TIGAR regulates intestinal mucus barrier integrity by inhibiting lactylation of G6PD/6PGD in ulcerative colitis

Domain: chemistry
Published: 2026-03-03
Editorial outcome: major_revision

Abstract
Oxidative stress and metabolic dysregulation in goblet cells are pivotal in ulcerative colitis (UC) pathogenesis. TIGAR promotes the synthesis of NADPH and contributes to mitigate oxidative stress, but how it regulates NADPH production and affects UC remains unclear. Here we demonstrate that TIGAR inhibits lactylation of the key NADPH-synthesizing enzymes G6PD (at K432) and 6PGD (at K38), thereby preserving their enzymatic activities by promoting G6PD homodimer formation and 6PGD binding to NADP+. In male UC mice, persistently low TIGAR expression elevates lactate levels, promoting the lactylation of G6PD and 6PGD and impairing their function. This process suppresses NADPH synthesis, exacerbating goblet cell oxidative stress. The resulting decline in Trx1 reductase activity induces S-nitrosylation of the mucin-processing enzyme AGR2, thereby inhibiting mature MUC2 production and compromising the intestinal mucus barrier. Our findings elucidate a mechanistic pathway through which TIGAR maintains cellular redox homeostasis, presenting it as a potential therapeutic target for UC.

Introduction
Ulcerative colitis (UC) is a chronic, nonspecific inflammatory bowel disease (IBD) of unknown etiology, primarily affecting the distal colonic mucosa and submucosa. Characterized by a prolonged and often recurrent course, UC significantly impacts patients' quality of life and can lead to colitis-associated cancer1, 2. UC is associated with various factors, including the intestinal environment, immune response, inflammation, genetics, diet, and gut microbiota, etc3, 4.

Recently, the relationship between the intestinal mucus barrier and UC has garnered increasing attention5, 6. Intestinal mucus serves as the first line of defense against the invasion of harmful factors and plays a crucial role in maintaining intestinal homeostasis7, 8. Current research primarily focuses on the effects of UC on intestinal mucus synthesis, mucus constituents, and the glycosylation modifications of mucins9, 10. However, whether UC affects the critical issue of converting mucin precursors to their mature forms remains unclear.

In the endoplasmic reticulum, a protein modification enzyme known as anterior gradient protein 2 (AGR2) plays a crucial role in the maturation of MUC211, 12. Both basic and clinical studies have confirmed that abnormal expression or dysfunction of AGR2 can impair mucins synthesis and modification, which are essential factors in the pathogenesis of UC11, 12. However, there are no reports on whether UC affects the enzymatic activity of AGR2. The activity of AGR2 depends on its redox balance, and oxidative stress can reduce AGR2’s activity, thereby impeding mucin synthesis and modification13. Oxidative stress in goblet cells is widely recognized as a significant contributor to mucin modification disorders and the onset of UC, making antioxidative strategies crucial for UC therapy14, 15. Additionally, abnormal cellular metabolism constitutes another key factor in UC. Studies have shown that intestinal epithelial cells in animal models and patients with UC exhibit a distinctive metabolic pattern characterized by reduced oxygen consumption, increased glucose utilization, and elevated lactate production16. This suggests that UC may induce metabolic reprogramming in intestinal epithelial cells, shifting glucose metabolism from oxidative phosphorylation to glycolysis17. It is now established that lactate can interfere with cellular metabolism and induce the lactylation of certain metabolic enzymes18. Recently, the lactylation of non-histone proteins has emerged as a focal point of research in this field19, 20. In this context, the TP53-induced glycolysis and apoptosis regulator (TIGAR) has gained attention for its regulatory effects on glucose metabolism and oxidative stress21, 22.

TIGAR exhibits phosphatase activity, catalyzing the dephosphorylation of fructose-2,6-bisphosphate to generate fructose-6-phosphate, which can be converted into glucose-6-phosphate. The latter then enters the pentose phosphate pathway (PPP) to synthesize reduced coenzyme II (NADPH)23. NADPH is one of the most crucial reducing agents in the body, providing hydrogen essential for maintaining the stability of various redox systems, including the glutathione and thioredoxin systems24. In living organisms, more than 60% of NADPH is synthesized via the PPP, where glucose-6-phosphate dehydrogenase (G6PD) and 6-phosphogluconate dehydrogenase (6PGD) serve as the rate-limiting enzymes regulating NADPH production25, 26. TIGAR's role in promoting glucose entry into the PPP and facilitating NADPH synthesis has been widely recognized. However, the relationship between TIGAR and these two rate-limiting enzymes remains underexplored. Only one study has demonstrated that TIGAR can promote the synthesis of G6PD but did not involve its enzymatic activity27. Furthermore, no investigation has been reported into whether TIGAR influences 6PGD activity. Given TIGAR's roles in inhibiting glycolysis and reducing lactate synthesis, we hypothesize that TIGAR may regulate the activities of these two enzymes through lactylation modification, thereby impacting NADPH synthesis.

Our findings revealed that TIGAR significantly inhibits the lactylation of both G6PD and 6PGD, and identified the lactylation sites at lysine residues 432 and 38 in their respective amino acid sequences. Low expression of TIGAR was found to be a key factor contributing to the enhanced lactylation of G6PD and 6PGD, resulting in decreased NADPH synthesis, induced S-nitrosylation of AGR2, impaired MUC2 maturation, and subsequent damage to the intestinal mucus barrier in male UC mice. This study establishes TIGAR as a critical molecule that influences MUC2 modification and mucus integrity by affecting cellular redox homeostasis and glucose metabolism. These findings provide experimental evidence supporting that TIGAR as a potential therapeutic target for UC.

Results
1. Intestinal TIGAR deletion aggravates colitis via oxidative stress in UC mice
The symptoms of DSS-induced colitis in mice closely resemble those of human ulcerative colitis28. Following 7 days of DSS induction, the HE staining results demonstrated that the crypt interstice was markedly widened (Supplementary Fig. 1a), and the crypt height was significantly reduced (Supplementary Fig. 1c). Correspondingly, the UC mice showed more severe disruption of the intestinal mucosal epithelium barrier accompanied by increased inflammatory cell infiltration (Supplementary Fig. 1b). AB-PAS staining indicated reduced neutral and acidic mucus synthesis (Supplementary Fig. 1a, d, g). We utilized Cy3-conjugated EUB338 probes, which specifically target bacterial DNA, and the MUC2-N antibody, which recognizes mature MUC2, to evaluate the damage to the mucus barrier in UC mice29. Both the proximal and distal colons of UC mice exhibited damage to the inner mucous layer (Supplementary Fig. 1e, f), thinning the outer mucous layer, and bacteria migrated to the inner mucous layer (Supplementary Fig. 1e, f). Additionally, we observed that the expression of TIGAR began to decrease from day 5 and showed a more pronounced reduction by day 7 (Supplementary Fig. 1h). Therefore, our results suggested that the reduced expression of TIGAR in colon tissue may play a significant role in the pathogenesis of ulcerative colitis.

To investigate the relationship between TIGAR and UC, we generated mice with conditional knockout of TIGAR in the intestinal tract (TIGARf/fVil1-Cre) by crossing T IGARf/f mice with Vil1-Cre mice, enabling conditional TIGAR knockout in the gut system (Fig. 1j), and verified by western blot (Fig. 1l). Following the same procedure of DSS administration, the TIGARf/fVil1- Cre mice exhibited symptoms including mucopurulent bloody stool, highly distorted crypts, increased inter cryptal spaces, and decreased intestinal mucus synthesis (Fig. 1a, b). These changes appeared two days earlier and were associated with more severe pathological damage in TIGARf/fVil1-Cre mice compared to TIGARf/f mice (Fig. 1a), suggesting that TIGAR deletion accelerated the onset and progression of UC.

Oxidative stress induced by DSS damages the mucus barrier30. TIGAR is an important glucose-regulating enzyme that promotes NADPH synthesis and inhibits oxidative stress31. To evaluate TIGAR's impact on the colonic mucus barrier, we measured oxidative stress markers (GSH, NADPH, NO) and activities of Trx1 and TrxR in colon tissues of TIGARf/fVil1-Cre mice treated with DSS. Compared to TIGARf/f mice, GSH content, GSH/GSSG ratio, NADPH content, NADPH/NADP+ ratio, and Trx1 reductase activity were significantly reduced (Fig. 1c-g), while NO levels were elevated (Fig. 1i). TrxR1 activity and protein levels of Trx1 and TrxR1 remained unchanged (Fig. 1h, k). Oxidative stress can induce the misfolding of ER proteins32, and GRP78 and CHOP levels were significantly elevated in colon tissues of TIGARf/fVil1-Cre mice (Fig. 1l). ER stress also caused MUC2 precursor misfolding, leading to their accumulation and reduced mature MUC2 synthesis (Fig. 1m). We established HT29 CL.16E cells with stable TIGAR knockout using CRISPR/Cas9 (Supplementary Fig. 2i). In vitro experiments mirrored the findings in TIGARf/fVil1-Cre mice (Supplementary Fig. 2a-i). TIGAR knockout in HT29 CL.16E cells increased ROS, NO, mitochondrial ROS, and superoxide levels (Fig. 1n-q). These results suggest that TIGAR deletion exacerbates intestinal oxidative stress and disrupts MUC2 maturation in UC mice.

2. TIGAR deficiency accumulate 6-phosphogluconate and lactate by reprograming the PPP and glycolysis
TIGAR has long been considered to primarily function as a fructose-2,6-bisphosphatase, which suppresses glycolysis and subsequently promotes the flux of the PPP to generate NADPH to reduce intracellular ROS21. However, there remains a lack of understanding regarding the specific mechanisms by which TIGAR regulates the distribution metabolic flux between glycolysis and the PPP. For this reason, we first conducted untargeted metabolomics to investigate the static changes of metabolites. We found a dramatic increase in 6-phosphogluconate (6-PG) levels in TIGAR-knockout HT29 CL.16E cells (Fig. 2a), which was identified by the mzVault library (Fig. S2b) and comparing the retention time of the standard. Then, we also observed a consistent increase in 6-PG in TIGAR-knockout cells (HT29 CL.16E and HIEC-6) and colon tissues from TIGARf/fVil1-Cre mice, with DSS induction further exacerbating the elevation of 6-PG (Fig. 2c). At the same time, our research revealed elevated lactate levels in both TIGAR-knockout cell lines and colon tissues from TIGARf/fVil1-Cre mice and DSS-induced TIGARf/fVil1 -Cre mice (Fig. 2d). The KEGG enrichment analysis of metabolite revealed that the PPP and glycolysis were both affected pathways following TIGAR-knockout in HT29 CL.16E cells (Supplementary Fig. 3a). These results suggest that 6-PG and lactate, key metabolites in the PPP and glycolysis, may have increased synthesis or suppressed catabolism without TIGAR. Furthermore, DSS induction appears to exacerbate this metabolic imbalance. To investigate the specific mechanisms underlying the reprogramming of glucose metabolism following TIAGR depletion, we need to perform isotope tracing analysis. First, we must ascertain whether the absence of TIGAR affects glucose uptake in cells or animals. Notably, our results indicated that the absence of TIGAR does not influence cellular glucose uptake (Supplementary Fig. 3b, c). The glucose oxidation rate appears to remain comparable between TIGARf/fVil1-Cre and TIGARf/f mice, as inferred from analyses of oxygen consumption and respiratory quotient (RQ) (Supplementary Fig. 3d-f)33, 34.

