
==== Front
J Biol Chem
J Biol Chem
The Journal of Biological Chemistry
0021-9258
1083-351X
American Society for Biochemistry and Molecular Biology

S0021-9258(24)02170-7
10.1016/j.jbc.2024.107669
107669
Research Article
Urolithin A attenuates hexavalent chromium-induced small intestinal injury by modulating PP2A/Hippo/YAP1 pathway
Guo Ping 12‡
Yang Rongfang 1‡
Zhong Shiyuan 1‡
Ding Yingying 1
Wu Jingnan 1
Wang Ziwei 13
Wang Huiqi 1
Zhang Jiaxin 1
Tu Nannan 1
Zhou Hongwei 4
Chen Shen 1
Wang Qing 1
Li Daochuan 1
Chen Wen 1
Chen Liping chliping@mail.sysu.edu.cn
1∗
1 Guangdong Provincial Key Laboratory of Food, Nutrition and Health, Department of Toxicology, School of Public Health, Sun Yat-sen University, Guangzhou, China
2 School of Public Health, Guangzhou Medical University, Guangzhou, China
3 Department of Pathology, Stony Brook Cancer Center, Stony Brook, New York, USA
4 Department of Toxicology, Guangzhou Center for Disease Control and Prevention, Guangzhou, China
∗ For correspondence: Liping Chen chliping@mail.sysu.edu.cn
‡ These authors contributed equally to this work

10 8 2024
9 2024
10 8 2024
300 9 1076697 3 2024
15 7 2024
© 2024 The Authors
2024
https://creativecommons.org/licenses/by-nc-nd/4.0/ This is an open access article under the CC BY-NC-ND license (http://creativecommons.org/licenses/by-nc-nd/4.0/).
Hexavalent chromium (Cr(VI)) exposure has been linked with gastrointestinal toxicity, whereas the molecular pathways and key targets remain elusive. Computational toxicology analysis predicted the correlation between protein phosphatase 2A (PP2A) and genes regarding Cr(VI)-induced intestinal injury. Here, we generated a mouse model with intestinal epithelium-specific knock out of Ppp2r1a (encoding PP2A Aα subunit) to investigate the mechanisms underlying Cr(VI)-induced small intestinal toxicity. Heterozygous (HE) mice and matched WT littermates were administrated with Cr(VI) at 0, 5, 20, and 80 mg/l for 28 successive days. Cr(VI) treatment led to crypt hyperplasia, epithelial cell apoptosis, and intestinal barrier dysfunction, accompanied by the decline of goblet cell counts and Occludin expression in WT mice. Notably, these effects were aggravated in HE mice, indicating that PP2A Aα deficiency conferred mice with susceptibility to Cr(VI)-induced intestinal injury. The combination of data analysis and biological experiments revealed Cr(VI) exposure could decrease YAP1 phosphorylation at Ser127 but increase protein expression and activity, together with elevated transcriptional coactivator with PDZ-binding motif protein driving epithelial crypt cells proliferation following damage, suggesting the involvement of Hippo/YAP1 signaling pathway in Cr(VI)-induced intestinal toxicity. Nevertheless, the enhanced phosphorylation of YAP1 in HE mice resulted in proliferation/repair defects in intestinal epithelium, thereby exacerbating Cr(VI)-induced gut barrier dysfunction. Notably, by molecular docking and further studies, we identified urolithin A, a microbial metabolite, attenuated Cr(VI)-induced disruption of intestinal barrier function, partly by modulating YAP1 expression and activity. Our findings reveal the novel molecular pathways participated in Cr(VI)-caused small intestinal injury and urolithin A could potentially protect against environmental hazards-induced intestinal diseases.

Keywords

hexavalent chromium
intestinal toxicity
protein phosphatase 2A
Hippo/YAP1 pathway
urolithin A
Abbreviations

ALP alkaline phosphatase

Cr(VI) hexavalent chromium

HE heterozygous

HO homozygote

IEC intestinal epithelial cell

IPA ingenuity pathway analysis

MOA mutagenic mode of action

PP2A protein phosphatase 2A

TAZ transcriptional coactivator with PDZ-binding motif

ZO1 zonula occludens-1

Reviewed by members of the JBC Editorial Board. Edited by Donita C. Brady
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pmcHexavalent chromium (Cr(VI)) compounds are proven toxin, widely used in steel manufacturing, painting, wood treatment, and refractory industries (1). It pollutes air, soil, and water, and eventually enters humans and animals by skin contact, oral ingestion, or inhalation. The Chinese Center for Disease Control and Prevention reported that the blood chromium in Chinese populations was 1.74 μg/L in 2009 to 2010 years (2). The epidemiological data revealed that Cr(VI) has been linked to adverse health effects, such as organ damage, asthma, rhinitis, and cancer (3, 4). The absorption of chromium in digestive system mainly occurs in small intestine, followed by large intestine (5). Accumulating evidence suggests that Cr(VI) exposure was closely correlated with gastrointestinal diseases, including irritation, diarrhea, acute hemorrhage, even cancer (6, 7). Animal studies also indicated that short-term oral exposure to high dose of Cr(VI) led to gastrointestinal irritation symptoms, ulcers, or anemia, while long-term low-dose exposure to Cr(VI) could increase the risk of gastrointestinal cancer (8). Previously, we also showed Cr(VI) exposure resulted in intestinal epithelial integrity damage via an in vitro gastrointestinal digestion model (9). The mechanisms for cytotoxicity and carcinogenicity of Cr(VI) involved increased levels of oxidative stress, cell apoptosis, DNA damage, and aberrant epigenetic pattern, etc (10, 11). Nevertheless, the key targets and molecular pathways regarding small intestinal injury induced by Cr(VI) exposure remain elusive.

