is widely believed that disruption in gene expression related to UA production in the liver and its excretion in the kidneys plays a significant role (16). The increased production of UA in the liver due to the upregulation of the key enzyme xanthine oxidase (XOD), as well as the decreased excretion of UA in the kidneys caused by the downregulation of transport proteins like organic anion transporter 3 (OAT3) and ATP-binding cassette sub-family G member 2 (ABCG2), can contribute to elevated serum UA levels (17, 18). The dysregulation of other associated proteins is also linked to the initiation and progression of HUA. Changes in enzyme activity resulting from increased hypoxanthine guanine phosphoribosyl transferase levels and decreased phosphoribosyl pyrophosphate synthase 1 levels can also lead to increased UA production in the body (19).

The Hippo signaling pathway is crucial in regulating cellular proliferation, maintaining tissue equilibrium, and impacting organ dimensions (20, 21). WWC1 (WW and C2 domain containing 1), a protein prominently associated with the kidney and brain, is predominantly expressed in both the brain and kidney (22). WWC1 exhibits expression within various components of the kidney, including glomerular podocytes, tubular structures, and collecting ducts (23). The protein WWC1 is an established element of the Hippo pathway (24). Specifically, WWC1 has been documented to interact with LATS1 (large tumor suppressor 1) and SAV1 (salvador homolog 1), thereby facilitating LATS1 phosphorylation (25). Consequently, the phosphorylated LATS1 triggers the subsequent inactivation of YAP1 (Yes-associated protein 1) via phosphorylation (26). YAP1, a potent transcriptional co-activator, plays crucial roles in development, stem cell maintenance, and maintaining normal tissue homeostasis and regeneration (27, 28). The protein levels of WWC1 are implicated as potential modulators of Hippo signaling (29). Moreover, there is supporting evidence suggesting that the activation of the Hippo signaling pathway by WWC1 leads to podocyte injury in vitro (30). However, the biological functions of WWC1 in kidney excretion of UA remain to be fully elucidated.

Our study utilized rat, mouse, and cell models to conduct animal and cell experiments, revealing the significant role of WWC1 in the onset and progression of HUA. Additionally, our findings demonstrated that the upregulation of WWC1 in renal tubular epithelial cells led to the phosphorylation and subsequent inactivation of the transcription factor YAP1 via the Hippo pathway. Consequently, this inhibited YAP1 from entering the cell nucleus and the transcription of ABCG2, thereby suppressing UA excretion, and leading to elevated levels of UA in the body. Our findings offered valuable insights into the intricate mechanisms that underlie the pathogenesis and progression of HUA.

Result

Uox knockout rats exhibited a reduction in renal UA excretion concurrent with an elevation in renal WWC1 levels

We developed a rat model with a deletion mutation of urate oxidase (Uox), denoted as Uox−/−, to serve as an appropriate model for HUA that more closely mimics purine metabolism in humans (Fig. 1A). The Uox gene in rats comprises eight exons. To facilitate allele discrimination during Cas9 cleavage, we designed a guide RNA (gRNA) TRGT specifically targeting a region within exon 3. Samples from Uox−/− and Uox+/+ rats exhibited distinct single peaks in Sanger sequencing (Fig. S1A). After aligning with Uox+/+ rat sequences, a 5 bp deletion was discovered in the Uox gene of Uox−/− rats (Fig. S1B). Samples of serum and tissue were collected from 6-month-old Uox−/− and Uox+/+ rats for analysis. The kidney somatic index of the Uox−/− group was comparably assessed against the wild-type (Uox+/+) group. No statistically significant differences were observed between the Uox−/− and Uox+/+ groups (Fig. 1B). The Uox−/− group exhibited noteworthy elevations in serum levels of creatinine (CREA), blood urea nitrogen (BUN), and UA (Fig. 1, C–E). Serum XOD levels, a key enzyme in UA metabolism and closely associated with UA production, were significantly higher in the Uox−/− group compared to those in the Uox+/+ group (Fig. 1F). UA is a crucial indicator of HUA. The liver synthesizes UA, which is then excreted through the kidneys. Inadequate excretion of UA can contribute to kidney damage. The renal tissues in Uox+/+ group exhibited a normal morphology, while Uox−/− group showed characteristic histological alterations, including renal tubule swelling (Fig. 1, G and H). To assess changes in renal UA transporters, we examined the UA excretion proteins OAT3 and ABCG2. The Uox−/− group showed reduced levels of OAT3 and ABCG2 proteins compared to the Uox+/+ group (Figs. 1I and S1C). The Uox−/− group exhibited elevated levels of both mRNA and protein of WWC1 (Figs. 1, J and K and S1D). Furthermore, immunohistochemical staining revealed the presence of WWC1 in renal tubules, with a significantly elevated expression level observed in Uox−/− rats. (Fig. S1E). In summary, we observed a reduction in UA excretion and an enhancement in renal WWC1 expression in Uox−/− rats.