Then, we employed U-13C-glucose for stable isotope tracing to analyze metabolic flux changes within the glycolysis and PPP (Fig. 2e). Following incubation with U-13C-glucose 2h, we observed a reduced carbon flux through the PPP and increased carbon flux through glycolysis upon TIGAR-knockout in HT29 CL.16E cells, with the fractions of 13C-labeled glucose 6-phosphate (G6P), fructose-6-phosphate (F6P), 2-phosphoglycerate (2-PG), pyruvate (PYR), and lactate accumulated in TIGAR-deficient. In contrast, the fractions of 13C-labeled 6-PG and ribose 5-phosphate (R5P) in the PPP pathway were remarkably decreased (Fig. 2f). Next, we observed the temporal patterns of isotope incorporation into metabolites multiple times points in cells. The isotope exchange rate for 6-PG was lower in TIGAR-deficient cells compared to the control group. At the same time, the total 6-PG (labeled and unlabeled) levels remained consistently higher in TIGAR-deficient cells (Fig. 2g). This suggested that both the production and consumption of 6-PG were inhibited in TIGAR-deficient cells, with the inhibition of the consumption pathway being more pronounced. In contrast, lactate exhibited a different pattern. While the total lactate levels were higher in TIGAR-deficient cells compared to controls, the isotope exchange rate of lactate was also elevated, indicating that the production pathways of lactate are activated in TIGAR-deficient cells (Fig. 2h). To confirm the consistency of this metabolic reprogramming pattern caused by TIGAR deficiency in intestinal epithelial cells, we validated the findings in HIEC-6 cells and the colons of the TIGARf/fVil1-Cre mice. The results were consistent with those observed in HT-29 cells (Fig. 2j, Supplementary Fig. 3g-o).

To further investigate the underlying mechanism of this metabolic reprogramming induced by TIGAR deficiency, we performed RNA sequencing on the colons of the TIGARf/f and TIGARf/fVil1-Cre mice. RNA sequencing analysis demonstrated that there were 1261 differentially expressed genes (p-adjust<0.05, FC>1.2). KEGG pathway enrichment analysis indicated that these differentially expressed genes were mainly enriched in pathways metabolism, human diseases, cellular processes, etc (Supplementary Fig. 4a). Given that TIGAR is a gene involved in the regulation of cellular metabolism, we analyzed the differentially expressed genes within the metabolism module, which consisted of a total of 107 genes. The absence of TIGAR was found to upregulate the expression of multiple glycolysis-related genes, including Hk2, Pkm and LdhAa (Supplementary Fig. 4b). Gene set enrichment analysis (GSEA) revealed substantial alterations in glycolysis and gluconeogenesis pathways associated with TIGAR deficiency (Supplementary Fig. 4c, d). An increase in lactate flux can be attributed to the upregulation of metabolic enzymes involved in glycolysis at the RNA level (Supplementary Fig. 4b). However, the RNA levels of metabolic enzymes upstream and downstream of 6-PG in RNA-seq showed no significant changes with TIGAR deficiency, including G6PD and PGD (Supplementary Fig. 4b). Additionally, no significant differences were observed in the protein levels of G6PD and 6PGD (Supplementary Fig. 4e-g). Therefore, we speculate that the accumulation of 6-PG may be related to changes in G6PD and 6PGD enzymatic activity.

3. TIGAR deficiency exacerbates oxidative stress by promoting the lactylation of G6PD and 6PGD
We next investigated the activities of G6PD and 6PGD, and their enzymatic activities were significantly reduced in TIGARf/fVil1-Cre mice (Supplementary Fig. 5a, b). Specifically, the dimerization of G6PD was markedly decreased in TIGAR knockout cells and mice (Supplementary Fig. 5c, d), whereas the dimerization of 6PGD remained stable (Supplementary Fig. 5e). These findings suggested that TIGAR regulated the activity of these two enzymes, potentially through distinct mechanisms, thereby impacting NADPH synthesis. Our research revealed elevated lactate levels in both colon tissues from UC mice and TIGARf/fVil1-Cre mice and in TNFα-induced HT29 CL.16E and HIEC-6 cells (Fig. 2d). Given lactate's role as a substrate for protein lactylation35, we hypothesized that TIGAR knockout induces lactylation of G6PD and 6PGD. Pan-Kla levels were elevated in UC mice (Fig. 3a, b), TNFα-induced cells (Fig. 3c and Supplementary Fig. 5h-j), TIGARf/fVil1-Cre mice colon tissues (Fig. 3a, b), and TIGAR-knockout HT29 CL.16E cells (Fig. 3c and Supplementary Fig. 5j). Western blot analysis showed significantly elevated lactylation of G6PD and 6PGD in DSS-induced mouse colon tissues (Fig. 3d), with similar results in cell experiments (Fig. 3e).

To investigate the effect of lactylation on mucus synthesis in G6PD and 6PGD, we conducted proteomics analysis in HT-29 cells. LC-MS revealed lactylation sites at G6PD K432 and 6PGD K38 (Supplementary Fig. 5f, g). G6PD and 6PGD activity depends on dimerization and NADP+ binding. TIGAR-knockout reduced G6PD dimer formation in HT29 CL.16E cells (Fig. 3f). Molecular docking simulations predicted lactylation effects on their activity. G6PD K432 was near the dimer interface, forming a hydrogen bond with M411 of the opposing monomer, suggesting that lactylation may disrupt dimer formation (Fig. 3g). Previous studies reported that lactylation at K45 hinders G6PD dimer formation and possibly affects NADP+ binding36. However, our results showed that K45 was close to the NADP+ binding domain, likely affecting NADP+ binding rather than dimerization (Fig. 3h). 6PGD K38, located in the catalytic NADP+ binding domain, forms a hydrogen bond with NADP+, indicating that its lactylation may affect NADP+ binding (Fig. 3i). Thus, the lactylation of 6PGD K38 may affect the binding to NADP+. We explored the impact of lactylation on dimer formation and NADP+ binding by mutating specific sites. The WT and mutants of G6PD were expressed in G6PD knockout HT29 CL.16E cells by same sense mutation. Mutations at K45 and K432 inhibited G6PD lactylation (Fig. 3j), K45 primarily affected NADP+ binding (Fig. 3k). While K432R impacted dimer formation (Fig. 3l). This was due to their locations: K432 was in the dimerization domain (Fig. 3g), and K45 was in the NADP+-binding domain (Fig. 3h). Moreover, lactate was found to inhibit the activity of G6PD (Fig. 3m). However, when the lactylation site K432 was mutated, the enzymatic activity was restored (Fig. 3m). We utilized FX11 to inhibit LDH activity and observed that FX11 was capable of restoring G6PD activity (Fig. 3n). Mutating K38 of 6PGD also inhibited its lactylation (Fig. 3o) and hindered NADP+ binding without affecting dimer formation (Fig. 3o, p). Similarly, treatment with lactate inhibited the activity of 6PGD, however, when the lactylation site K38 was mutated, the enzymatic activity was restored (Fig. 3q). And following the inhibition of LDH, the activity of 6PGD was restored (Fig. 3r). Inhibiting lactylation of G6PD and 6PGD promotes MUC2 maturation (Fig. 3s, t). Subsequently, we generated G6PD or 6PGD-deficient HT29 CL.16E cells using the CRISPR-Cas9 method and found that G6PD or 6PGD-deficiency increased ROS, NO, and superoxide (Supplementary Fig. 5k-p). Re-expressing wild type or mutant G6PD/6PGD decreased these levels induced by TIGAR knockout (Supplementary Fig. 5q-s), with the mutants showing a more pronounced effect (Supplementary Fig. 5q-s).

4. TIGAR deletion leads to AGR2 S-nitrosylation and inhibits MUC2 maturation
Oxidative stress leads to a decrease in intestinal mucus synthesis, and AGR2 plays a crucial role in the synthesis and modification of MUC211. The synthesis of mature MUC2 was reduced in the colon tissue of UC mice (Supplementary Fig. 6a). We utilized the MUC2-VNTR antibody, which targets immature MUC229, to investigate its interaction and cellular co-localization with AGR2. The interaction between AGR2 and the MUC2 precursor has been validated through immunoprecipitation (Supplementary Fig. 6b, c), and their co-localization has been confirmed by immunofluorescence analysis of cell and colon tissue (Supplementary Fig. 6d-f). We then generated AGR2-knockout HT29 CL.16E cells and measured precursor MUC2 and mature MUC2 levels. The results demonstrated that AGR2 knockout led to an accumulation of precursor MUC2, whereas the level of mature MUC2 was reduced (Supplementary Fig. 6g). Our previous studies have confirmed that AGR2 S-glutathione modification influences mucus processing and synthesis.13 We measured total protein S-glutathione levels in the colon tissue of UC mice. Although total protein S-glutathione showed a slight increase, this change was not statistically significant (Fig. 4a). This indicates that other posttranslational modification modifications may affect the function of AGR2 in UC mice.

Previous studies have shown that changes in intracellular NO levels due to oxidative stress are linked to protein S-nitrosylation37. Given the significant increase in NO levels in UC mice (Fig. 4b), we hypothesized and confirmed elevated S-nitrosylation of proteins in the colon tissues of UC mice (Fig. 4c-e). Additionally, AGR2 sulfhydryl groups were significantly reduced (Fig. 4f, g), while AGR2 S-nitrosylation was increased (Fig. 4h). AGR2 facilitates MUC2 maturation by forming disulfide bonds with MUC2 precursors via its Cys81 residue11, 38. Our results show that increased AGR2 S-nitrosylation in UC mice reduce its binding affinity to MUC2, as confirmed by immunoprecipitation (Fig. 4i, j). Mutation at Cys81 led to the accumulation of MUC2 precursors and a decrease in mature mucin synthesis (Fig. 4k, l). We also observed increased S-nitrosylation of proteins and AGR2 in TIGARf/fVil1-Cre mice (Fig. 4m, n). These findings indicate that AGR2 Cys81 S-nitrosylation inhibits its interaction with MUC2 precursors, disrupting MUC2 maturation. TIGAR promotes MUC2 maturation by preventing AGR2 S-nitrosylation and maintaining the mucus barrier.