In addition to nutrient absorption, the intestinal epithelium exerted a crucial barrier function to prevent the penetration of hazardous substances (12). In response to small intestinal injury, epithelial repair process will be initiated by crypt cell proliferation and differentiation to renew the damaged cells (13). Recent evidence demonstrated that impaired intestinal epithelial barrier was a common risk factor driving the occurrence of intestinal diseases such as colorectal cancer, inflammatory bowel disease, or Crohn’s disease (14, 15). Previous studies have reported crypt regenerative hyperplasia and genetic damage as key events in Cr(VI)-induced intestinal damage (16). Therefore, uncovering the perturbed pathways with respect to intestinal epithelium injury is critical for identifying the molecular events in Cr(VI)-induced intestinal injury.

The Cr(VI) toxicity in drinking water has been studied extensively, and the available datasets of in vivo and in vitro studies combined with advanced toxicological tools permits us to identify the signaling pathways and key targets in Cr(VI)-induced small intestine toxicity. Preliminary analysis predicted that most disturbed pathways underlying Cr(VI) toxicity was associated with protein phosphatase 2A (PP2A). PP2A accounts for the majority of phospho-serine/threonine phosphatase activity in eukaryote. Each holoenzymes contains a catalytic C subunit, a structural scaffolding A subunit, and a variable B regulatory subunit. PP2A scaffold subunit contains two isoforms, Aα and Aβ, which are 86% identical (17). Aα scaffold subunit is responsible for 90% of PP2A holoenzyme assemblies. PP2A dysregulation have been linked to several human disease, including cancer, cardiovascular disease, and Alzheimer’s disease (18, 19). The genetic variants in PP2A-Aα gene have been reported to increase the risk for a variety of human colorectal cancers (20), suggesting the critical role of PP2A in determining susceptibility to gut disease in populations. Our prior studies also showed that PP2A regulated chromium-induced cytotoxicity by modulating the induction of metallothionein or heat shock proteins (21, 22). Specially, we found the decreased PP2A Aα level in small intestine tissue upon Cr(VI) exposure (unpublished data), leading to the speculation that PP2A might be involved in the regulation of small intestinal toxicity of Cr(VI). Therefore, the understanding of signaling pathways governed by PP2A will help us to discover novel molecular events with regard to Cr(VI)-induced small intestinal injury.

Ingenuity Pathway Analysis (IPA) based on differentially expressed genes in published literatures underlying chromium toxicity predicted the association between PP2A and Yes1-associated transcriptional regulator (YAP1). YAP1, a major downstream effector of the Hippo signaling pathway, was indispensable for development, stem cell maintenance, and tissue homeostasis (23). Accumulating evidence has established the role of YAP1/Hippo signaling pathway in the development of a variety of human cancers (24). Particularly, YAP1 was implicated in the regulation of intestinal homeostasis (25, 26). Besides, we previously also showed the enhanced YAP1 phosphorylation in mouse model with PP2A Aα deficiency in hepatocytes (27). In this context, we suggested that PP2A might be implicated in regulating the integrity of intestinal epithelium by YAP1. However, whether PP2A/YAP1/Hippo signaling pathway participated in the small intestinal toxicity of Cr(VI) still remains undefined.

In this study, we established a mouse model with intestinal epithelium-specific deletion of Ppp2r1a (encoding PP2A Aα subunit) and characterized the regulatory role of PP2A in small intestinal toxicity of Cr(VI). Mechanistically, PP2A was involved in modulating the destruction or repair of gut barrier by YAP1/Hippo signaling pathway upon Cr(VI) exposure. Especially, urolithin A could alleviate Cr(VI)-induced disruption of intestinal barrier function by targeting YAP1. Our findings reveal the novel molecular pathways involved Cr(VI)-induced small intestinal injury and may open new avenues for the precaution and therapy of intestinal injury or intestine diseases.

Results

The establishment of mouse model with intestinal epithelium-specific deletion of Ppp2r1a gene

The mouse model with intestinal epithelium-specific deletion of Ppp2r1a gene were successfully generated by crossing Ppp2r1aloxp/+ with Vill-Cre mice. The offspring were genotyped to identify Ppp2r1a−/− homozygote (HO) and heterozygote (HE). The Vill−/− mice obtained from the same breeding were used as WT control (Fig. 1A). As expected, PP2A Aα mRNA expression in small intestinal tissues from HE and HO mice exhibited 54.43% and 87.9% reduction, respectively (Fig. 1B). In addition, PP2A Aα protein levels of HE and HO mice exhibited 45% and 98% reduction, respectively, in small intestinal tissues. In particular, Aβ expression was upregulated in HE and HO mouse intestinal epithelium (Fig. 1C), consistent with previous findings that PP2A Aα subunit could determine the protein expression and stability of Aβ subunit (28). Likewise, immunohistochemical staining revealed that Aα-positive staining in small intestine tissue was reduced by 58.5% in HE mice, and Aα only existed in lamina propria and muscularis of HO mice (Fig. 1, D and E). Consistently, tdTomato fluorescence analysis confirmed that Villin gene only expressed in intestinal epithelial cells (IECs), but not lamina propria and muscularis (Fig. S1A), indicating that Ppp2r1a was specifically deleted in intestinal epithelium. These data indicate that we successfully generated a mouse model with PP2A function deficiency in intestinal epithelium. HE mice displayed no obvious changes in terms of villi length, crypt depth, intestinal epithelium permeability, and organ function within 3 months after birth (Figs. 1, F–H and S2). Nevertheless, HO mice exhibited crypt hyperplasia, inflammatory cell infiltration, and absence of most villous epithelial structures (Fig. 1F). Moreover, we previously showed that Aα subunit downregulation led to a significant reduction in PP2A-attributable phosphatase activity (29). Therefore, to explore the role of PP2A in the regulation of Cr(VI)-induced small intestinal injury, we used HE mice in the following experiments.Figure 1 Generation of mouse model with intestinal epithelium-specific deletion of Ppp2r1a gene.A, DNA genotyping in WT (671 bp), HE (671 bp and 825 bp), and HO (825 bp) mice by PCR analysis. B, relative mRNA levels of PP2A Aα in small intestinal tissues from WT, HE, and HO mice (n = 5). C, protein levels of PP2A Aα and Aβ in small intestinal tissues from WT, HE, and HO mice. D, representative images shown for small intestinal sections stained with Aα antibody in WT, HE, and HO mice (200× magnification). E, quantification of PP2A Aα-stained area was expressed as percentage of small intestinal fields occupied compared with WT mice (n = 3). F, representative panoramic images of H&E-stained small intestinal sections from WT, HE, and HO mice (40× and 200× magnification). A total of 15 random fields were analyzed in each group, and the (G) length of intestinal villi and (H) depth of intestinal crypt were calculated. ∗p < 0.05, compared with WT mice. HE, heterozygous; HO, homozygote; PP2A, protein phosphatase 2A.