Figure 1.

Figure 1

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Uox deficiency rat model revealed elevated serum UA and renal expression of WWC1.A, Western blotting analysis of hepatic expression of UOX in Uox+/+ and Uox−/− rats. B, kidney somatic index comparison between Uox+/+ and Uox−/− rats. C–F, comparison of serum CRE (B), BUN (C) UA (D), and XOD (E) levels in Uox+/+ and Uox−/− rats. G and H, images of H&E staining in kidney sections from Uox+/+ and Uox−/− rats (G) with quantification of relative kidney tubule area (H). G: glomerulus; T: renal tubule. Scale bar: 200 μm and 100 μm. I, Western blotting analysis of renal expression of UA transporters (OAT3 and ABCG2) in Uox+/+ and Uox−/− rats. J, quantification of WWC1 mRNA expression assessed by RT–qPCR in kidneys from Uox+/+ and Uox−/− rats. K, analysis of renal expression of UA transporters OAT3 and ABCG2 using Western blotting in Uox+/+ and Uox−/− rats. GAPDH was identified as the loading control. The data presented here were representative of a minimum of three independent experiments. Values were expressed as the mean ± SD. ∗p < 0.05, ∗∗p < 0.01, ∗∗∗p < 0.001.

The renal excretion of UA was reduced, accompanied by an elevation in the level of renal WWC1 in an HUA mouse model

To induce a HUA mouse model, we administered intraperitoneal injections of PO (potassium oxonate) at a dosage of 350 mg/kg and provided the mice with 10% fructose water for 28 days (Fig. 2A). Mice in the HUA model group showed a statistically significant increase in kidney somatic index compared to the control group (Fig. 2B). As depicted in Figure 2E, PO treatment led to a marked increase in serum UA levels when compared to the control group. We observed a substantial increase in serum CREA and XOD levels in the HUA model group (Fig. 2, C and F). Based on histological examination, the HUA model group exhibited significant tubular dilation and hydropic degeneration of renal tubules compared to the control group (Fig. 2, G and H).

Figure 2.

Figure 2

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The renal WWC1 level was increased in an HUA mouse model.A, schematic illustration depicting the generation of the HUA mice model using PO and fructose water. B, comparison of kidney somatic index between control and HUA mice. C–F, evaluation of serum CRE (C), BUN (D), UA (E), and XOD (E) levels in control and HUA mice. G and H, images of H&E staining in kidney sections from control and HUA mice (E) with quantification of the relative kidney tubule area (G). G: glomerulus; T: renal tubule. Scale bar: 200 μm and 100 μm. I, analysis of renal expression of UA transporters OAT3 and ABCG2 using Western blotting in both control and HUA mice. J, quantification of Wwc1 mRNA expression assessed by RT-qPCR in kidneys from control and HUA mice. K, western blotting analysis depicting the renal expression of WWC1 in both control and HUA mice. GAPDH was identified as the loading control. The data presented here were representative of a minimum of three independent experiments. Values were expressed as the mean ± SD. ∗p < 0.05, ∗∗p < 0.01.

To characterize changes in renal UA transporters, we examined the renal proteins OAT3 and ABCG2. The HUA model group exhibited reduced expression of OAT3 and ABCG2 proteins compared to the control group (Figs. 2I and S2A). Regarding Wwc1 mRNA, the HUA model group showed elevated Wwc1 mRNA expressions compared with the control (Fig. 2J). Similarly, HUA mouse models increased renal WWC1 protein levels in the kidney (Figs. 2K and S2B). In combination, mice with HUA exhibited decreased renal UA excretion and increased renal WWC1 expression.

WWC1 expression was elevated in an HUA cell model

We created a cellular model of HUA in HK2 (human renal cortex proximal tubule epithelial cells) cells by subjecting them to adenosine treatment (2.5 mM) for 30 h, followed by XOD exposure (2.5 U/L) for 8 h. A schematic representation of the experimental protocol employed to treat HK2 cells with these inducers is presented in Figure 3A. The HUA model showed a significant increase in extracellular UA concentration compared to the control group (Fig. 3B). To characterize transporter changes, we conducted tests on the UA excretion proteins ABCG2 and OAT3. The HUA model cells exhibited reduced levels of OAT3 and ABCG2 proteins compared to the control (Figs. 3C and S3A). Regarding WWC1 mRNAs, the HUA model cells displayed elevated WWC1 mRNA expressions compared to the control (Fig. 3D). In line with the mRNA results, the protein level of WWC1 was elevated in the cells of the HUA model (Figs. 3E and S3B). Furthermore, immunofluorescence analysis demonstrated an elevated cytoplasmic expression of WWC1 in cells of the HUA model (Fig. 3F). The results demonstrated that the upregulation of WWC1 in HK2 cells of HUA was associated with a decrease in cellular UA excretion. In short, these findings provided preliminary evidence suggesting that elevated WWC1 levels led to reduced UA excretion in renal tubular epithelial cells, providing novel insights into the mechanisms underlying UA regulation.