5. TIGAR deletion facilitates the Trx1-mediated S-nitrosylation of AGR2
Trx1 is a key intracellular oxidoreductase involved in denitrosylation and transnitrosylation, with its function regulated by the intracellular redox state39. In this study, we observed increased S-nitrosylation of Trx1 in the colon tissue of UC mice (Supplementary Fig. 7a), accompanied by reduced reductase activity and denitrosylation function (Supplementary Fig. 7b). Similar results were found in TNFα-induced HT29 CL.16E cells (Supplementary Fig. 7c, d). Overexpression of Trx1 increased SNO-protein levels, while knockdown decreased them (Supplementary Fig. 7e), indicating Trx1 primarily acts as an oxidase under inflammatory conditions. The redox status of Cys32 and Cys35 in Trx1 modulates its ability to S-nitrosylate or denitrosylate target proteins39, 40. TIGAR depletion increased Trx1 S-nitrosylation (Supplementary Fig. 7f-h), and redox electrophoresis showed disulfide bond formation in Trx1 Cys32-Cys35 and Cys62-Cys69 upon TNF-α treatment (Supplementary Fig. 7i), confirming loss of reductase activity. Trx1 was nitrosylated at Cys73, crucial for transnitrosylation41. Mass spectrometry confirmed S-nitrosylation at Cys73, a conserved site in mammals (Supplementary Fig. 7j, k). The NADPH regulates the activity of Trx1 oxidoreductase42. NADPH supplementation in TIGAR knockout cells reduced protein and Trx1 S-nitrosylation (Supplementary Fig. 7l, m), indicating TIGAR maintains Trx1 reductase activity by enhancing NADPH synthesis.

Trx1 transfers SNO to target proteins after S-nitrosylation at Cys7339. The primary sequence motif analysis on peptides containing identified S-nitrosylation sites revealed that proteins harboring CXXXXA, CXXI, and EC motifs were more susceptible to S-nitrosylation, with Cys81 of AGR2 located within these motifs (Supplementary Fig. 8a-c). Trx1-mediated transnitrosylation depends on target protein interaction39, 41. Immunoprecipitation and immunofluorescence confirmed Trx1-AGR2 interaction (Fig. 5a, b). Trx1 knockdown reduced AGR2 S-nitrosylation (Fig. 5c and Supplementary Fig. 8d), while overexpression increased it (Fig. 5d and Supplementary Fig. 8e), suggesting Trx1 transfers nitroso to AGAR2 in inflammation. Using plasmids with synonymous mutations, we first attenuated endogenous Trx1 in HT29 CL.16E cells using shRNA and subsequently expressed Flag-Trx1 (Supplementary Fig. 8f). Following Trx1 attenuation, both SNO-protein levels and AGR2 S-nitrosylation decreased (Supplementary Fig. 8g, h). In contrast, these S-nitrosylation levels increased upon Trx1 re-expression (Supplementary Fig. 8g, h). We found mutating Cys32 or Cys35 reduced SNO-protein (Fig. 5e, g), SNO-Trx1 (Fig. 5h), and SNO-AGR2 (Fig. 5f, i), while simultaneous mutation of Cys32 and Cys35 increased their S-nitrosylation (Fig. 5e-i), and reduced the synthesis of MUC2 (Fig. 5i). Mutating Cys73 significantly reduced their S-nitrosylation levels (Fig. 5g-i) and increased the synthesis of MUC2 (Fig. 5i), indicating Cys32 and Cys35 are involved in denitrosylation, while Cys73 is critical for transnitrosylation. Next, we found that NADPH addition after TIGAR deletion or Trx1 transnitrosylation inhibition with PX12 reduced AGR2 S-nitrosylation and increased the synthesis of MUC2 (Fig. 5j). Exogenous NADPH had no effect on AGR2 S-nitrosylation and the synthesis of MUC2 after Trx1 knockdown or simultaneous mutation of Cys32 and Cys35 (Fig. 5k, l), indicating NADPH modulates AGR2 S-nitrosylation by altering Trx1 reductase activity. In summary, TIGAR reduces AGR2 S-nitrosylation by promoting NADPH synthesis to maintain Trx1 reductase activity, ultimately enhancing mucus synthesis.

6. TIGAR maintains Trx1 and AGR2 activity through G6PD and 6PGD
Our results reveal an important role of G6PD and 6PGD in TIGAR-triggered oxidative stress, the NADPH/NADP+ ratio and GSH/GSSG ratio were declined by knockdown of G6PD and 6PGD (Supplementary Fig. 9a-d). We next investigated whether G6PD and 6PGD regulate the activity of Trx1 and AGR2. There was a substantial reduction in the reductase activity of Trx1 after G6PD and 6PGD knockout in HT29 CL.16E cells (Fig. 6a, b), accompanied by an over-oxidation (Fig. 6c). Furthermore, the S-nitrosylation of total protein, Trx1, and AGR2 was significantly elevated in G6PD or 6PGD deficient HT29 CL.16E cells (Fig. 6d-h and Supplementary Fig. 9e, f), but decreased after activating G6PD (Supplementary Fig. 9e, f). Moreover, the S-nitrosylation of Trx1 was reduced in G6PD or 6PGD knockout cells cultured in the presence of NADPH (Fig. 6i, j). These results confirmed that G6PD and 6PGD were crucial enzymes for promoting NADPH synthesis and maintaining the reductase activity of Trx1.

To further substantiate that TIGAR sustains Trx1 reductase activity through 6PGD and G6PD, we initially performed a knockout of endogenous TIGAR in HT29 CL.16E cells, followed by the re-expression of TIGAR, which resulted in a decline of Trx1 and AGR2 S-nitrosylation (Fig. 6k-n,). However, upon inhibiting G6PD or 6PGD, the S-nitrosylation levels of Trx1 and AGR2 were markedly elevated (Fig. 6k-n). We further verified the effect of G6PD and 6PGD lactylation on the S-nitrosylation of Trx1 and AGR2. Restriction of their lactylation can reduce the S-nitrosylation of Trx1 and AGR2 and restore Trx1 activity impaired by TIGAR knockout (Fig. 6o-r and Supplementary Fig. 9g, h), while also preventing MUC2 synthesis disorder (Supplementary Fig. 9i, j). These findings indicated that TIGAR preserves the activities of Trx1 and AGR2 by mitigating G6PD and 6PGD lactylation, thereby maintaining the synthesis of mature MUC2.

7. Mutant G6PD and 6PGD in TIGAR-KO mice alleviate DSS-induced colitis
To validate the lactylation sites of G6PD (K432) and 6PGD (K38) identified in this study and to assess the literature-reported G6PD (K45) site36, we generated K432R and K45R mutations in G6PD and K38R in 6PGD, introduced these mutants into TIGAR-knockout mice, and induced colitis with DSS four weeks later. The K432 and K38 mutations effectively prevented lactylation of G6PD and 6PGD, respectively (Fig. 7g, h), and significantly restored their enzymatic activities in TIGAR-knockout mice (Fig. 7e, f). In contrast, the K45 mutation had no significant effect on G6PD activity (Fig. 7e). Further investigation revealed that either the K432 or K38 mutations significantly reduced S-nitrosylation levels of Trx1 and AGR2, and markedly enhanced mature MUC2 synthesis (Fig. 7g, h). Morphological analysis showed that mutation of G6PD at K432 or 6PGD at K38 ameliorated colitis-associated intestinal damage, enhanced mucus production, preserved mucosal layer integrity (Fig. 7a-d), and markedly reduced bacterial load in the inner mucus layer (Fig. 7c, d).

Another finding of this study is that even unmutated wild-type G6PD or 6PGD exhibited some biological activity upon expression, but the magnitude of their effect was substantially lower than that of the K432R or K38R mutants, respectively. In contrast to the control (wild-type), the G6PD K45R mutant displayed no marked alterations in lactylation modification, enzymatic activity, and biological functions (Fig. 7a-h). Collectively, these findings indicate that although K45 is a site of G6PD lactylation, as supported by both published evidence36 and our cellular experiments, it has a comparatively modest impact on G6PD’s post-translational modification, activity, and antioxidant capacity relative to K432.

Methods
Cell lines and culture
The 293T cell line (ATCC, CRL-3216, fetal) was maintained in our laboratory and was cultured in DMEM medium (Gibco, Carlsbad, CA, USA) supplemented with 10% FBS (Gibco, Carlsbad, CA, USA) at 37℃ in a 5% CO2 incubator.
The HIEC-6 cell line (CRL-3266, fetal) was purchased from ATCC and cultured in DMEM medium (Gibco, Carlsbad, CA, USA) supplemented with 5% FBS (Gibco, Carlsbad, CA, USA), 10 mM HEPES, 5 mg/ml EGF, 100 U/ml penicillin/streptomycin, 4 mM Glutamax at 37℃ in a 5% CO2 incubator.
Sodium butyrate was utilized to differentiate HT29 cells into mature goblet cells, thereby obtaining mucus-secreting HT29 CL.16E cells50. Briefly, Human colon cancer cell line HT29 (ATCC, HTB-38, female) was cultured in a 5 mmol/L sodium butyrate solution for 9 days. Subsequently, the cells were passaged and maintained in a medium supplemented with sodium butyrate for 14 days to generate the HT-29 Cl.16E cell line. Over time, these cells differentiate into goblet cell phenotypes characterized by their ability to produce substantial amounts of mucus. The HT-29 Cl.16E cells were cultured at 37°C in a humidified atmosphere containing 5% CO2. The culture medium consisted of DMEM supplemented with 10% FBS, 100 U/mL penicillin/streptomycin, and 1 mM sodium pyruvate.