The effects of PP2A Aα deficiency on the Cr(VI)-induced small intestinal injury

To address the role of PP2A in the regulation of Cr(VI)-induced small intestinal injury, HE mice and matched WT mice were treated with Cr(VI) at dose of 0, 5, 20, and 80 mg/l for 28 days. The chromium concentration in blood was shown in Table S3. There was no significant difference in term of body weight and small intestinal length between WT and HE mice following 28 days Cr(VI) exposure (Table S4). H&E staining showed that the intestinal villi in WT mice upon Cr(VI) exposure exhibited atrophy and increasing blunting and thickening in a dose-dependent manner (Fig. 2A). In accordance, the intestinal villi length was decreased by 5.3%, 10.9%, and 19.6%, respectively, in WT mice compared with that in control mice (ptrend < 0.05). Since the shortening of villi represents damage to villous epithelium, these results suggest that Cr(VI) exposure disrupted the structure of small intestinal epithelium. Particularly, the decline of villus length in HE mice was more profound than that in WT mice (Fig. 2B), indicating that HE mice with PP2A Aα deficiency were more susceptible to Cr(VI) exposure.Figure 2 PP2A was involved in Cr(VI)-induced small intestinal injury. WT and HE mice were treated with 0, 5, 20, and 80 mg/l Cr(VI) in drinking water, respectively, for 28 days (n = 8∼11). A, representative panoramic images of small intestinal tissues sections stained with H&E in mice (40× and 200× magnification). B, the length of intestinal villi (n = 3), (C) depth of intestinal crypt (n = 3), and (D) ratio of intestinal villi length/crypt depth were quantified by ImageJ, a total of 45 randomized areas were analyzed in each group. Results were shown as mean ± SD. ∗p < 0.05, compared with WT control mice. #p < 0.05, compared with HE control mice. Cr(VI), hexavalent chromium; HE, heterozygous; PP2A, protein phosphatase 2A.

The intestinal epithelium repair depends on uniform proliferation and differentiation of crypt, and elongation of crypt is regarded as a signature of cell proliferation (30). As shown in Figure 2A, the elongation of crypt was found in a dose-dependent manner in WT mice, indicating that crypt proliferation might be an adaptive response to Cr(VI) exposure. In contrast, we found no obvious crypt elongation in HE mice, even shortened in 20 and 80 mg/l Cr(VI) group. Notably, HE mice at 20 and 80 mg/l Cr(VI) group displayed more apparent structural abnormality in small intestine, such as villus atrophy or crypt hyperplasia. In parallel, crypt depth in Cr(VI)-treated WT mice were increased in a dose-response manner (ptrend < 0.05), which maybe account for Cr(VI)-stimulated crypt hyperplasia to compensate for damaged epithelial cells. However, the enhancement in crypt depth was inhibited in HE mice upon Cr(VI) exposure (Fig. 2C), suggesting that dysregulation of PP2A might suppress the proliferation/repair of small intestinal epithelium. Furthermore, the intestinal villi length/crypt depth ratio, an indicator for the maturation degree of small intestine, were descended with the increasing dose in WT mice (ptrend < 0.05) (Fig. 2D), indicating the integrity and function of small intestine were affected by Cr(VI). In particular, there effects were more profound in HE mice, even in 5 mg/l group. These observations together suggest that PP2A Aα suppression in intestinal epithelium appeared to inhibit crypt cell proliferation, exacerbating Cr(VI)-induced small intestinal epithelial injury.

PP2A was involved in the regulation of intestinal epithelial function upon Cr(VI) exposure

As described above, Cr(VI) exposure damaged the structure of villi and crypt, which were responsible for the barrier function of intestinal epithelium. We next examined the intestinal barrier function following Cr(VI) exposure. As illustrated in Figure 3, A and B, the goblet cell counts were decreased by 21.3%, 34.30%, and 47.9%, respectively, in WT mice with 5, 20, or 80 mg/l Cr(VI) treatment compared to that in control mice (ptrend < 0.05). In HE mice, the goblet cell counts were declined by 19.7%, 52.1%, and 62.0%, respectively, indicating that the impaired intestinal barrier in HE mice was more remarkable than WT mice. In line with these alterations, the mRNA level of intestinal secretory protein Muc2 was decreased by 28.0% in WT mice, whereas showed a 38.8% decrease in HE mice at 80 mg/l group (Fig. 3C). Collectively, PP2A Aα deficiency seems to suppress Cr(VI)-induced mucus secretion, thereby aggravating Cr(VI)-induced intestinal barrier injury.Figure 3 The effects of PP2A Aα deficiency on the intestinal barrier function upon Cr(VI) exposure. WT and HE mice treated with 0, 5, 20, and 80 mg/l Cr(VI) in drinking water, respectively, for 28 days. A, representative images were shown in small intestinal tissues stained with AB-PAS (200× magnification). B, the number of goblet cells in indicated group (n = 3) was quantified by ImageJ. C, relative mRNA levels of Muc2 in WT and HE mice treated with Cr(VI) at indicated dose. D, representative images were shown in small intestine stained with an antibody against occludin (green). The nuclei (blue) were stained with 4,6-diamino-2-phenyl indole (200× magnification). E, occludin expression was quantified and expressed as relative fluorescence intensity normalized by control (n = 3). F, the intestinal permeability was determined by the concentration of FITC-dextran in serum (n = 3). ∗p < 0.05, compared with WT control mice. #p < 0.05, compared with HE control mice. AB-PAS, alcian blue/periodic acid-Schiff; Cr(VI), hexavalent chromium; HE, heterozygous; PP2A, protein phosphatase 2A.