Figure 3.

Figure 3

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The expression of WWC1 was upregulated in a cellular model of HUA.A, schematic illustration depicting the generation of the HUA HK2 cell model using inducers. B, comparison of extracellular UA levels between control and HUA model cells. C, analysis of expression of UA transporters OAT3 and ABCG2 using Western blotting in control and HUA model cells. D, quantification of WWC1 mRNA expression via RT-qPCR in control and HUA model cells. E, evaluation of expression of WWC1 through Western blotting in control and HUA model cells. GAPDH was identified as the loading control. F, immunofluorescence images depicting WWC1 in control and HUA model cells. Scale bar: 10 μm and 5 μm, respectively. The data presented here were representative of a minimum of three independent experiments. Values were expressed as the mean ± SD. ∗p < 0.05, ∗∗∗∗p < 0.0001.

WWC1 knockdown enhanced the UA secretion in HK2 cells

We silenced WWC1 in HK2 cells using small interfering RNAs (siRNAs) (Figs. 4, A and B and S4, A and B). Upon WWC1 knockdown, we observed a significant increase in extracellular UA levels accompanied by a corresponding decrease in intracellular UA levels (Fig. 4, C and D). This suggested that the level of UA excretion was elevated in WWC1 knockdown HK2 cells. To characterize UA transporter changes, we examined the levels of OAT3 and ABCG2 proteins. Silencing WWC1 resulted in a noteworthy elevation of OAT3 and ABCG2 protein levels in HK2 cells (Figs. 4E and S4C). These results indicated that WWC1 knockdown promotes UA secretion.

Figure 4.

Figure 4

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Knockdown of WWC1 promoted UA secretion in HK2 cells.A, quantification of WWC1 mRNA expression via RT-qPCR in si-NC and si-WWC1 HK2 cells. B, analysis of expression of WWC1 using Western blotting in HK2 cells after WWC1 siRNA treatment. C and D, extracellular (C) and intracellular (D) UA levels in si-NC and si-WWC1 HK2 cells. E, analysis of expression of UA transporters OAT3 and ABCG2 using Western blotting in si-NC and si-WWC1 HK2 cells. GAPDH was identified as the loading control. The data presented here were representative of a minimum of three independent experiments. Values were expressed as the mean ± SD. ∗p < 0.05.

The overexpression of WWC1 in HK2 cells resulted in the inhibition of UA secretion

Following the transfection of the WWC1 plasmid to induce WWC1 overexpression in HK2 cells, both WWC1 mRNA and protein levels were significantly increased (Figs. 5, A and B and S5A). Elevated WWC1 expression led to a notable reduction in extracellular UA levels and a concurrent increase in intracellular UA levels, indicating a potential decrease in UA excretion in WWC1 overexpressed HK2 cells (Fig. 5, C and D). To characterize alterations in UA transporters, we assessed the levels of OAT3 and ABCG2 proteins in HK2 cells. WWC1 overexpression led to a notable decrease in endogenous OAT3 and ABCG2 protein levels (Figs. 5F and S5B). These results indicated that the increase of WWC1 inhibited the excretion of UA in HK2 cells.

Figure 5.

Figure 5

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Overexpression of WWC1 decreased UA secretion in HK2 cells.A, quantification of WWC1 mRNA expression via RT-qPCR in HK2 cells following WWC1 plasmid transfection. B, analysis of expression of WWC1 using Western blotting in HK2 cells after WWC1 plasmid transfection. C and D, extracellular (C) and intracellular (D) UA levels after WWC1 plasmid transfection. E, analysis of expression of UA transporters OAT3 and ABCG2 using Western blotting in HK2 cells after WWC1 plasmid transfection. GAPDH was identified as the loading control. The data presented here were representative of a minimum of three independent experiments. Values were expressed as the mean ± SD. ∗p < 0.05, ∗∗p < 0.01.

WWC1 decreased the transcription of UA excretion transporter OAT3 and ABCG2 by suppressing YAP1 activity in the renal tubular epithelial cells

We assessed the protein levels of LATS1, p-LATS1, YAP1 and p-YAP1 in the kidneys of both rats and mice. The Uox−/− group demonstrated a significant increase in p-LAST1/LATS1 protein levels compared to the Uox+/+ group, whereas the levels of total YAP1 and YAP1/p-YAP1 were significantly decreased (Figs. 6A and S6A). The HUA group exhibited a significant increase in p-LAST1/LATS1 protein levels in mouse kidneys, accompanied by a noteworthy decrease in total YAP1 and YAP1/p-YAP1 levels (Figs. 6B and S6B).