Animals and Experimental Models
Healthy male C57BL/6N mice (22-25 g, 8-10 weeks of age) were obtained from the Third Military Medical University Laboratory Animal Center. The animals were housed under SPF conditions in individually ventilated cages, with a 12-hour light/dark cycle and ad libitum access to food and water. Before the experiment, the mice were acclimatized to a standardized diet for one week. All procedures adhered to institutional guidelines and regulations and the experimental protocol was approved by the Experimental Animal Welfare and Ethics Committee of the Third Military Medical University (Approval number: AMUWEC20210636).
TIGARf/f mice have two lox sites on both sides of exon 2 in the TIGAR locus. Floxed mice were crossed with Vil1-Cre mice to obtain tissue-specific knockout of TIGAR. Mice were administered orally with 2.5% DSS dissolved in sterile distilled water for 7 days to establish the UC model. The mice were randomly assigned to the Cont, UC (DSS), TIGARf/f, DSS+TIGARf/f and DSS+TIGARf/fVil1-Cre groups. Following the experimental period, the mice were euthanized, and colon tissues were collected for subsequent analysis.
The adeno‐associated viruses (AAVs) used in the overexpression of G6PD (WT, K45R and K432R) and 6PGD (WT and K38R) were manufactured by Cyagen Biosciences Inc (Shanghai, China). 2 × 1011 genomes of the AAV9 vector for G6PD (WT, K45R and K432R) and 6PGD (WT and K38R) overexpression and control viruses were injected intraperitoneally into mice. Four weeks later, DSS was used to induce colitis for 5 days.
The animals used in this study were male. Unless otherwise specified, all references to animals in the text refer to males.

Stable knockout，knockdown and overexpressing cell lines
Stable knockout cell lines of TIGAR, Trx1, and G6PD were generated utilizing CRISPR/Cas9 technology. The double-stranded oligonucleotide complementary to the target sequence was cloned into the lentiCRISPRv2 vector and co-transfected with the packaging plasmid into HEK293 cells. Subsequently, a 48h viral supernatant was collected to infect cells with polybrene. The infected cells were treated with a specific concentration of puromycin for 7 days, and the particular gRNA sequence information was presented in Supplementary Table 1.
The shRNA sequence was cloned into the pLKO.1-Puro vector and co-transfected with packaging plasmids to construct stable knockdown cell lines. The viral supernatant was collected 48 hours post-transfection to infect cells, and infected cells were selected using puromycin for 7 days. Specific shRNA sequence information is provided in Supplementary Table 1.
Mammalian expression plasmids for Flag-, TIGAR, Trx1, AGR2, G6PD, 6PGD, and their mutants were constructed using standard molecular biology techniques. The sequences of WT or mutant were cloned into the pLV4ltr-Puro-CMV vector. The shRNA approach was used to construct stable overexpressed cell lines.

Mucin detection
HT29 Cl.16E cells or colonic tissues were lysed using a lysis buffer containing 1 mmol/L EDTA, 10 mmol/L Tris-HCl, 10% SDS, 5 mmol/L sodium chloride, and 20% Triton X-100, supplemented with 5 mmol/L phenylmethylsulfonyl fluoride (PMSF) and a protease inhibitor cocktail (Sigma-Aldrich). The lysates were then subjected to compound AgPAGE separation and transferred to a nitrocellulose membrane. The membrane was probed with two MUC2 precursor antibodies: MUC2-VNTR (Invitrogen, MA5-12345) and MUC2-VNTR (Novus, NBP2-25221), capable of detecting immature MUC2. The amount of mature MUC2 was detected using a MUC2-N antibody (Abcam, ab90007) in colonic tissue and HT29 CL.16E cells.

H&E staining, AB-PAS staining, and Histological scoring
Intestinal slices with a thickness of 5 μm were fixed overnight in 4% Carnoy's solution and then paraffin-embedded. H&E staining was performed by staining the sections with hematoxylin and dehydrating them in graded alcohols and xylene. For AB-PAS staining, dehydrated paraffin sections were stained with periodic acid, incubated with Schiff's reagent, and then counterstained with hematoxylin before dehydration. The histological score for colonic epithelial injury was determined as follows: crypt structure was scored from normal (0) to severe crypt distortion with complete crypt loss (3); the degree of inflammatory cell infiltration was scored from normal (0) to dense inflammatory infiltration (3); the score for muscle thickening ranged from normal (0) to evident apparent muscle thickening (3); depletion of goblet cells was scored from absent (0) to present (1); crypt abscesses were scored from absent (0) to show (1).
To highlight the images of the mucosal layer changes, we selected different mouse samples for staining and statistical analysis of crypt height changes. The statistics of crypt height were derived from five mice in each group. For each mouse, we measured the lengths of six consecutive crypts and calculated their average. Each point in the figure represents the average length of six consecutive crypts in one section of each mouse.

Determination of ROS, NO levels, and related redox couples
Cellular ROS levels were quantified using H2DCFDA (MCE, HY-D0940). Briefly, a 10 mM stock solution of H2DCFDA was prepared by dissolving it in DMSO and subsequently diluted for working concentrations. Adherent cells were incubated with PBS containing 5 μM H2DCFDA at 37°C for 30 minutes and harvested with 0.05% trypsin-EDTA. The cells were then resuspended in a fresh culture medium and immediately analyzed using NovoCyte flow cytometry (excited by a 488 nm laser) to measure fluorescence intensity. The MitoSox probe and DHE probe were used to detect mitochondrial ROS and superoxide, and the incubation method was consistent with that for ROS detection.
The concentration of NO was detected using a commercial kit (Beyotime, S0023). Briefly, cells and tissue samples were lysed with the NO detection-specific lysis buffer (Beyotime, S3090), and the reaction solution was prepared according to the instructions. After thorough mixing, the samples were incubated at room temperature for 10 minutes. Then, the OD value at 540 nm was measured, and the NO concentration was calculated based on the standard curve. The DAF-FM DA fluorescent probe (Beyotime, S0020S) was used to detect the intracellular NO level. Briefly, a staining working solution with a final concentration of 5 μM DAF-FM DA was prepared. The cell culture medium was removed and added the staining working solution. The cells were incubated in a 37 °C cell culture incubator for 20 minutes. The cells were washed three times with PBS to thoroughly remove the residuary DAF-FM DA. Fluorescence images were collected using a laser confocal microscope.
NADPH/NADP+ ratios in colon tissues and cells were measured using commercial kits (Beyotime, S0180S). The cell or tissue is first lysed using NADPH/NADP+ extractive buffers. The supernatant was then collected for analysis after centrifugation at 12000 × g and 4 ℃ for 5 min. Generate standard curves according to instructions. The concentration of NADPH in the sample was determined after the decomposition of NADP+ in a water bath at 60 ℃ for 30 min. The ratio of NADPH to NADP+ is calculated as follows: NADPH/NADP+ = (NSADPtotal- NADP+)/NADP+. The ratio of GSH/GSSG in colon tissues and cells was detected using commercial kits (Beyotime, S0053). Standard curves calculated the concentrations of GSH and GSSG.

Trx1 redox state monitoring and determination of Trx1, TrxR activity
To investigate the redox status of Trx1 in colon tissues and cultured cells, protein samples were prepared by precipitation with trichloroacetic acid (Macklin, T885181). The samples were labeled using the -SulfoBiotics Protein Redox State Monitoring Kit Plus (Dojindo, SB12). Cell and colon tissue extracts were subjected to SDS-PAGE and irradiated with ultraviolet light on a radiator for 10 minutes to eliminate protein shifters. Proteins were then electrophoretically transferred to polyvinylidene fluoride membranes. The membranes were blocked with 5% skim milk, incubated overnight with an anti-Trx1 antibody, and subsequently incubated with a secondary antibody. Signal strength was quantified using an ECL luminescence system. The activities of Trx1 and TrxR were quantified using the Thioredoxin Fluorometric Activity Assay Kit (Cayman, 500228) and Oxidized Thioredoxin Reductase(TrxR) Activity Assay Kit (Solarbio, BC1155), respectively, according to the manufacturer’s protocols.

Immunofluorescence
Cells were fixed with 4% paraformaldehyde for 15 minutes, permeabilized with 0.5% Triton X-100 for 15 minutes, and blocked with 5% normal serum for 1 hour at room temperature. Primary antibody was added and incubated overnight at 4 °C. After washing with TBST, fluorescent secondary antibody was applied and incubated at 37 °C for 1 hour. DAPI staining was performed in the dark for 10 minutes, and specimens were sealed with anti-fluorescence quencher solution before imaging under a fluorescence microscope.

Western blot
The cell or tissue protein was collected, and its concentration was determined using the BCA method. Subsequently, the protein was transferred to a PVDF membrane through SDS-PAGE electrophoresis. The membrane was then blocked with 5% skim milk powder and incubated overnight at 4℃ with the appropriate dilution of primary antibody working solution. Following this, the membrane was washed for 10 minutes each with TBST buffer, then incubated at room temperature for 90 minutes with the secondary antibody working solution. Subsequently, the film should be washed three times with TBST buffer and observed using an ECL luminescent solution. Antibodies: anti-TIGAR (Santa Cruz, sc-166290), anti-Trx1 (Abcam, ab273877), anti-Trx1 (ABclonal, A4024), anti-TrxR (Abcam, ab124954), anti-AGR2 (Abcam, ab76473), anti-G6PD (Abcam, ab993), anti-6PGD (Invitrogen, PA5-80894), Anti-L-Lactyl Lysine (PTM Bio, PTM-1401RM), anti-MUC2 (Invitrogen, MA5-12345), anti-MUC2 (Novus, NBP2-25221), anti-MUC2 (Abcam, ab90007), anti-Flag (protein tech, 20543-1-AP), anti-Flag (protein tech, 66008-4-Ig), anti-CHOP (Abcam, ab11419), anti-GRP78 (Abcam, ab108615), anti-GAPDH (protein tech, 60004-1-Ig), anti-Tubulin (protein tech, 66031-1-Ig).

Co‑immunoprecipitation (Co‑IP)
The co-immunoprecipitation assay used corresponding antibodies with the Pierce™ Classic Magnetic IP/Co-IP Kit (Thermo Fisher Scientific, 88804). In Brief, the cells were lysed on ice using an IP buffer containing a protease inhibitor, followed by centrifugation at 14000 x g for 20 min at 4 ℃. The resulting supernatant was collected. Incubation with 5 μg of antibody was carried out overnight in a shaker at 4 ℃, followed by the addition of 25 μl of clean Protein A/G magnetic beads. The mixture was then shaken for 4 h at 4 ℃, and the magnetic beads were collected. Then, he collected magnetic beads, which were washed with IP buffer and wash solution buffer and finally eluted with 100 μl of eluent to obtain proteins for western blot analysis.

Biotin switch assay for detection of S-nitrosylation
The S-nitrosylation modification of proteins was detected with minor improvements51. Cells were lysed using HENS buffer (250 mM HEPES, 4 mM EDTA, 0.1 mM neocuproine, 1% SDS), and equal protein amounts were incubated with MMTS at 50 °C for 20 minutes to block free cysteine thiols. After acetone precipitation and centrifugation, excess MMTS was removed using 70% acetone. The precipitate was resuspended in HENS buffer, mixed with biotin-HPDP and sodium ascorbate, and incubated for 1 hour in the dark. Following acetone precipitation and washing, the sample was neutralized and incubated with streptomycin at 4°C for 12-18 hours. After low-speed centrifugation, the sample was washed, eluted, and analyzed by SDS-PAGE.