Occludin and zonula occludens-1 (ZO1) are important components of tight junction function for intestinal barrier (31). In agreement, in presence of Cr(VI), the intercellular boundaries of intestinal epithelium became blurred (Fig. 3D). Meanwhile, occludin expression was decreased by 9.43%∼44.60% (p < 0.05) (Fig. 3, D and E). The similar results regarding the expression and distribution of ZO1 were also observed (Fig. S3). Importantly, HE mice exhibited a lower abundance of occludin and ZO1 expression in small intestine. Since tight junction reorganization allows for IEC renewal without loss of barrier function (32), we suggest that PP2A Aα deficiency inhibits the ability regarding IECs to migrate and renew, thereby impairing intestinal barrier. In agreement, Cr(VI) exposure resulted in 8.78% and 24.39% increase in intestinal permeability, respectively, in WT and HE mice (Fig. 3F), indicating that PP2A Aα defect enhanced Cr(VI)-induced intestinal barrier dysfunction. Taken together, these observations demonstrate the indispensable role of PP2A in maintaining the intestinal epithelial barrier function in response to Cr(VI) exposure.

PP2A regulated Cr(VI)-induced small intestinal toxicity by Hippo/YAP1 signaling pathway

To illuminate the molecular mechanism underlying Cr(VI)-induced small intestinal epithelial injury, we subjected 6022 genes regarding Cr(VI) exposure in vitro and 1511 DGEs in vivo to IPA analysis (List in Data S1). The functional annotations revealed that these genes were relevant to cell death/proliferation (Data S2). The upstream analysis identified 3962 regulatory molecules, of which 111 were associated with PP2A (Data S3). Notably, PP2A showed a strong association with YAP1 in the interaction network (Fig. S4). Consequently, we next explored whether Hippo/YAP1 signaling pathway was involved in Cr(VI)-induced intestinal toxicity. As shown in Figure 4, A and B, YAP1 was increased by 42.32% in WT mice upon Cr(VI) exposure, which were abolished in HE mice (Fig. 4, A and B). Meanwhile, Cr(VI) treatment led to 39.93% increase in transcriptional coactivator with PDZ-binding motif (TAZ) protein level. By contrast, there was no obvious change in HE mice (Fig. 4, C and E). In parallel, the downstream genes including Clu, Cry61, Ctgf, and Tnfrsf12a mRNA level were increased in WT mice upon Cr(VI) exposure, whereas the induction was abolished in HE mice (Fig. S5), suggesting the involvement of Hippo/YAP1 pathway in Cr(VI)-induced intestinal toxicity. Prior studies have reported the phosphorylation of YAP1 at Ser127 resulted in the cytoplasmic retention or proteasome degradation of YAP1, thereby suppressing YAP1 activity or expression (33, 34). As shown in Figure 4, D and F, the phosphorylation levels of YAP1 at Ser127 appeared to be downregulated in WT mice upon Cr(VI) exposure, whereas it was significantly elevated in HE mice. Besides, Cr(VI) exposure led to a significant enhancement of phosphorylated YAP1 at Ser109 in WT mice, whereas a slight decrease in HE mice (Fig. 4C). Together, these results suggest that PP2A participated in Cr(VI)-induced intestinal toxicity by affecting the phosphorylation of YAP1 at Ser127.Figure 4 The involvement of Hippo/YAP1 signaling pathway in the intestinal injury induced by Cr(VI).A, representative panoramic images of immunofluorescence staining with an antibody against YAP1 (green) in small intestinal sections from WT and HE mice treated with 20 mg/l Cr(VI) (40× and 200× magnification). The nuclei (blue) were stained with 4,6-diamino-2-phenyl indole (DAPI). B, YAP1 expression was quantified and expressed as relative fluorescence intensity normalized by to DAPI density. C, protein levels of indicated protein in small intestinal tissues were determined by immunoblotting analysis. D, representative images shown for small intestinal sections stained with p-YAP1(Ser127) antibody (200× magnification). E, quantification was shown for the fold change of indicated protein level relative to WT control mice. F, quantification of p-YAP1(Ser127)-stained area was expressed as percentage of small intestinal fields occupied (n = 3). ∗p < 0.05, compared with WT control mice. #p < 0.05, compared with HE control mice. Cr(VI), hexavalent chromium; HE, heterozygous; PP2A, protein phosphatase 2A.

YAP1 mainly regulates cell proliferation, cell survival, and regenerative repair (35). As expected, Ki-67–positive cells were increased by 20.56%, 55.08%, and 77.60%, respectively, in mice intestinal crypts following Cr(VI) exposure (ptrend < 0.05) (Fig. 5, A and B), suggesting that Cr(VI) exposure triggered the proliferation of crypt cells by upregulating YAP1 expression. However, apoptotic cells was also increased by 1.02-, 1.86-, and 2.72- fold in WT mice, and the positive cells were mainly located in epithelial layer of villi (Fig. 5, C and D). Importantly, YAP1 expression were positively correlated with the levels of Ki-67 (r = 0.59, p < 0.05) and number of apoptotic cells (r = 0.64, p < 0.05) (Fig. 5, E and F), suggesting that excessive YAP1 expression might not facilitate the intestinal epithelium repair. By contrast, the Ki-67–positive cells displayed no obvious changes in HE mice, but the apoptotic cells were enhanced by 1.93-, 2.58- and 2.82- fold, respectively, indicating that PP2A Aα deficiency suppressed crypt cell proliferation and exacerbated IEC apoptosis. Collectively, these results indicate that suppression of PP2A Aα disturbed the renewal and repair of intestinal epithelium via Hippo/YAP1 signaling pathway.Figure 5 Hippo/YAP1 signaling pathway mediated Cr(VI)-induced intestinal injury by regulating cell proliferation and apoptosis. Representative images of staining with (A) Ki-67 and (C) TUNEL in small intestinal sections WT and HE mice upon 20 mg/l Cr(VI) treatment (200× magnification). The nuclei (blue) were stained with 4,6-diamino-2-phenyl indole. The numbers of (B) Ki-67–positive cells and (D) apoptotic cells were quantified and expressed as the average number of positive cells every five visual filed per mouse. Spearman correlation analysis was performed to analyze the association between YAP1 expression and the number of (E) Ki-67–positive cells or (F) apoptotic cells, respectively. ∗p < 0.05, compared with WT control mice. #p < 0.05, compared with HE control mice. Cr(VI), hexavalent chromium; HE, heterozygous; TUNEL, terminal-deoxynucleotidyl transferase-mediated nick end labeling.