Figure 6.

Figure 6

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The upregulation of WWC1 inhibited the activity of YAP1, subsequently suppressing both the transcription of UA excretion proteins.A, analysis of renal expression of LATS1, p-LATS1, YAP1, and p-YAP1 using Western blotting in Uox+/+ and Uox−/− rats. B, analysis of renal expression of LATS1, p-LATS1, YAP1, and p-YAP1 using Western blotting in both control and HUA mice. C, analysis of expression of LATS1, p-LATS1, YAP1 and p-YAP1 using Western blotting in HK2 cells after WWC1 siRNA treatment. D, analysis of expression of LATS1, p-LATS1, YAP1 and p-YAP1 using Western blotting in HK2 cells WWC1 plasmid transfection. GAPDH was identified as the loading control. E, quantification of SLC22A8 and ABCG2 mRNA expression via RT-qPCR in HK2 cells after WWC1 siRNA treatment. F, quantification of SLC22A8 and ABCG2 mRNA expression via RT-qPCR in HK2 cells after WWC1 plasmid transfection. G, analysis of renal expression of YAP1 in the cytoplasm and nucleus using Western blotting in HK2 cells after WWC1 plasmid transfection. GAPDH was identified as the loading control of cytoplasm, LAMINB served as a loading control of the nucleus. H and I, extracellular (H) and intracellular (I) levels of UA after co-transfection of WWC1 and YAP1 (or YAP5SA) plasmids. J, analysis of expression of UA transporters OAT3 and ABCG2 using Western blotting in HK2 cells after co-transfection of WWC1 and YAP1 (or YAP5SA) plasmids. GAPDH was identified as the loading control. K and L, quantification of SLC22A8 (K) and ABCG2 (L) mRNA expression via RT-qPCR in HK2 cells after co-transfection of WWC1 and YAP1 (or YAP5SA) plasmids. M, this schematic diagram illustrated a potential mechanism by which WWC1 affects the levels of renal UA excretion protein. The data presented here were representative of a minimum of three independent experiments. Values were expressed as the mean ± SD. ∗p < 0.05, ∗∗p < 0.01.

Compared to si-NC HK2 cells, si-WWC1 cells exhibited decreased levels of phosphorylated LATS1, accompanied by an increase in total YAP1 levels. The notably elevated YAP/p-YAP ratio in si-WWC1 cells, compared to si-NC cells, suggested heightened YAP1 activity in the si-WWC1 cells (Figs. 6C and S6C). Conversely, overexpressed WWC1 HK2 cells exhibited higher levels of phosphorylated LATS1 and lower total YAP1 levels. The significantly reduced YAP/p-YAP ratio in WWC1-overexpressed cells suggested diminished YAP1 activity in these cells (Figs. 6D and S6D).

Following WWC1 knockdown, the results revealed significantly increased levels of SLC22A8 (encoding OAT3 protein) and ABCG2 mRNA in WWC1 knockdown HK2 cells (Fig. 6E). Conversely, overexpression of WWC1 resulted in a marked reduction in SLC22A8 and ABCG2 mRNA levels (Fig. 6F). Cellular cytoplasmic and nuclear proteins were extracted from both control and overexpressing WWC1 HK2 cells. YAP1 was significantly decreased in both cytoplasm and nucleus, suggesting that the level of transcription factor YAP1 was decreased and the amount entering the nucleus was reduced after WWC1 was highly expressed (Fig. 6G).

We performed YAP1 and YAP5SA (constitutively expressed YAP1 protein) overexpression rescue experiments on HK2 cells overexpressing WWC1. The results showed a significant increase in extracellular UA levels following elevation of YAP1/YAP5SA levels (Fig. 6H), accompanied by a significant decrease in intracellular uric acid levels (Fig. 6I). With the overexpression of YAP1/YAP5SA, the levels of UA excretion proteins OAT3/ABCG2 significantly increased in HK2 cells (Figs. 6J and S6, F–H). Similarly, in cells overexpressing YAP1/YAP5SA, mRNA levels of SLC22A8 and ABCG2 also showed a significant increase (Fig. 6, K and L). This indicated that YAP1, acting as a transcription factor, could promote the expression of UA excretion proteins OAT3 and ABCG2. Taken together, our data indicated that elevated WWC1 levels indirectly reduced the transcription of UA transporters encode genes (SLC22A8 and ABCG2) in renal tubular epithelial cells by inhibiting the activity of transcription factor YAP1 through the activation of the Hippo pathway (Fig. 6M).