Untargeted Metabolomics Analysis
For cellular samples, 1 mL of pre-chilled (at -80°C) 80% (v/v) methanol of MS grade was added to the culture dish, and cell debris was scraped off using a spatula. The cells were disrupted using a low-temperature ultrasonic disrupter for 60 seconds at -20°C, followed by a pause of 30 seconds, with this cycle repeated three times. 25 mg of colon tissue was weighed and mixed with 750 μL of pre-chilled 80% methanol for mouse colon samples. A low-temperature tissue grinder was utilized to homogenize the tissue for 60 seconds at -20°C, with a 30-second pause, repeated three times. The resulting homogenates from cells and colons were transferred and incubated at -20°C for 60 minutes. Subsequently, samples were centrifuged at 18,410 x g for 10 minutes at 4°C. The supernatant was collected after centrifugation, dried under a nitrogen gas, reconstituted, and centrifuged again to obtain the supernatant for subsequent analysis. A 20 μL aliquot from each sample was combined to prepare a quality control (QC) mixture for mass spectrometry analyses.
An Orbitrap Exploris 120 Mass Spectrometer (Thermo Fisher Scientific) was used to analyze the hydrophilic metabolites extracted from cells and colon tissues. The column temperature was 45 °C. In negative ion mode, mobile phase A (5% ACN, 10 mM ammonium bicarbonate, pH 9) and mobile phase B (95% ACN, 10 mM ammonium bicarbonate, pH 9) were eluted by a gradient according to proportional mixing on a BEH Amide HILIC column (2.1 × 150 mm, 1.7 μm, Waters). The linear gradient was as follows: 0 min, 99% B; 0.1 min, 99% B; 6 min, 30% B; 6.5 min, 99% B; 10 min, 99% B. The spray voltage was 2.5 kV for negative ion mode. The capillary temperature was 350 °C for negative ion mode. The mass range (m/z) was 70–1,050 for negative ion mode.
The raw files of untargeted metabolomics were searched using Compound Discoverer 3.3 (Thermo Fisher Scientific), which is based on the mzCloud and mzVault databases. The mass tolerance for precursor ions was set at 10 ppm, while the mass tolerance for fragment ions was defined as 15 ppm. For quantification purposes, a retention time shift of up to 0.25 minutes was permitted. For detailed methods, please refer to the Supplymentary information.

Isotope Tracing Analysis
Cells were cultured in a glucose-free medium supplemented with 10 mM D-glucose (MCE, HY-B0389) and U-13C-labeled glucose (MCE, HY-B0389A). Samples were collected at 1, 2, 4, 8, and 24 hours post-treatment. For metabolite extraction, 1 mL of pre-chilled 80% (v/v) methanol was added to each culture dish, and subsequent sample processing was performed according to established protocols for untargeted metabolomics.
In the in vivo experiment, mice were subjected to a 12-hour fasting period before treatment. 100 μL 1 M Unlabeled glucose and U-13C-labeled glucose were administered intravenously via the tail vein at 15-minute intervals. After the final injection, the mice were euthanized, and colon tissue samples were harvested for further analysis.
Isotope tracing analysis was performed using an Orbitrap Exploris 120 Mass Spectrometer (Thermo Fisher Scientific) with a column temperature maintained at 30 °C. In negative ion mode, gradient elution was conducted at a flow rate of 0.5 mL/min on a Premier ZHILIC column (2.1 × 100 mm, 1.7 μm, Waters) using mobile phase A (5% ACN, 15 mM ammonium acetate, pH 9) and mobile phase B (95% ACN, 15 mM ammonium acetate, pH 9). The linear gradient was as follows: 0 min, 90% B; 5 min, 65% B; 6 min, 65% B; 8 min, 90% B; 10 min, 90% B. The ion source used was H-ESI, operating in negative ion mode at 3 kV, with a sheath gas of 60 Arb, an auxiliary gas of 20 Arb, and a sweep gas of 2 Arb. The ion transfer tube temperature was maintained at 380°C, while the ion source evaporation temperature was set at 350°C. The collected data were analyzed using Compound Discoverer 3.3 software following the isotope-tracing analysis workflow. For detailed methods, please refer to the Supplementary information.

Pan antibody-based PTM enrichment and LC-MS/MS analysis
The HT29 CL.16E cells were exposed to lactic acid for 24 hours to enhance the Kla-modified peptides. The TNFα treated the HT29 CL.16E cells with Trx1 overexpression for 24 h to improve the S-nitrosylated peptides. The tryptic digest was resuspended in IP buffer (1 mM EDTA, 50 mM Tris-HCl, 100 mM NaCl, 0.5% NP-40, pH 8.0) and subsequently incubated with pre-washed Anti-L-Lactyl Lysine Antibody Conjugated Agarose Beads (PTM Bio, PTM-1404) or Anti-TMT Antibody beads (Thermo Scientific, 90076) at 4°C for 12 hours. The bound peptides were eluted using 0.1% trifluoroacetic acid (TFA). Following vacuum-drying, the eluted peptides were desalted using Millipore C18 ZipTips before LC-MS/MS analysis. LC-MS/MS analysis was supported by Jingjie PTM BioLabs. For detailed methods, please refer to the Supplementary information.

RNA-Seq
Total RNA was extracted from colon tissues of three wild-type (WT) mice and three TIGAR knockout (KO) mice using TRIzolTM reagent (Invitrogen). The purity and concentration of RNA were assessed using the Agilent 2100 Bioanalyzer system (Agilent Technologies). The mRNA was isolated from total RNA using the Dynabeads® mRNA Purification Kit (Invitrogen). The mRNA was fragmented using a fragmentation buffer, and a cDNA library was constructed using the fragmented mRNA as a template. Sequencing was performed on the Illumina NovaSeq6000 platform at the Beijing Genomics Institute, and the data were aligned to the UCSC mm10 reference genome using TopHat.

Determination of G6PD/6PGD activity and the G6PD/6PGD dimer
The activity of G6PD was measured using a commercial Kit (Beyotime, S0189). In brief, 1 × 106 cells or 20 mg of tissue samples were collected and lysed with G6PD extraction solution at 4 °C. The protein concentration was normalized and quantified using a BCA assay kit (Beyotime, P0010). 50 μL of the protein sample and 50 μL of the G6PD test solution were added to each well of a 96-well plate. Following a 10-minute incubation at 37 °C in the dark, the OD value was measured at 450 nm using a microplate reader. The 6PGD activity was measured using the 6-Phosphogluconate dehydrogenase Activity Assay Kit according to the manufacturer’s protocols (Solarbio, BC2105).
G6PD or 6PGD dimer was detected using the DSS crosslinking method. Briefly, the culture medium was aspirated, and cells were washed twice with PBS. Cells were subsequently scraped in PBS containing an EDTA-free protease inhibitor cocktail. DSS was added to the samples, followed by cross-linking through rotation at 37 °C for 30 minutes. An equal volume of 1M Tris (pH 7.5) was then added to achieve a final concentration of 20 mM, and the mixture was incubated at room temperature for 15 minutes. Cell lysis was performed using M-PER buffer and centrifugation at 16,000 × g for 10 minutes at 4°C. The supernatant was collected for subsequent western blot analysis.