Urolithin A attenuates Cr(VI)-induced small intestinal toxicity by regulating Hippo/YAP1 signaling pathway

Recent studies have indicated that microbial metabolites could be as promising therapeutics against the loss of gut barrier integrity (36, 37). Then, we predicted the binding between 15 microbial metabolites and YAP1 by molecule docking (Table S5). Among of them, urolithin A have the lowest binding energy (−6.19 kcal/mol) with YAP1 (Fig. S6). To determine whether urolithin A has protective effects against Cr(VI)-induced gut barrier dysfunction, mice were treated with 20 mg/l Cr(VI) in presence or absence of urolithin A for 28 days. As shown in Fig. S7, urolithin A treatment led to 26.65% increase in YAP1 expression but 28.70% decline in phosphorylated YAP1 at Ser127, together with the enhanced transcription of YAP1 target genes, including Areg, Gprc5a, and IL1rn, suggesting that urolithin A could modulate YAP1 activity and expression. Notably, compared to that in Cr(VI) exposure group, cotreatment urolithin A (Cr(VI)-urolithin A) abolished Cr(VI)-triggered excessive induction of YAP1 as well as the downstream genes. Besides, cotreatment with urolithin A significantly decreased the villi shortening and crypt depth by 16.3% and 21.7%, respectively (Fig. 6, A–C), suggesting that urolithin A treatment relieved the Cr(VI)-induced intestinal epithelial damage. In particular, the reduction of goblet cells was restored in Cr(VI)-urolithin A group, and the brush border was relatively intact (Fig. 6, D and E). Correspondingly, cotreatment with urolithin A rebated the decline in occludin expression induced by Cr(VI) (Fig. 6, F and G). In accordance, urolithin A treatment reversed Cr(VI)-induced intestinal permeability increase by 44.7% (Fig. 6H). These results indicate that urolithin A could alleviate the impairment of intestinal epithelial integrity upon Cr(VI) exposure. In addition, we also measured the alkaline phosphatase (ALP) activity to assess the gut epithelial polarity, closely related to gut barrier function. As a result, Cr(VI) led to the decreased epithelial cell polarization by 45.02%, whereas urolithin A cotreatment notably abolished the Cr(VI)-induced ALP decrease (Fig. 6I). Taken together, urolithin A might exert a protective role against Cr(VI)-induced intestinal function dysregulation. Meanwhile, cotreatment with urolithin A reduced Ki-67–positive cells and apoptotic cells by 30.6% and 47.4%, respectively, compared to that in Cr(VI) exposure group (Fig. 7, A–D). These results further support the notion that urolithin A could attenuate Cr(VI)-induced hyperplasia and apoptosis by regulating YAP1 expression, leading to the maintenance of intestinal homeostasis upon Cr(VI) exposure.Figure 6 Urolithin A attenuated Cr(VI)-induced small intestinal injury by regulating Hippo/YAP1 signaling pathway. Mice were treated with 20 mg/l Cr(VI) in the presence or absence of urolithin A (20 mg/kg body weight) for 28 days (n = 10). A, representative panoramic images were shown for small intestinal sections stained with H&E (10× or 100× magnification). B, the length of intestinal villi (n = 3) and (C) depth of intestinal crypt (n = 3) were quantified by ImageJ. D, representative images were shown in small intestinal tissues stained with AB-PAS (200× magnification). E, AB-PAS staining was performed to count the number of goblet cells (n = 3). F, representative images of immunofluorescence staining with an antibody against occludin (green) in small intestinal tissues. The nuclei (blue) were stained with 4,6-diamino-2-phenyl indole. G, occludin expression was quantified and expressed as fluorescence intensity relative to control group (n = 3). H, the intestinal permeability was determined by the concentration of FITC-dextran in serum. I, the alkaline phosphatase (ALP) level was normalized to intestinal protein and expressed as unit/g protein. ∗p < 0.05, compared with control mice. #p < 0.05, compared with urolithin A mice. AB-PAS, alcian blue/periodic acid-Schiff; Cr(VI), hexavalent chromium.

Figure 7 Urolithin A treatment abolished the hyperplasia and apoptosis.A, representative images of immunohistochemistry staining with Ki-67 and (B) immunofluorescence staining with TUNEL in small intestinal sections (200× magnification) in each group (n = 3). The nuclei (blue) were stained with 4,6-diamino-2-phenyl indole. The numbers of (C) Ki-67–positive cells and (D) apoptotic cells were expressed as the average number of positive cells every five visual filed per mouse. ∗p < 0.05, compared with control mice. #p < 0.05, compared with urolithin A mice. TUNEL, terminal-deoxynucleotidyl transferase-mediated nick end labeling.

Discussion

The understanding of key targets and molecular events underlying Cr(VI)-induced small intestinal injury was critical for elucidating the mode of action for small intestinal toxicity of Cr(VI). In this study, we revealed that PP2A regulated proliferation, repair, and apoptosis of IECs by Hippo/YAP1 signaling pathway, participating in the Cr(VI)-induced small intestinal injury. Particularly, urolithin A could ameliorate Cr(VI)-induced disruption of intestinal barrier function and abnormal proliferation of crypt cells by modulate YAP1 activity or expression. These findings advance the understanding of mechanism regarding Cr(VI)-induced small intestinal toxicity and highlight a novel role of Hippo signaling pathway, and microbial metabolite urolithin A might potentially be used to protect against environmental hazards–caused intestinal impairments or intestine diseases.