Discussion

The homeostasis of UA depends on a balance of complex processes involving hepatic production and renal tubule/intestinal excretion (31). The kidneys play a crucial role in excreting UA, accounting for approximately two-thirds of its total excretion (32, 33). WWC1 is expressed in multiple tissues, exhibiting greater expression levels in both human and mouse kidneys (34). Nevertheless, limited knowledge exists regarding the functional role of WWC1 in kidney tubule physiology and its involvement in the pathogenesis of HUA.

The function of UA transporters in renal tubule epithelial cells is pivotal for maintaining UA homeostasis in the human body (35, 36). ABCG2 acts as a multi-drug efflux pump, functioning as a UA efflux transporter on the apical membrane of proximal renal tubule cells (37, 38). On the other hand, OAT3, encoded by SLC22A8, mediates the secretion of UA from the basolateral side to the intracellular proximal renal tubule cells, ultimately facilitating its excretion in the urine (39, 40).

During purine metabolism in most mammalian species, UA is synthesized and subsequently degraded by the enzyme uricase. This process leads to the formation of allantoin, which is more soluble. Nevertheless, due to the absence of functional uricase in humans and the significant reabsorption of filtered UA, UA levels in human plasma are roughly tenfold higher than those found in the majority of other mammals (41, 42). Owing to their size and cognitive and physiological traits, rats serve as a valuable model for exploring cardiovascular diseases, neurological disorders, and metabolic disorders (43). We generated HUA rat models with Uox deletion mutations (Uox−/−) to simulate human UA metabolism and induce HUA. In the Uox−/− group, the renal UA efflux transporters, specifically ABCG2 and OAT3, exhibited a significant decrease in expression levels compared to the Uox+/+ group. And the Uox−/− group exhibited elevated expression levels of WWC1. This finding indicated that elevated levels of renal protein WWC1 are linked to decreased renal UA excretion.

In comparison to reports on Uox knockout mice, our observations reveal that although serum UA levels significantly increase in Uox−/− rats, the magnitude of the increase is smaller compared to Uox−/− mice. Evaluation through serum CREA and kidney histology indicated lesser renal damage in Uox−/− rats. Additionally, Uox knockout did not adversely affect the survival of rats during the experimental process (44). It is speculated that this difference may be attributed to variances between species, such as differences in size and lifestyle habits between rats and mice, which could result in rats exhibiting greater tolerance to the UA elevation caused by Uox gene knockout.

Several studies have demonstrated the inhibitory potential of PO on uricase activity (45). Therefore, we utilized PO to induce HUA in mice. Consistent with prior research, the mice exhibited a significant elevation in UA levels following drug-induced administration (46). Similar to the rats experiment, mice with HUA had decreased levels of UA excretion proteins OAT3 and ABCG2 in their kidneys, while the WWC1 level was elevated. Human proximal tubule epithelial cells encompass a diverse range of UA transporters and serve as the primary site for renal tubule UA excretion (47, 48). Consistently, in HUA HK2 cells induced by adenosine and XOD, we observed decreased expression of UA excretion proteins OAT3 and ABCG2, along with increased levels of WWC1. Furthermore, elevated WWC1 levels in HK2 cells have been found to impede the excretion of UA.

As a transcription factor, YAP1 has been shown to regulate ABCG2 by binding to its promoter and modulating its transcription (49). Similarly, YAP1 facilitates the expression of renal collecting duct proteins AQP-2, -3, and -4 to maintain water homeostasis in vivo (50). Functioning as an upstream regulator of the Hippo pathway, WWC1 activates it, thereby promoting the phosphorylation-dependent degradation of YAP1 (51). In line with previous studies, our animal and cell experiments revealed that elevated levels of WWC1 promote LATS1 phosphorylation, thereby inhibiting YAP1 activity. The reduction in YAP1 levels resulted in decreased transcription and expression of UA excretion proteins OAT3 and ABCG2 in the proximal renal tubules, consequently leading to reduced renal UA excretion.

Overexpression of YAP1 could increase the levels of uric acid excretion proteins, promoting uric acid excretion. Phosphorylation of YAP1 is crucial for Hippo pathway signal transduction (52), and a YAP1 mutant with serine substitutions (Mt-YAP5SA) at the LATS1 kinase recognition site can resist degradation (53). Our experiments demonstrated that YAP5SA was constitutively overexpressed in cells, with significantly higher levels than wild-type YPA1 overexpression. However, compared to YPA1, YAP5SA did not significantly enhance uric acid excretion. It was hypothesized that overexpression of YAP1 saturated the transcription and translation levels of UA excretion proteins, and although YAP5SA had higher cellular levels, it still could not effectively promote UA excretion.