Statistical analysis
All statistical analyses were conducted using GraphPad Prism (v 8.0.1, GraphPad Inc.) or R v.4.1.1. Unless otherwise stated, results are reported as mean ± standard deviation (SD). Between-group comparisons were conducted using the unpaired Student’s t-test, unpaired t-test with Welch's correction, or the Mann–Whitney U test, depending on the data's normality and the equality of variances across groups. Multigroup comparisons were analyzed through one-way analysis of variance (ANOVA) with Tukey’s honestly significant difference (HSD) post hoc test or two-way repeated-measures ANOVA. All hypothesis tests were two-tailed, and a predetermined significance level of α = 0.05 was used as the threshold for statistical significance across all analyses.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "This study revealed that TIGAR reduces 6-phosphogluconate and lactate levels by reprogramming the pentose phosphate\npathway (PPP) and glycolysis. Its deficiency exacerbates oxidative stress by promoting the lactylation of G6PD and 6PGD.\nSpecifically, lactylation at lysine residues K435 in G6PD and K38 in 6PGD impairs NADPH synthesis. This impairment\ninduces S-nitrosylation of AGR2, disrupts MUC2 maturation, and subsequently damages the intestinal mucus barrier in\nulcerative colitis (UC) mouse models. Overall, this work elucidates the role of TIGAR in UC pathogenesis and establishes a\ncompelling connection between TIGAR and G6PD/6PGD through high-quality data and well-designed experiments.\nHowever, several key concerns should be addressed prior to publication:\n1. Human Tissue Validation:It would be valuable to include analysis of human UC biopsy samples to validate whether\nTIGAR expression correlates with disease severity and whether G6PD/6PGD lactylation is elevated in UC patients. This\nwould significantly strengthen the translational relevance of the findings.\n2. Functional Validation of Lactylation Sites:While the study identifies K435 (G6PD) and K38 (6PGD) as critical lactylation\nsites, direct functional evidence is required to confirm that lactylation at these specific sites directly impairs enzyme function.\nEnzymatic activity assays comparing wild-type (WT) proteins to non-lactylatable mutants (e.g., lysine-to-arginine mutants)\nwould provide this critical evidence.\n3. Mechanism of Lactylation Induction:The hypothesis that lactate accumulation drives G6PD/6PGD lactylation requires\nexperimental validation. Pharmacological inhibition of lactate dehydrogenase (LDH) could test whether reducing lactate\nlevels rescues enzyme activity. Furthermore, investigating the potential involvement of the glyoxalase pathway and enzymes\nlike GLO2/S-lactoylglutathione (SLG) hydrolase in this lactylation is warranted, given recent findings that SLG can induce D-\nlactylation in immune cells.\n4. Figure Labeling Consistency:There are inconsistencies in figure labeling:\nIn Figure 2, the letter labels for subfigures show variations in capitalization (e.g., \"A\" vs. \"a\").\nIn Extended Data Figure 8, the subfigure labels (e.g., a, b, c) display variations in font size.", "reviewer_2": "In this study, the authors aimed to characterize how TIGAR modulates glucose metabolism in the context of ulcerative colitis.\nThey combined a variety of approaches including classical biochemistry, mass spectrometry-based metabolomics and\nproteomics, stable isotope tracing, RNA sequencing, and mouse physiology, and uncovered a signaling axis whereby\nTIGAR deletion elevated lactylation of key pentose phosphate pathway (PPP) enzymes, leading to reduced Trx1 activity,\ndecreased S-nitrosylation of AGR2, and impaired mucin maturation. The study is very interesting, and the data are generally\nof reasonable quality.\nA major concern with the current version of the manuscript is the lack of direct evidence demonstrating that AGR2 and Trx1\nare S-nitrosylated, and that 6PGD and G6PD are lactylated in vivo, particularly in disease-relevant mouse tissues. While the\ncell-based data are convincing, validating these post-translational modifications in primary tissues would significantly\nstrengthen the manuscript and enhance its suitability for publication in Nature Communications. See detailed comments\nbelow:\n1. In Extended Data Fig. 1a, c, the H&E staining does not seem to show a reduction of crypt height by half, as stated. Also, it\nis unclear whether the data points in Extended Data Fig. 1c represent measurements from different mice or multiple sections\nfrom the same mouse. Please clarify this in the figure caption.\n2. Lines 137–139: “These changes appeared two days earlier and more serious pathological damage in TIGARf/fVil1-Cre\nmice compared to TIGARf/f mice” — If the authors wish to argue that TIGAR deletion accelerates the onset and progression\nof colitis, they should provide comparative data at earlier time points (e.g., day 5 vs. day 7 of DSS treatment) between\nTIGARf/fVil1-Cre and TIGARf/f mice.\n3. In Fig. 2a, the authors should label all metabolites related to the PPP in the volcano plot. While the KEGG pathway\nanalysis highlights PPP as the most significantly regulated pathway, visualizing the changes in individual PPP metabolites\nwould enhance the interpretability of the data. In Extended Data Fig. 3a, the main text refers this data as being from HT29\nCL.16E cells, but the legend says it’s from TIGARf/fVil1-Cre mice. Please clarify.\n4. In Fig. 2h, the data clearly show increased lactate production in TIGAR-deficient cells. However, based on the current\ndata, the authors cannot conclude that lactate consumption is also increased.\n5. In Extended Data Fig. 4, please describe how many genes were changed.\n6. In Fig. 3, it is critical to demonstrate that 6PGD K38 and G6PD K432 are indeed lactylated in vivo. The authors should\nperform pull-down of these two enzymes from mouse tissue followed by mass spectrometry–based proteomic analysis to\nprovide direct evidence of site-specific lactylation in vivo. Also, the authors should provide proteomic evidence of S-\nnitrosylation of AGR2 in mouse tissues in Fig. 4 and also Trx1 if possible.\n7. In Fig. 3p, it’s not convincing that 6PGD K38R increases the MUC2 band from the data presented.\n8. Extended Data Fig. 4f and g are missing.\n9. In Extended Data Fig. 6d, it’s very hard to tell if MUC2 and ARG2 colocalize from the staining. The authors should\nconsider adding quantifications to this figure.\n10. Add protein ladders to all blots.\n11. In Fig. 4i, the authors should include data from experiments using TIGAR knockout and control mice under the same\nexperimental conditions. This comparison would help clarify whether the interaction between MUC2 precursors and AGR2\nis dependent on TIGAR.\n12. In Extended Data Fig. 6b, the authors should complement the current data by performing immunoprecipitation of MUC2\nfollowed by immunoblotting for AGR2. This reciprocal approach would provide stronger evidence for a direct interaction\nbetween MUC2 and AGR2.\n13. In Extended Data Fig.7, it’s not convincing that TIGAR deletion increases the SNO-Trx1 band from the data presented.\n14. Extended Data Fig.7h is pretty cool data.\n15. In Fig. 5, the authors should include immunoblotting of MUC2 under Trx1 perturbation conditions to further support the\nconclusion that Trx1-mediated S-nitrosylation of AGR2 acts upstream of MUC2 maturation.\n16. The authors should add a model figure to help navigate the TIGAR-lactate-6PGD/ G6PD-Trx1-AGR2-MUC2 signaling\naxis, which is a pretty interesting mechanism.\nMinor comments:\n1. Lines 133-135: “we generated mice with conditional knockout of TIGAR in the intestinal tract (TIGARf/fVil1-Cre) by\ncrossing TIGARf/f mice with Vil1-Cre mice (Fig. 1k)”. Data in Fig 1k doesn’t match with texts.\n2. In Fig. 1i, I can’t find anywhere how the authors quantified NO from their samples.\n3. Line 203: “we performed RNA sequencing on the colons of the TIGARf/fVil1-Cre mice”. Please add TIGARf/f mice as well."}, "author_response": "Dear Reviewers and Editor,\nThank you for your letter and the reviewers’ meticulous review and professional\ncomments on our manuscript entitled “TIGAR Regulates Intestinal Mucus Barrier\nIntegrity by Inhibiting Lactylation of G6PD/6PGD in Ulcerative Colitis” (ID:\nNCOMMS-25-24116-T). We have completed the revision in full accordance with the\nrequirements of the editorial department, and revisions in the text are shown using red\nhighlight, including: (1) We have provided explanations for each of the questions raised\nby the reviewers. (2) We have supplemented the animal experiments to make our data\nmore convincing. (3) We revised vague and non-specific descriptions in the article\nbased on the reviewers' comments. (4) We have supplemented and modified the\nexperimental methods in the manuscript according to the suggestions of the reviewers.\n(5) We have made the modifications according to the Nature Communications\nformatting instructions.\nWe sincerely appreciate the valuable feedback provided by reviewers. Two respected\nreviewers not only pointed out the shortcomings of our research, but also provided\ncorresponding improvement methods. Under his/her guidance, we successfully\ncompleted the supplementary experiment and revised our paper, which greatly\nimproved the quality of the manuscript. For that, we express our deep gratitude and\nrespect.\nThe following summarizes what we responded to reviewer comments.\nReviewer #1:\nThis study revealed that TIGAR reduces 6-phosphogluconate and lactate\nlevels by reprogramming the pentose phosphate pathway (PPP) and glycolysis. Its\ndeficiency exacerbates oxidative stress by promoting the lactylation of G6PD and\n6PGD. Specifically, lactylation at lysine residues K432 in G6PD and K38 in 6PGD\nimpairs NADPH synthesis. This impairment induces S-nitrosylation of AGR2,\ndisrupts MUC2 maturation, and subsequently damages the intestinal mucus\nbarrier in ulcerative colitis (UC) mouse models. Overall, this work elucidates the\nrole of TIGAR in UC pathogenesis and establishes a compelling connection\nbetween TIGAR and G6PD/6PGD through high-quality data and well-designed\nexperiments. However, several key concerns should be addressed prior to\npublication:\nReply: We greatly appreciate your positive evaluation and valuable suggestions.\nYour insightful comments helped us identify certain limitations in our manuscript.\nUnder your guidance, we have conducted additional experiments to address these\nconcerns, thereby strengthening the rigor and persuasiveness of our study. We are\ndeeply grateful for your constructive feedback. Below, we provide point-by-point\nresponses to your comments.\n1. Human Tissue Validation: It would be valuable to include analysis of human UC\nbiopsy samples to validate whether TIGAR expression correlates with disease\nseverity and whether G6PD/6PGD lactylation is elevated in UC patients. This\nwould significantly strengthen the translational relevance of the findings.\nReply: We fully agree with your viewpoint that our study would be significantly\nstrengthen the translational relevance if supported by clinical data. Actually, our overall\nexperimental design was to first conduct animal studies to confirm the role and\nunderlying mechanisms of TIGAR, followed by clinical investigations. Your\nprofessional comment highlights that our original study design could be further\nimproved. Indeed, the manuscript would be more convincing if supported by\npreliminary clinical sample analysis. However, in China, the process for ethical\napproval is complex and time-consuming, requiring at least three months. Patient\nrecruitment, biospecimen collection, laboratory testing, and data analysis require a\nminimum of six months to complete, it extended the time required for manuscript\nrevision considerably. We therefore plan to initiate the corresponding clinical research\nas soon as possible after publication of this work. Although current conditions do not\npermit us to conduct clinical studies, your suggestion has prompted us to consider\nwhether clinical evidence supporting our findings could be obtained through alternative\\approaches. To this end, we have surveyed some available databases and carried out a\nfurther literature search.