The maintenance of apical-basal polarity of IECs depends on a dynamic balance among multiple cellular activities, including proliferation, differentiation, migration, and apoptosis, which are tightly regulated by distinct signaling networks (38). Disruption of intestinal mucosal homeostasis and epithelium integrity occurred commonly in various critical pathological conditions. It has therefore been argued that the tumors observed following chronic oral exposure to Cr(VI) arose by a mutagenic mode of action (MOA) involving DNA mutation as an early key event in the carcinogenic process (7). However, prior study showed that Cr(VI) administration caused villous cytotoxicity in mice, followed by crypt proliferation without DNA damage (39), and reported a MOA involving villous enterocyte cytotoxicity as an initial event in Cr(VI) toxicity (40). In agreement, we here also showed that Cr(VI) exposure damaged villous epithelium and stimulated crypt hyperplasia. We previously also indicated that epithelial integrity damage might be as a sensitive marker for the intestinal toxicity of oral exposure to Cr(VI) by a Caco-2/HT29-MTX coculture model (9), suggesting a possibility that destruction of intestinal epithelium led to a small amount of Cr-VI leakage, stimulating crypts proliferation. Together, our results support the MOA of Cr(VI)-induced intestinal injury regarding villous enterocyte cytotoxicity.

It has been demonstrated that aberrant activation of protein kinases was associated with Cr(VI) toxicity (41). As a serine/threonine phosphatase that antagonizes the effects of multiple kinases, PP2A was proved to be implicated in cellular stress response to environmental pollutants. For instance, PP2A deficiency enhanced carcinogenesis of Lgr5+ intestinal stem cells treated with carcinogens (42). We previously found that suppression of PP2A activity sensitized cells to Cr(VI)-induced cytotoxicity (22). Here, we also showed that PP2A Aα deficiency aggravated Cr(VI)-induced intestinal epithelial barrier dysfunction, suggesting that PP2A might be a modulator for intestine toxicity induced by oral contaminants. In particularly, in mice with ablated alpha4, a key regulator of PP2A, the small intestinal mucosa exhibited crypt hyperplasia, villus shrinkage, defective differentiation of Paneth cells, and reduced IECs migration along the crypt–villus axis (43). The epithelial barrier function is regulated by an apical junctional complex, and accumulating evidence has established the role of PP2A in the regulation of epithelial tight function (44), supporting the involvement of PP2A in maintaining intestinal epithelium integrity. Notably, we showed that HO mice (Ppp2r1aflox/flox; Vill+/−) exhibited crypt hyperplasia, inflammatory cell infiltration, and absence of most villous epithelial structures. Thus, these observations together indicate the pivotal role of PP2A in maintaining intestinal epithelium homeostasis and provide the novel sight into PP2A-mediated signaling pathways in the development of intestinal disease.

In this study, we revealed the involvement of YAP1/Hippo pathway in Cr(VI)-induced small intestinal injury. YAP1 was the main downstream effector proteins of Hippo signaling pathway. When Hippo pathway was in an off state, the activation of YAP1/TAZ mainly mediated cell proliferation and apoptosis, regulating cancer growth and organ size (45, 46). It is evident that YAP1 exerted a crucial role in epithelial renewal and remission of inflammatory bowel disease (47). For example, YAP1 deletion in macrophages could alleviate inflammatory bowel disease by regulating M1/M2 macrophage polarization (48). YAP1/YAP and EGL-44/TEAD were required for resistance to infections with pathogenic bacteria when intestinal barrier was disrupted in worms and mice (49). Interestingly, we showed that exposure to Cr(VI) triggered a number of adaptive responses such as induction of YAP1, triggering the proliferation of crypt cells to renew damaged IECs and differentiation into goblet cells, which conferred organism with resistance to Cr(VI)-induced enterotoxicity. However, YAP1 overexpression led to the dysplasia of crypt and enhanced cytotoxicity, suggesting a dual role of YAP1 in regulating cellular events, varied in different context. In agreement with our results, prior study also indicated that YAP1 could trigger Wnt/β-catenin signaling, and not only facilitated enterocyte self-renewal and crypt regeneration after colitis but also promoted colitis-associated colorectal cancer development via chronic inflammation and excessive regeneration (50). Together, these observations suggest that appropriate YAP1 level was crucial for the repair of damaged intestinal epithelium.

It has been reported that YAP1 regulation was tightly associated with posttranslational modifications including phosphorylation (51), such as phosphorylation of Ser127 resulted in 14-3-3 binding and cytoplasmic retention of YAP1 (52). Furthermore, NEK1 also phosphorylated YAP on Ser127 residue and promoted its proteasomal degradation (34). In contrast, the phosphorylation YAP1 Ser109 by LATS1/2 affected the transcriptional activity of YAP1 (53). In the current study, we showed that the dephosphorylation of YAP1 at Ser127 in response to Cr(VI) led to the upregulation of YAP1, sequentially triggered the abnormal crypt cell proliferation. In particular, loss of PP2A Aα subunit elevated the phosphorylation of YAP1 at Ser127 to disturb the Hippo/YAP1 pathway, thereby aggravating small intestine injury. Consequently, we conclude that YAP1 dephosphorylation at Ser127 is prerequisite for the Cr(VI) toxicity. In accordance, prior study has demonstrated that PP2A was a critical regulator of YAP1 activation during loss of epithelial barrier (49) and activated YAP1 sequentially triggered crypt cell proliferation to renew damaged IECs (28). These observations provides evidence that YAP1/Hippo signaling pathway mediated by PP2A contributed to Cr(VI)-induced intestinal toxicity. In addition, we also showed the enhanced phosphorylation of YAP1 at Ser109 following Cr(VI) exposure, whereas the biological function and role in Cr(VI) toxicity remains unclear and needs to be further characterized.