In summary, our research revealed a pivotal involvement of WWC1 in the onset and advancement of HUA. Our investigation revealed that elevated levels of WWC1 reduced OAT3 and ABCG2 expression by suppressing YAP1 activity in the kidney tubule. Elevated WWC1 levels inhibited the expression of UA excretion proteins (OAT3/ABCG2) in renal tubular epithelial cells, resulting in suppressed renal UA excretion. Collectively, this study illustrated that the increased expression of WWC1 contributed to the onset and advancement of HUA. These findings emphasized the crucial roles played by WWC1 and YAP1 in regulating UA excretion. Our study enhanced our understanding of HUA and provided important insights into the effects and mechanisms of UA regulation. Furthermore, it provided fresh perspectives for devising strategies aimed at preventing and treating HUA.

Experimental procedures

Rats and treatment

The Uox knockout (Uox−/−) rats were kindly provided by Haibing Chen, from the Department of Endocrinology and Metabolism, Shanghai 10th People's Hospital, School of Medicine, Tongji University, Shanghai, China. The CRISPR-Cas9 system was utilized as a targeted approach to edit the Uox gene in rats. A guide RNA (gRNA) was designed to specifically target exons 3 of the Uox gene, and a corresponding targeting vector was constructed. Rat zygotes were microinjected with a mixture of transcribed Cas9 and gRNA. Genomic DNA was extracted from 7-day-old rat tail tissue. Genomic DNA was extracted by dissolving each rat tail in 400 μl of SNET lysis buffer (100 mM Tris-Cl, 100 mM NaCl, 25 mM EDTA, 0.5% SDS, 0.1 mg/ml proteinase K) overnight at 55 °C. The genomic DNA was extracted from the lysis mixture by adding 3 volumes of ethanol, followed by dissolution in 150 μl of double-distilled H2O at 55 °C for a minimum duration of 4 h. Genotyping of the modified rats was performed using the following PCR primers: Uox-forward primer: 5′-CCCAGGCTAAACTCTGAGGCT-3′; reverse primer: 5′-TGTCAGGGAAACAGTCATTTCACA-3′. PCR reactions were conducted in a Veriti 96-well Thermal Cycler from Applied Biosystems (BIO-RAD) using Multiplex PCR Mix (Sangon Biotech), under the following conditions: initialization at 93 °C for 3 min; denaturation for 35 cycles at 95 °C for 30 s, annealing at 62 °C for 30 s, and extension at 72 °C for 30 s; and final extension at 72 °C for 10 min. The PCR products were then subjected to sequencing analysis. Five wild-type (Uox+/+) and five Uox−/− rats, utilized in this study, were accommodated at the Laboratory Animal Center, Xiamen University, with unrestricted access to food and water. On the sixth month, rats were sacrificed after a 12 h fast. Blood samples were collected using 1.5 ml centrifuge tubes, and then centrifuged at 3500 rpm for 15 min at 4 °C to collect serum. The samples were stored at −80 °C for subsequent biochemical assays. The kidney tissues were promptly excised, weighed, and divided into two sections: one portion was fixed in 4% paraformaldehyde for histopathological examination, while the other was stored at −80 °C for subsequent biochemical assays.

All animal experiments adhered to ethical standards. All animal experiments were approved by the Xiamen University Institutional Animal Ethics Committee (Acceptance No. XMULAC20170361).

Mice and treatment

All mice in this study were of the Kunming genetic background. The experimental cohort exclusively comprised male mice. The age range of the mice spanned from 6 to 8 weeks. Experimental mice were sourced from Beijing Vital River Laboratory Animal Technology Co, Ltd. The experiments were conducted in standard barrier unit facilities, where the animals were housed under a controlled environment with a 12-h light/dark cycle, consistent room temperature, and unrestricted access to food and water. All mice were randomly allocated into two groups (n = 8): (1) Control group; (2) HUA model group. Mice were induced to develop HUA through intraperitoneal potassium oxonate (PO) (Sigma, cat. # 156124) injections and ad libitum access to 10% fructose water. PO was dissolved in saline and administered to mice via intraperitoneal injection at a daily dosage of 350 mg/kg for 28 days. Mice in the control group were treated with saline (vehicle) as a control. On the 29th day, mice were sacrificed following a 12-h fast. The kidney tissues were immediately excised, weighed, and divided into two sections: one section was fixed in 4% paraformaldehyde, while the other section was stored at −80 °C.

Cell cultures and transfection

The HK2 (human renal cortex proximal tubule epithelial cells) were procured from Procell Life Science & Technology Co, Ltd and cultured in Dulbecco’s Modified Eagle Medium F-12 (Pricella, cat. # PM150312) supplemented with 10% fetal bovine serum (VivaCell, cat. # C04001) and 1% penicillin/streptomycin (Solarbio, cat. # P8420/S8290) within a damped atmosphere containing 5% CO2.