\nBased on the data we have obtained, TIGAR is a novel glycolytic regulatory\nenzyme, and current research in UC patients remains limited to genetic-level, with a\nlack of the protein levels and functional study. Two ulcerative colitis gene expression\nprofile datasets (GSE11223 and GSE47908) were obtained from the GEO database and\nanalyzed to assess TIGAR expression changes. We found that TIGAR expression in\nUC patients did not show significant alteration in GSE11223. These data differ from\nour findings, which may be attributed to species differences or to the fact that the data\noriginate from different stages of the disease. However, through the analysis of\nGSE47908, we found that TIGAR was down-regulated in colonic mucosal biopsy from\npatient with ulcerative colitis compared with the control individual. Although species\ndifferences exist, this clinical data partly supports our animal findings that\ndownregulation of TIGAR in UC is a key factor contributing to impaired antioxidant\ncapacity in intestinal tissues.\nRegarding the issue of G6PD/6PGD lactylation modifications in UC patients,\nthere are no reports available either in databases or in the literature. This lack of existing\ndata indicates that our study is a worthwhile area for exploration in elucidating UC\npathogenesis and identifying potential therapeutic targets. Although, due to current\nlimitations, we are unable to conduct related clinical studies in the near term, your\nsuggestion has identified an important avenue for future research. On behalf of all\nauthors, I would like to express our sincere gratitude to you.\nThe expression of TIGAR in GSE47908.\n2. Functional Validation of Lactylation Sites: While the study identifies K432\n(G6PD) and K38 (6PGD) as critical lactylation sites, direct functional evidence is\nrequired to confirm that lactylation at these specific sites directly impairs enzyme\nfunction. Enzymatic activity assays comparing wild-type (WT) proteins to non-\nlactylatable mutants (e.g., lysine-to-arginine mutants) would provide this critical\nevidence.\nReply: Thank you for your suggestions. We acknowledge that this point was\noverlooked in our manuscript, which represents a limitation in our work. We have\nsupplemented this experiment. We re-expressed wild-type and mutant G6PD (K432R)\nand 6PGD (K38R) in cells with knocked down G6PD and 6PGD. The results confirmed\nthat the lactylation site do indeed affect the enzymatic activities of G6PD and 6PGD.\nThe results were shown in new Fig. 3 m and q. Wild-type and mutant G6PD (WT and\nK432R) or 6PGD (WT and K38R) were re-expressed by same sense mutation in the\ncells with G6PD or 6PGD knocked down, the results showed that lactate was found to\ninhibit the activity of G6PD and 6PGD (Fig. 3m and q). However, when the lactylation\nsite K432 and K38 were mutated, the enzymatic activities of G6PD and 6PGD were\nrestored. Thank you once again for your valuable suggestions, which have significantly\ncontributed to enhancing the conclusion of our article.\nFig. 3m, q The activities of G6PD and 6PGD are enhanced following mutation of the\nlactylation site.\n3. Mechanism of Lactylation Induction: The hypothesis that lactate accumulation\ndrives G6PD/6PGD lactylation requires experimental validation. Pharmacological\ninhibition of lactate dehydrogenase (LDH) could test whether reducing lactate\nlevels rescues enzyme activity. Furthermore, investigating the potential\ninvolvement of the glyoxalase pathway and enzymes like GLO2/S-\nlactoylglutathione (SLG) hydrolase in this lactylation is warranted, given recent\nfindings that SLG can induce D-lactylation in immune cells.\nReply: Thank you for your professional comments. Following your suggestion, we\ninhibited lactate dehydrogenase (LDH) activity and found that the lactylation levels of\nG6PD and 6PGD were significantly decreased, while their enzymatic activities were\nincreased (Fig. 3n, r). These results demonstrate that lactylation indeed suppresses the\nactivity of these two enzymes.\nAdditionally, we verified the D-lactylation levels of G6PD and 6PGD in cells and\ntissues. Under normal conditions, low levels of D-lactate modification are present, but\ntheir levels do not show marked changes following treatment with TNF-α or DSS.\nThis indicates that the enhanced protein lactylation in UC is primarily driven by increased\nL-lactate level, rather than D-lactate. Although we did not further investigate the\nunderlying mechanisms due to the lack of significant changes in D-lactylation, your\ncomment has prompted us to reflect on our study design. We recognize that our previous\napproach was not fully comprehensive. We sincerely thank you for your professional\nand insightful suggestion, which will guide our future work. The results are presented\nin the figure below:\nFig. 3n, r The activities of G6PD and 6PGD increase following the inhibition of LDH\nThe D-lactylation changes of G6PD and 6PGD in TNFα-treated cells and UC colon tissues.\n4. Figure Labeling Consistency:There are inconsistencies in figure labeling:\nIn Figure 2, the letter labels for subfigures show variations in capitalization (e.g.,\n\"A\" vs. \"a\").\nIn Extended Data Figure 8, the subfigure labels (e.g., a, b, c) display variations in\nfont size.\nReply: Thank you for your reminder, we sincerely apologize for these clerical\nerrors, we have made corrections to these imprecise parts in Figure 2 and\nSupplementary Figure 8.\nReviewer #2:\nIn this study, the authors aimed to characterize how TIGAR modulates glucose\nmetabolism in the context of ulcerative colitis. They combined a variety of\napproaches including classical biochemistry, mass spectrometry-based\nmetabolomics and proteomics, stable isotope tracing, RNA sequencing, and mouse\nphysiology, and uncovered a signaling axis whereby TIGAR deletion elevated\nlactylation of key pentose phosphate pathway (PPP) enzymes, leading to reduced\nTrx1 activity, decreased S-nitrosylation of AGR2, and impaired mucin maturation.\nThe study is very interesting, and the data are generally of reasonable quality.\nA major concern with the current version of the manuscript is the lack of direct\nevidence demonstrating that AGR2 and Trx1 are S-nitrosylated, and that 6PGD\nand G6PD are lactylated in vivo, particularly in disease-relevant mouse tissues.\nWhile the cell-based data are convincing, validating these post-translational\nmodifications in primary tissues would significantly strengthen the manuscript\nand enhance its suitability for publication in Nature Communications. See detailed\ncomments below:\nReply: We sincerely thank you for your supportive comments and constructive\nsuggestions, which have been instrumental in improving the quality of our manuscript.\nWe are deeply impressed by your profound expertise and scholarly rigor. Your\ncomments reflect an exceptional ability to grasp both the overall framework and the\nfine details of scientific research, which we truly admire. You sharply pointed out a\ncentral limitation in our study: indeed, the study of post-translational modifications of\nkey proteins and their functional implications should not be limited to cell-based\nexperiments alone. We fully agree with this assessment.\nIn response, we have focused our additional experiments on validating, in vivo\nusing UC mouse models, the post-translational modifications of G6PD, 6PGD, Trx1,\nand AGR2, as well as their protective effects on the intestinal mucus barrier. These new\ndata have substantially strengthened the rigor and credibility of our conclusions. On\nbehalf of all authors, I would like to express our sincere and heartfelt gratitude for your\ninsightful guidance, which has greatly improved the quality of our work.\n1. In Extended Data Fig. 1a, c, the H&E staining does not seem to show a reduction\nof crypt height by half, as stated. Also, it is unclear whether the data points in\nExtended Data Fig. 1c represent measurements from different mice or multiple\nsections from the same mouse. Please clarify this in the figure caption.\nReply: Thank you for your careful observation and highly insightful question. First,\nall figures in this study, including but not limited to Supplementary Data Fig. 1a, 1c,\nwere derived from different mice specimens. In this study, for each mouse, we measured\nthe lengths of six consecutive crypts and calculated the average. Each data point in\nFigure 1c represents the mean crypt length from one tissue section of an individual\nmouse. The measurement strategy and rationale have been described in the Methods\nsection (line 557-561), and further clarified in the figure legend.\nSecond, we sincerely appreciate your astute observation. Indeed, in the\nrepresentative images we provided, the degree of crypt atrophy does not reach 50%.\nHowever, quantitative analysis of the full dataset confirms that crypt length is reduced\nby half, reaching the level of 50% atrophy. The apparent discrepancy between the\nrepresentative images and the statistical results arises because the images were selected\nto emphasize changes in the mucus layer, which was the primary focus of this study,\nrather than to illustrate the full extent of crypt atrophy. This approach aligns with the\ncentral aim of our research: to investigate mucus barrier dysfunction in UC. Notably, in\nsamples with severely damaged crypts, the mucus layer is completely absent. To enable\nreaders to better observe the differences in mucus synthesis, we selected well-stained\ntissue sections that clearly show mucus production.\nRepresentative images of HE staining that used to analyze the changes in the length of the\ncrypts of different mice.\n2. Lines 137–139: “These changes appeared two days earlier and more serious\npathological damage in TIGARf/fVil1-Cre mice compared to TIGARf/f mice” —\nIf the authors wish to argue that TIGAR deletion accelerates the onset and\nprogression of colitis, they should provide comparative data at earlier time points\n(e.g., day 5 vs. day 7 of DSS treatment) between TIGARf/fVil1-Cre and TIGARf/f\nmice.\nReply: Thank you for this suggestion. We agree that added the comparative data\non day 5 and day 7 would help strengthen our conclusion that TIGAR deletion\naccelerates the onset and progression of colitis.\nIn fact, we have generated comparative data for Day 5 and Day 7. We acknowledge\nthat presenting these results in separate figures may have contributed to potential\nmisinterpretations. As shown in Supplementary Fig. 1a, we already assessed mucus\nsynthesis after 7 days of DSS treatment and observed a significant reduction in\nTIGARf/fVil1-Cre mice compared to TIGARf/f controls. Moreover, in Fig. 1a, after 5\ndays of DSS treatment, wild-type mice showed only a slight decrease in intestinal\nmucus synthesis, whereas TIGAR-knockout mice exhibited significant changes at this\ntime point. These results support our conclusion that intestinal injury in TIGAR-\nknockout mice occurs approximately two days earlier than in control mice and is\naccompanied by more severe pathological damage.\nAs suggested, we have now incorporated the 7-day DSS treatment results into\nFigure 1a to provide a clearer and more intuitive comparison between genotypes across\ntime points, thereby improving the coherence and persuasiveness of our findings.\nFig. 1a Colon HE staining (left) and AB-PAS staining (right)\n3. In Fig. 2a, the authors should label all metabolites related to the PPP in the\nvolcano plot. While the KEGG pathway analysis highlights PPP as the most\nsignificantly regulated pathway, visualizing the changes in individual PPP\nmetabolites would enhance the interpretability of the data. In Extended Data Fig.\n3a, the main text refers this data as being from HT29 CL.16E cells, but the legend\nsays it’s from TIGARf/fVil1-Cre mice. Please clarify.\nReply: We sincerely apologize for the lack of clarity in our manuscript. We have\nmarked all the metabolites related to PPP in Fig. 2a. In Supplementary Fig. 3a, the data\nis from HT29 CL.16E cells, and we have made the correction in the legend.\nFig. 2a Changes in PPP metabolites\n4. In Fig. 2h, the data clearly show increased lactate production in TIGAR-\ndeficient cells. However, based on the current data, the authors cannot conclude\nthat lactate consumption is also increased.