Urolithin A is a metabolite generated from ellagic acid and ellagitannins by gut microbiota. It possesses a variety of pharmacological effects, including antioxidant, anti-inflammation, and antiaging ability. In particular, urolithin A was found to restore colon damage induced by high-fat diet and regulate the gut microbiota (54). Urolithin A administration improved the intestine morphology architecture and significantly alleviated the disturbance of gut microbiota in mice upon ionizing radiation (55). Hence, these observations suggest the potential therapeutic applications of urolithin A in the treatment of inflammatory bowel disease. Mechanistically, microbial metabolites could enhance barrier function and significantly mitigate systemic inflammation and colitis via activating AhR- or Nrf2-dependent pathways to upregulate tight junction proteins (56, 57). Besides, urolithin A protected against iAs3+-induced disruption of tight junction proteins in IECs by the blockade of oxidative stress and inflammation (58). Together, these findings support a protective role of urolithin A for intestine against environmental hazards. In agreement, we also showed that urolithin A treatment could ameliorate gut barrier dysfunction upon Cr(VI) exposure. Subsequent experiments suggest that urolithin A could regulate the expression of YAP1, thereby maintaining the normal proliferation of crypt, contributing to alleviate the Cr(VI)-induced intestinal injury. In addition, the role regarding inducing autophagy of urolithin A was increasingly recognized, which appeared to become a therapeutic potential natural compound for aging and aging-related disease (59, 60). Moreover, Hippo signaling pathway has been extensively shown to regulate autophagy (61). In this regard, we suggest that urolithin A modulated Hippo/YAP1 activity via autophagy in intestinal epithelium cells upon exposure to Cr(VI). However, the deep mechanism need further study to elucidate.

In this study, we revealed the involvement of PP2A-mediated Hippo signaling pathway might exert a role in Cr(VI)-induced impairment of intestinal barrier function. These finding further supported the notion that barrier dysfunction may be an important molecular events in MOA for Cr(VI)-induced intestinal toxicity. Furthermore, the microbial metabolite urolithin A could attenuate Cr(VI) toxicity by modulating the YAP1 expression. Of note, given that disruption of human intestinal mucosal homeostasis and epithelial integrity has serious pathological consequences, urolithin A might be a promised therapeutic strategy for intestinal barrier dysfunction and intestines disease.

Experimental procedures

Generation of ppp2r1aflox/flox, Vill-Cre mice, and C57BL/6-Rosa26/tdTomato mice

Ppp2r1aloxp/+ mice, containing loxP sites flanking exons 5 to 6 of Ppp2r1a gene, were purchased from the Jackson Laboratory. Mice with intestinal epithelium-specific deletion of Ppp2r1a were generated by crossing Ppp2r1aloxp/+ mice with Vill-Cre mice. Ppp2r1aflox/+; Vill+/− mice were subsequently backcrossed with Ppp2r1aloxp/loxp mice to generate HO mice (Ppp2r1aflox/flox; Vill+/−) and HE mice (Ppp2r1aflox/+; Vill+/−). Ppp2r1aflox/flox; Vill−/− or Ppp2r1a+/+; Vill+/− mice obtained from the same breeding were used as controls and termed WT mice. The offspring were genotyped by PCR. The PCR specific primers are listed in Table S1. To determine the specificity of Villin expression, Rosa26-tdTomato mice (purchased from Jackson Laboratory) were crossed with Vill-Cre mice to generate Vill-tdTomato mice. All animal protocols were approved authorized by the Animal Care and Use Committee of the Animal Research Center of Sun Yat-sen University (No. 2020-073).

Animal treatments

Potassium chromate (K2CrO4, Aladdin) was dissolved in sterilized ultrapure water. WT and HE mice (8 weeks old) were randomly divided into four groups (n = 8∼11) and treated with 0 mg/l, 5 mg/l, 20 mg/l, and 80 mg/l Cr(VI) in drinking water, respectively. Of these, 5 mg/l was chosen according to the lowest-observed-adverse effect level based on animal experiments conducted by National Toxicology Program (62). Twenty milligrams per litre and 80 mg/l were determined by the Cr concentration of drinking water and groundwater in industrial chromium-contaminated area, respectively (63). As the daily intake of water is approximately 0.2 ml/g (body weight) of mice a day. The exposure dosages of mice consuming Cr(VI) containing drinking water can be estimated as 0.08 and 0.32 mg/kg/day in 20 mg/l and 80 mg/l water, respectively. For determining the effect of urolithin A, male C57BL/6 mice (n = 10) were administrated with 20 mg/l Cr(VI) in the presence or absence of urolithin A (20 mg/kg body weight) (Mackli) for four successive weeks.

Measurement of chromium content

Briefly, 100 μl whole blood was mixed with a solution comprised of 0.1% nitric acid to bring the total volume to 3 ml and rinsed with deionized water thoroughly. The supernatants were collected after centrifugation at 3000 rpm for 15 min at 4 °C and subjected to analysis by inductively coupled plasma-mass spectrometry (Agilent Technologies Co Ltd).

Histological analysis

The intestine was washed by ice-cold PBS, fixed in 4% paraformaldehyde, and prepared as Swiss rolls. The intestinal tissues (5 μm thick) were stained with H&E and examined under TissueFAXS Viewer system (Tissuegnostics). The length of villus and depth of crypts were quantified by software ImageJ (National Institute of Health, https://imagej.en.softonic.com). Alcian blue/periodic acid-Schiff staining was performed to examine the goblet cells according to manufacturer’s introductions. Fifteen random fields per section in mice (n = 3∼5) were counted.

Terminal-deoxynucleotidyl transferase-mediated nick end labeling staining

The apoptotic cells in intestinal tissues section (5 μm) from mice (n = 3∼5) were determined using a terminal-deoxynucleotidyl transferase-mediated nick end labeling kit (Beyotime Biotech). The slides were examined under TissueFAXS Viewer system and 15 random fields per slide were chosen to quantify apoptotic cells by ImageJ.