The SMARTPool siRNAs targeting WWC1 (si-WWC1) (NS-021962; Forward Primer: 5′- GGCUGAUCCUUAUCAACGA-3') and negative control siRNA (si-NC) were acquired from Tsingke Biotech. Donor sequences of the hWWC1 CDS flanked by homology arms were cloned into the pcDNA3.1-HA vector (Dahong, Guangdong, China, cat. # OENM_001161661). Donor sequences of the hYAP1/YAP5SA CDS flanked by homology arms were cloned into the pcDNA3.1-Flag vector (mailgene, cat. # MG240311002). Transient transfection was carried out using Lipofectamine 2000 (Thermo, cat. # 11668-019).

Serum biochemical analysis

Following the manufacturer's instructions, commercially available assay kits from Nanjing Jiancheng Bioengineering Institute were used to assess the concentrations of UA, creatinine, blood urea nitrogen, and XOD in rat and mouse serum. Additionally, assay kits were utilized to measure uric acid levels in both intracellular and extracellular compartments of HK2 cells.

Histopathological examination

Kidney tissues from rat and mouse were fixed in 4% paraformaldehyde for 24 h, followed by embedding in paraffin. Tissue sections (5 μm thick) were then stained with hematoxylin and eosin (HE) using standard procedures. Subsequently, kidney histological lesions were evaluated under a light microscope (OLYMPUS) at magnifications of 100 × and 200 × .

Real-time quantitative PCR (RT-qPCR)

According to the manufacturer's instructions, total RNA was extracted from rat and mouse kidney samples as well as HK2 cells using TRIzol (Accurate, Shanghai, China, catalog #AG21102). The RNA concentration was measured using a Quawell UV-Vis spectrophotometer Q5000 (Tripbiotech). RNA (0.5 μg) was used for cDNA synthesis with the TransScript First Strand cDNA Synthesis SuperMix Kit (Accurate, Shanghai, China, cat. # AG11706). The cDNA was then amplified using the ABScript II One Step SYBR Green RT-qPCR Kit (ABclonal, cat. # RK21204) and specific primers in the Gentier 96E/96R Real-Time PCR Detection system (Tianlong). The primer sequences are listed below:

hWWC1 forward, 5′-cgtggctgtccttccttgct-3′; hWWC1 reverse, 5′-cctgccctcctgcctcttct-3′; hβ-actin forward, 5′-CACCAGGGCGTGATGGT-3′; hβ-actin reverse, 5′-CTCAAACATGATCTGGGTCAT-3′; hABCG2 forward, 5′- CATCTTCTCCATTCATCAGCCTC-3′; hABCG2 reverse, 5′- ATCTTCTTCTTCTTCTCACCCCC-3′; hSLC22A8 forward, 5′- CCCCATCGGATCCAGACC-3′; hSLC22A8 reverse, 5′- CACCTCTCAGGCTTCCCATT-3′. RT-qPCR was carried out with the following conditions: 95 °C for 3 min, followed by 42 cycles of 95 °C for 15 s, 60 °C for 30 s, and 72 °C for 30 s. The relative mRNA expressions were calculated using the 2∧(−△△Ct) method, with β-actin as the housekeeping gene for normalization of the target genes.

Western blotting analysis

The kidney tissue of rat and mouse or HK2 cells were homogenized in radioimmunoprecipitation assay (RIPA) buffer containing 0.1% phenylmethanesulfonyl fluoride (PMSF) (Solarbio, Cat. No. #R0010), followed by centrifugation at 12,000 rpm for 20 min at 4 °C. The supernatant was collected, and protein concentrations were determined using a bicinchoninic acid (BCA) protein assay kit (Yamei, cat. # ZJ101). The protein samples were combined with 5 × loading buffer in a ratio of 4:1 and subsequently boiled for 10 min. Following this, the mixture was resolved using 8–10% sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) and subsequently transferred onto a polyvinylidene fluoride membrane (Merck, cat. # IPVH00010) for 1 to 2 h.

After the transfer, the membranes were incubated with a blocking solution containing 5% non-fat milk in triethanolamine-buffered saline with Tween (TBST) for 1 h at room temperature. Following that, the membranes were subjected to a 12 h incubation at 4 °C with antibodies targeting WWC1 (ABclonal, cat. # A17110), ABCG2 (CST, cat. # 42078), OAT3 (Proteintech, cat. # A3119), YAP1 (ABclonal, cat. # A1002), p-YAP1(S127) (Abcam, cat. # ab76252), LATS1 (CST, cat. # 3477), and p-LATS1 (Thr1079) (CST, cat. # 8654) at a dilution of 1:1000 each.