\nReply: Thank you for your valuable feedback. We thoroughly reviewed the data\nand indeed could not conclude that lactate consumption was increased. To investigate\nlactate consumption, we performed a two-way repeated measures ANOVA for\nunlabeled lactate (M+0) from Figure 2h. The analysis revealed no statistically\nsignificant difference between the groups (p = 0.1622). Consequently, the previous\nstatement suggesting that lactate consumption was increased in TIGAR-deficient cells\nis not supported by the data. We sincerely apologize for the lack of rigor in our work.\nWe have revised the corresponding section in the main text to ensure a more accurate\nrepresentation (lines 195-198). Thank you again for your insightful comment.\nThe changes in unlabeled lactate levels over time in TIGAR-deficient cells\n5. In Extended Data Fig. 4, please describe how many genes were changed.\nReply: There are 1261 genes were changed, we have made a detailed explanation\nin the line 204-211 of main text.\n6. In Fig. 3, it is critical to demonstrate that 6PGD K38 and G6PD K432 are indeed\nlactylated in vivo. The authors should perform pull-down of these two enzymes\nfrom mouse tissue followed by mass spectrometry–based proteomic analysis to\nprovide direct evidence of site-specific lactylation in vivo. Also, the authors should\nprovide proteomic evidence of S-nitrosylation of AGR2 in mouse tissues in Fig. 4\nand also Trx1 if possible.\nReply: Among all the questions you have raised, this is the most central and\ninsightful. Your insightful observations regarding the limitations in our current study\nwill greatly assist us in refining the experimental design and strengthening the overall\ncredibility of our manuscript. We fully concur with your recommendation that\ninvestigating protein modifications of G6PD, 6GPD, Trx1, and AGR2 solely at the\ncellular level is insufficient, and that validation through in vivo studies using animal\nmodels is essential. Furthermore, we maintain that supplementary experiments should\nnot only validate the occurrence of corresponding protein modifications in key proteins\nfrom UC animals, but also confirm the novel modification sites identified in this study\nthrough functional validation of their biological effects. To address the limitations of\nprevious research, a series of animal experiments was conducted.\nIn terms of research methods, we have made appropriate adjustments based on\npast experience and research objectives. The approach you proposed that extracting and\npurifying G6PD and 6PGD from animal tissues followed by proteomic analysis to\nconfirm the presence of lactoylation modification, is indeed a well-established method\nfor validating such post-translational modifications in vivo. However, this approach\ndoes not enable the assessment of functional consequences associated with these\nmodifications on the target proteins. Moreover, during the protein purification process,\nmultiple chromatography, elution and centrifugation are required. The physical and\nchemical factors in the experimental process are very likely to cause the modification\ngroups already bound to the proteins to fall off, resulting in false negative results. We\nhave encountered this phenomenon in our previous cell experiments. If the proteins are\nextracted and purified from tissues, the possibility of false negatives is estimated to be\neven greater.\nBased on all these considerations, we adopted another method. Using the adeno-\nassociated virus 9 (AAV9), the wild-type and mutant forms of 6PGD (WT and K38R)\nand G6PD (WT and K432R-K45R) were injected intraperitoneally into TIGARf/fVil1-\nCre mice. Four weeks later, the UC model was established by DSS. As shown in Fig.\n7, the K432 and K38 mutations effectively prevented lactylation of G6PD and 6PGD\n(Fig. 7g, h), respectively, and significantly restored their enzymatic activities in\nTIGAR-knockout mice (Fig. 7e, f). In contrast, the K45 mutation had no significant\neffect on G6PD activity (Fig. 7e). Further investigation revealed that either the K432\nor K38 mutations significantly reduced S-nitrosylation levels of Trx1 and AGR2, and\nmarkedly enhanced mature MUC2 synthesis (Fig. 7g, h). Morphological analysis\nshowed that mutation of G6PD at K432 or 6PGD at K38 ameliorated colitis-associated\nintestinal damage, enhanced mucus production, preserved mucosal layer integrity, and\nmarkedly reduced bacterial load in the inner mucus layer (Fig. 7a-d).\nGiven that AGR2 contains only one cysteine residue, its S-nitrosylation\nmodification site must be cysteine 81. In the case of Trx1, our data demonstrated that\nthe S-nitrosylation occurs at cysteine 73, which is consistent with previous reports[1-\n3]. These studies further support the reliability of our experimental findings.\nFinally, we sincerely thank you for your valuable comments. Your feedback\nhighlights the importance of in vivo experiments and will guide us in improving the\ndesign of experimental strategies in our future studies.\nFig. 7 Expression of WT and mutant G6PD and 6PGD in TIGAR-KO mice alleviates DSS-\ninduced colitis.\n[1] Nagarajan N, Oka SI, Nah J, et al. Thioredoxin 1 promotes autophagy through transnitrosylation\nof Atg7 during myocardial ischemia. J Clin Invest. 2023;133(3):e162326.\n[2] Wu C, Parrott AM, Liu T, et al. Distinction of thioredoxin transnitrosylation and denitrosylation\ntarget proteins by the ICAT quantitative approach. J Proteomics. 2011;74(11):2498-509.\n[3] Li H, Wan A, Xu G, et al. Small changes huge impact: the role of thioredoxin 1 in the regulation\nof apoptosis by S-nitrosylation. Acta Biochim Biophys Sin. 2013;45(3):153-61.\n7. In Fig. 3p, it’s not convincing that 6PGD K38R increases the MUC2 band from\nthe data presented.\nReply: Thank you for your reminder, we have replaced the image with a more\nrepresentative one.\nRepresentative images of MUC2 changes after 6PGD lactylation site mutation.\n8. Extended Data Fig. 4f and g are missing.\nReply: Thank you for your reminder. We have added the symbol f and g in\nSupplementary Figure 4 and made the corrections in the line 217-218 of the main text.\n9. In Extended Data Fig. 6d, it’s very hard to tell if MUC2 and ARG2 colocalize\nfrom the staining. The authors should consider adding quantifications to this\nfigure.\nReply: Thank you for your suggestion, we have added the quantification. In the\nprevious version, due to an oversight on our part, only mature mucus-producing cells\nwere annotated, whereas cells exhibiting co-localization were not indicated. In the\nrevised Supplementary Figure 6e, the cells exhibiting co-localization of MUC2 and\nAGR2 in the merged image appear yellow. We have annotated the cells with co-\nlocalization using yellow arrows and further confirmed the co-localization of MUC2\nand AGR2 in situ by quantitatively analyzing the co-localization coefficient, the\nPearson’s correlation coefficient are 0.91851 and the Manders’ Overlap coefficient are\n0.9631 (Supplementary Fig. 6f).\nSupplementary Fig. 6e, f The co-localization analysis of MUC2 and ARG2 in colon tissue.\n10. Add protein ladders to all blots.\nReply: We have added the protein ladders to all blots.\n11. In Fig. 4i, the authors should include data from experiments using TIGAR\nknockout and control mice under the same experimental conditions. This\ncomparison would help clarify whether the interaction between MUC2 precursors\nand AGR2 is dependent on TIGAR.\nReply: Thank you for your suggestion, we admittedly overlooked this aspect, we\nhave added this experiments. The results in the new Fig. 4i and j showed that the\ninteraction between MUC2 precursors and AGR2 was weakened in UC and TIGAR\nknockout mice compared with their respective control groups. This demonstrated that\nthe interaction is dependent on TIGAR.\nFig. 4i, j The interaction between MUC2 precursors and AGR2 was weakened in UC and\nTIGAR knockout mice.\n12. In Extended Data Fig. 6b, the authors should complement the current data\nby performing immunoprecipitation of MUC2 followed by immunoblotting for\nAGR2. This reciprocal approach would provide stronger evidence for a direct\ninteraction between MUC2 and AGR2.\nReply: According to your suggestion, we have conducted and included this\nexperiment. The results are presented in the newly added Supplementary Fig. 6b and\nc. As you pointed out, this reciprocal approach provides stronger evidence for a direct\ninteraction between MUC2 and AGR2.\nSupplementary Fig. 6b, c AGR2 interacts with MUC2.\n13. In Extended Data Fig.7, it’s not convincing that TIGAR deletion increases the\nSNO-Trx1 band from the data presented.\nReply: Thank you for your careful observation and highly insightful question. To\nfacilitate a more intuitive understanding of the differences, a quantitative analysis was\nconducted on the data presented in Supplementary Figure 7f and g. The results\nshowed that the nitrosylation of Trx1 was significantly upregulated after the knockout\nof TIGAR in mice and cells (Supplementary Figure 7f-h, m).\nSupplementary Fig. 7f-h TIGAR knockout aggravates Trx1 S-nitrosylation.\n14. Extended Data Fig.7h is pretty cool data.\nReply: Thanks for your affirmation.\n15. In Fig. 5, the authors should include immunoblotting of MUC2 under Trx1\nperturbation conditions to further support the conclusion that Trx1-mediated S-\nnitrosylation of AGR2 acts upstream of MUC2 maturation.\nReply: Thank you for your suggestion, we have added the experiments. As showed\nin the new Fig. 5, mutating Cys32 or Cys35 reduced SNO-protein (Fig. 5e, g), SNO-\nTrx1 (Fig. 5h), and SNO-AGR2 (Fig. 5f, i), while simultaneous mutation of Cys32 and\nCys35 increased their S-nitrosylation (Fig. 5e-i), and reduced the synthesis of MUC2\n(Fig. 5i). Mutating Cys73 significantly reduced their S-nitrosylation levels (Fig. 5g-i)\nand increased the synthesis of MUC2 (Fig. 5i), indicating Cys32 and Cys35 are\ninvolved in denitrosylation, while Cys73 is critical for transnitrosylation. Next, we\nfound that NADPH addition after TIGAR deletion or Trx1 transnitrosylation inhibition\nwith PX12 reduced AGR2 S-nitrosylation and increased the synthesis of MUC2 (Fig.\n5j). NADPH had no effect on AGR2 S-nitrosylation and the synthesis of MUC2 after\nTrx1 knockdown or simultaneous mutation of Cys32 and Cys35 (Fig. 5k, l), indicating\nNADPH modulates AGR2 S-nitrosylation by altering Trx1 reductase activity. Thank\nyou again for your valuable suggestions, which have significantly contributed to\nenhancing the conclusion of our article.\nFig. 5e-l TIGAR deletion facilitates the Trx1-mediated S-nitrosylation of AGR2.\n16. The authors should add a model figure to help navigate the TIGAR-lactate-\n6PGD/ G6PD-Trx1-AGR2-MUC2 signaling axis, which is a pretty interesting\nmechanism.\nReply: We have added the model figure. As shown in Fig. 7i, it’s the schematic\nillustration of TIGAR maintains the intestinal mucus barrier by inhibiting lactate\nproduction, reducing lactylation of G6PD and 6PGD, promoting NADPH synthesis,\ninhibiting nitrosylation of Trx1 and AGR2, and facilitating MUC2 maturation.\nFig. 7i Schematic illustration of TIGAR maintains the intestinal mucus barrier by inhibiting\nlactate production\nMinor comments:\n1. Lines 133-135: “we generated mice with conditional knockout of TIGAR in the\nintestinal tract (TIGARf/fVil1-Cre) by crossing TIGARf/f mice with Vil1-Cre mice\n(Fig. 1k)”. Data in Fig 1k doesn’t match with texts.\nReply: We sincerely apologize for the lack of clarity in our manuscript, we have\nadded a schematic diagram about the generation of mice with conditional knockout of\nTIGAR in the intestinal tract. The 135-136 line in the main text has also been modified.\nAnd the schematic diagram was shown in new Fig. 1j.\nFig. 1j Intestinal TIGAR deletion aggravates colitis via oxidative stress in UC mice.\n2. In Fig. 1i, I can’t find anywhere how the authors quantified NO from their\nsamples.\nReply: Thank you for your reminder, we apologize for overlooking the detailed\nmethods. Based on your suggestion, we have added the experimental procedures in line\n571-581 of the materials and methods section.\n3. Line 203: “we performed RNA sequencing on the colons of the TIGARf/fVil1-\nCre mice”. Please add TIGARf/f mice as well.\nReply: Thank you for your reminder, we have already made modifications to line\n203-204 of the text."}, {"round": 2, "reviewer_comments": {"reviewer_1": "Thanks for your rely.No more questions.", "reviewer_2": "The authors have satisfactorily addressed all my concerns. The manuscript is now of high quality and suitable for\npublication. I recommend acceptance."}, "author_response": ""}]

Decision letter