Assays of gut permeability

FITC-conjugated dextran (Sigma-Aldrich) dissolved in water was administered to mice with the dose of 60 mg/kg body weight by gavage as described previously (Woting and Blaut 2018). Four hours thereafter, blood was collected via puncture. The concentration of FITC-dextran in serum was determined by a plate reader with an excitation wavelength at 490 nm. The concentration of FITC-dextran in sera was determined by comparison to FITC-dextran standard curve.

ALP activity in intestinal tissue

The ALP activity assay in small intestinal tissue was conducted following manufacture’s protocol (Beyotime). Briefly, whole-cell lysates were prepared using ice-cold radio immunoprecipitation assay lysis buffer. After homogenization and centrifugation, the supernatant was incubated with p-nitrophenol for 10 min at 37 °C and then subjected to absorbance detection at 405 nm using a Microplate Spectrophotometer (BIOTEC). The ALP activity was expressed as unit per gram protein (U/g protein).

RNA extraction and quantitative real-time PCR

The relative mRNA expression of genes was determined as described previously (64). The primers for amplifying genes were provided in Table S2.

Immunoblotting analysis

Whole-cell lysates were prepared by using ice-cold radio immunoprecipitation assay lysis buffer (Beyotime Biotech) containing protease and phosphatase inhibitors (Roche). Proteins (40 μg) were subjected to 4 to 12% gradient acrylamide gel for SDS-PAGE before immunoblotting. The following antibodies were used: rabbit anti-PP2A Aα, anti-LATS1, anti-p-YAP1 (Ser109), p-YAP1 (Ser127) (Cell Signaling Technology), mouse anti-TAZ, rabbit anti-YAP1 (Proteintech group), and rabbit anti-PP2A Aβ (Bioworld).

Immunofluorescence and immunohistochemistry analysis

Immunohistochemistry staining was conducted on paraffin sections by incubation with the following primary antibodies: rabbit anti-PP2A Aα (1: 100, Cell Signaling Technology), p-YAP1 (Ser127) (Abcam), and rabbit anti-Ki-67 (1: 200, Servicebio). Immunofluorescence staining was performed on frozen slices by incubation with primary antibodies, including mouse anti-occludin (1: 100), rabbit anti-YAP1 (1: 100), and anti-ZO-1 (1: 100) (Cell Signaling Technology). The images were captured by TissueFAXS Viewer system. Quantification of positive regions was performed by ImageJ software and depicted as positive staining area or mean fluorescence intensity.

Bioinformatics analysis

The in vitro data underlying Cr(VI) toxicity was downloaded from Comparative Toxicogenomics Database (http://ctdbase.org/) by “Chemical-Gene/protein Interactions” function. In vivo data with Cr(VI) exposure was obtained from toxicogenomic analysis of mouse duodenal tissue treated with Cr(VI) in drinking water for 7 or 90 days (GSE87259). Differentially expressed genes were selected with |fold change|>2.0 and false discovery rate q < 0.05. The differentially expressed genes with dose-response manner were identified by BMDExpress v2.3 (65) and then subjected to IPA software (Qiagen, https://analysis.ingenuity.com/pa/installer/select), following the core analysis, including upstream regulators and interaction network. Gene Ontology enrichment analysis was performed in R version 4.2.3.

Molecular docking

The structure of candidate metabolites were downloaded from PubChem (https://pubchem.ncbi.nlm.nih.gov) and prepared by PyMol 2.3.2 software (https://pymol.org/2/). Protein crystal structure of YAP1 was attracted from Protein Data Bank (www.rcsb.org) (PDB ID: 6JK0). AutoDock Tool was applied to convert PDB file of YAP1 or candidate metabolites into AutoDock Ligand format (PDBQT) file. The target protein structure was pretreated by AutoTools, including removal of water molecules, addition with hydrogens and so on. Then, both ligands (candidate metabolites) and YAP1 were subjected to AutoDock 4.2.6 to predict the interaction. The Lamarckian algorithm was conducted with 50 runs. Molecular docking analysis was conducted by Lamarckian Generic Algorithm in Autodock Vina (Scripps Research Institute). Accordingly, the models with the lowest score and binding-energy value were selected and visualized for subsequent analysis.

Statistical analysis

Data are shown as the mean ± SD from three independent experiments. All statistical analysis was performed using SPSS 20.0 (IBM) and GraphPad Prism 7 (GraphPad, https://www.graphpad.com/support/prism-7-updates/). Statistical differences among groups were determined by either one-way ANOVA or two-way ANOVA analysis. Nonparametric spearman correlation analysis was performed to analyze the association between YAP1 expression and the number of apoptotic or Ki-67–positive cells. Differences were considered statistically significant when p < 0.05.

Data availability

Supplementary data to this article can be found online.

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.

Supporting information

Supporting Information

Supplemental Data S1–S3

Author contributions

P. G., R. Y., J. W., D. L., W. C., and L. C. conceptualization; P. G., R. Y., S. Z., Y. D., H. W., J. Z., and N. T. data curation; P. G. writing–original draft; P. G. and L. C. funding acquisition; R. Y., S. Z., Z. W., H. W., N. T., and H. Z. methodology; S. Z., Y. D., and J. Z. formal analysis; J. W. visualization; J. W. and Z. W. investigation; S. C., Q. W., D. L., and L. C. supervision; H. Z. and S. C. validation; Q. W., W. C., and L. C. writing–review and editing.

Funding and additional information

L. P. C. and P. G. were supported by 10.13039/100014717 National Natural Science Foundation of China 82073580 and 82304176 L. P. C. was supported by 10.13039/501100003453 Natural Science Foundation of Guangdong Province (China) 2021A1515010916 . P. G. and S. C. were supported by Open Project Program of Guangxi Key Laboratory of Environmental Exposomics and Entire Lifecycle Health, 10.13039/501100014969 Guilin Medical University (China) 2022-GKLEH-01 and 2022-GKLEH-03 .
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