After three TBST washes, the membranes were subjected to a 12 h incubation at room temperature with secondary antibodies, AffiniPure Goat Anti-Mouse IgG (H + L) (Boster, cat. # BA1038) and AffiniPure Goat Anti-Rabbit IgG (H + L) (Bostercat. # BA1039) at a 1:10,000 dilution. Subsequently, membranes were imaged using an enhanced chemiluminescence kit (Advansta, cat. # K-12045-D50), and gray scanning analysis was conducted with Image J software (National Institutes of Health, Bethesda, USA).

Cellular cytoplasmic and nuclear proteins extraction

Total protein was extracted using a lysis buffer (10 mM HEPES-NaOH [pH9], 10 mM KCl, 1.5 mM MgCl2, 0.5 mM β-mercaptoethanol, proteinase inhibitor cocktail). The mixture was then vortexed with the addition of 10% NP40. After centrifugation at 16000g and 4 °C for 15 min, the resulting supernatant was collected as cytoplasmic protein. The sediment was suspended again in a nuclei lysis buffer (10 mM Tris-HCl, 420 mM NaCl, 0.5% NP40, 1 mM Dithiothreitol 1 mM phenylmethylsulfonyl fluoride, 2 mM MgCl2, proteinase inhibitor cocktail). After centrifugation at 16,000g and 4 °C for 15 min, the resulting supernatant was collected as the nuclear protein.

Immunofluorescence staining

After a 24 h treatment, specific groups of HK2 cells, cultured in 24-well plates, were fixed using 4% paraformaldehyde. Afterwards, the cells were incubated with anti-hWWC1 antibody at 4 °C overnight (1:100, ABclonal, cat. # A17110) following blocking with 3% FBS. The cells underwent incubation with the secondary antibody, ABflo 555-conjugated Goat Anti-Rabbit IgG (H + L) (1:400, ABclonal, cat. # AS057), at 37 °C for 1 h. Subsequently, the cell nuclei were stained with DAPI (Sigma, cat. # D5942) for 5 min in the dark. Immunofluorescence images were collected and processed by Zen Software.

Immunohistochemistry staining

The rat kidney samples embedded in paraffin were sectioned, followed by deparaffinization and dehydration using a series of xylene and ethanol gradient concentrations. Following this, the sections were subjected to a 10-min treatment with 3% hydrogen peroxide to suppress endogenous catalase activity. Afterward, the sections were pre-incubated with 5% goat serum for 1 h and then subjected to overnight incubation at 4 °C with the WWC1 antibody (1:200, ABclonal, cat. # A17110). The sections were subsequently incubated with a horseradish peroxidase-conjugated secondary antibody, AffiniPure Goat Anti-Rabbit IgG (H + L) (Bioss) at 37 °C for 1 h, followed by subsequent treatment with streptavidin HRP AffiniPure Goat Anti-Rabbit IgG (H + L) (Boster, cat. # BA1039). The WWC1 expression in the kidney was microscopically analyzed using a ZEISS microscope (Oberkochen, Germany) and assessed with Image J software.

Statistics and reproducibility

Group differences were assessed for statistical significance using Prism 8.3.0 (GraphPad Software Inc). We used two-tailed unpaired Student’s t-tests to compare between two groups. Statistical significance was defined as p-values < 0.05. Data are expressed as mean ± SD. Representative findings, unless otherwise specified, were derived from at least three independent experiments.

Data availability

The data used to support the findings of this study are available from the corresponding author upon request.

Supporting information

This article contains supporting information.

Conflict of interest

The authors declare that they have no conflicts of interest with the contents of this article.

Acknowledgments

Author contributions

Z. Z., H. C., W. H., J. H., Changshun Han, Chengyong He, and C. Z. conceptualization; T. P., Z. L., Changshun Han, and X. D. formal analysis; Changshun Han and Chengyong He writing–original draft; Changshun Han and Chengyong He writing–review & editing.

Funding and additional information

The work is funded by Natural Science Foundation of Fujian Province (2022J011356) and Fujian Clinical Research Center for Chronic Glomerular Disease (2021Y2018).

Reviewed by members of the JBC Editorial Board. Edited by Paul Shapiro

Contributor Information

Jiyi Huang, Email: hjy0602@163.com.

Weiping Hu, Email: hwp1227@126.com.

Supporting information

Supporting Information
mmc1.docx (1.5MB, docx)

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Associated Data

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

Supplementary Materials

Supporting Information
mmc1.docx (1.5MB, docx)

Data Availability Statement

The data used to support the findings of this study are available from the corresponding author upon request.

Articles from The Journal of Biological Chemistry are provided here courtesy of American Society for Biochemistry and Molecular Biology

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