d -Allulose Ameliorates Hyperglycemia Through IRE1α Sulfonation-RIDD- Sirt1 Decay Axis in the Skeletal Muscle

Abstract

Aims:

The skeletal muscle maintains glucose disposal via insulin signaling and glucose transport. The progression of diabetes and insulin resistance is critically influenced by endoplasmic reticulum (ER) stress. d-Allulose, a low-calorie sugar substitute, has shown crucial physiological activities under conditions involving hyperglycemia and insulin resistance. However, the molecular mechanisms of d-allulose in the progression of diabetes have not been fully elucidated. Here, we evaluated the effect of d-allulose on hyperglycemia-associated ER stress responses in human skeletal myoblasts (HSkM) and db/db diabetic and high-fat diet-fed mice.

Results:

d-allulose effectively controlled glycemic markers such as insulin and hemoglobin A1c (HbA1c), showing anti-diabetic effects by inhibiting the disruption of insulin receptor substrate (IRS)-1 tyrosine phosphorylation and glucose transporter 4 (GLUT4) expression, in which the phosphatidylinositol-3 kinase (PI3K)/protein kinase B (Akt) pathway is involved. The levels of glucose dysmetabolism-based NADPH oxidase, such as NADPH-dependent oxidoreductase (Nox) 4, were highly increased, and their interaction with IRE1α and the resultant sulfonation-regulated IRE1-dependent decay (RIDD)-Sirt1 decay were also highly increased under diabetic conditions, which were controlled with d-allulose treatment. Skeletal muscle cells grown with a high glucose medium supplemented with d-allulose showed controlled IRE1α sulfonation-RIDD-Sirt1 decay, in which Nox4 was involved.

Innovation and Conclusion:

The study observations indicate that d-allulose contributes to the muscular glucose disposal in the diabetic state where ER-localized Nox4-induced IRE1α sulfonation results in the decay of Sirt1, a core factor for controlling glucose metabolism. Antioxid. Redox Signal. 37, 229–245.

Introduction

Insulin resistance in the skeletal muscles is a crucial factor contributing to glucose dysmetabolism in type 2 diabetes mellitus. Transportation of glucose in the skeletal muscles is driven by the activation of phosphatidylinositol-3 kinase (PI3K)/protein kinase B (Akt), which essentially participates in the translocation of glucose transporter 4 (GLUT4) from the intracellular pool to the plasma membrane. The impairment of GLUT4 translocation or reduced GLUT4 activity in the muscles results in systemic insulin resistance (52, 56), and overproduction of reactive oxygen species (ROS) is a key triggering factor (26). The endoplasmic reticulum (ER), mitochondria, and NADPH oxidase are responsible for excessive ROS production, and superoxide production in adipocytes and myotubes is a typical characteristic observed in insulin resistance mouse models (21, 57).

Innovation

The study observations provide strong evidence to support a preclinical rationale and molecular basis to explore potential therapeutic applications of oral d-allulose as a simple, effective, and safe treatment for obesity, diabetes, and other metabolic syndromes.

An increase in the activity of NADPH oxidase enhances the production of superoxides, leading to an increase in oxidative stress with hyperglycemia. NADPH oxidase-associated ROS are assumed to be the cause of the reduced insulin responsiveness (14, 44), ER dysfunction, and ROS production in the ER (4, 57). The ER is an important cellular organelle that facilitates protein folding. The chaperone-based electron transfer system is directly involved in this process (48, 53).

Unlike the adaptive ER stress response, redox imbalance or ROS accumulation is linked to prolonged or pathological ER stress, which influences post-translational modification of proteins and subsequent modifications of protein function (11, 20, 51). Protein dysfunction is influenced by sulfonation, which involves the oxidation of specific cysteine residues mediated by endogenous or exogenous ROS (23). ER-derived H₂O₂ is produced from NADPH-dependent oxidoreductase (Nox)4 and leads to ER stress (54). IRE1α sulfonation is activated by the Nox4-ER-ROS axis and is linked to cellular dysfunction (25).

The IRE1α post-translational modification-linked signaling axis, known as regulated IRE1-dependent decay (RIDD), has diverse substrates, including proteins associated with glucose metabolism. d-Allulose is a low-calorie C-3 epimer of D-fructose found naturally in vegetables and fruits and exhibits 70% of the sweetness of sucrose (2, 37). Previously, several studies have demonstrated the influence of d-allulose on lipid metabolism. One study showed that the action of d-allulose is similar to that of D-glucose; however, it stimulates glucose uptake from the liver while suppressing hepatic lipogenic enzyme activity (2).

Another study showed that d-allulose reduces intra-abdominal fat accumulation and subdues blood glucose levels during glucose loading (24). Most previous studies conducted with animal models demonstrated beneficial effects of d-allulose on body weight, fat mass, and energy intake (36, 39). Moreover, the skeletal muscle is a key site of glucose metabolism, and d-allulose might potentially act as a strong activator of glucose transport in the skeletal muscle, in addition to its essential role in adipocytes and hepatocytes. However, the effects of d-allulose on the skeletal muscles have not been completely elucidated. The control of insulin resistance and associated insulin signaling, including GLUT4, mediated by d-allulose, has been recently studied; however, glucose dysmetabolism-based ROS signaling was not examined (29, 50).

Moreover, animal models efficiently demonstrate impaired glucose tolerance when subjected to a glucose challenge test, and these models are well accepted for investigating type II diabetes. However, compared with mice models showing high blood glucose levels (>500 mg/dL), blood sugar levels of diabetic patients are consistently higher than the target range (200–350 mg/dL in adults and 200–240 mg/dL in children) (42).

Typically, patients manage their blood sugar level to stay within the optimal range and make all possible efforts to adjust the levels with drugs. Physiological differences may also be present in animals. However, db/db and high-fat diet (HFD)-fed mice models show characteristics of overt insulin resistance, progressive development of hyperglycemia, and β cell dysfunction with age. These models have been considered clinically more relevant than other animal models. In addition, muscle cells can consume up to 70% of glucose, indicating the suitability of the cells for use as in vitro diabetes models. Thus, we applied db/db and HFD-fed mice and human skeletal myoblasts (HSkM) for in vitro investigations. These models are relatively relevant for exploring the molecular mechanism underlying the anti-diabetic effect of d-allulose mediated against Nox-IRE1α sulfonation-RIDD.

Results

d-Allulose controls glucose metabolism in db/db mice

Diabetic db/db mice were administered various doses of d-allulose in different formulations, namely, allulose powder (AP), allulose liquid (AL), or vehicle to evaluate glucose homeostasis and insulin sensitivity. The two formulations did not affect the weekly average weight gain (Supplementary Fig. S1a) and food intake (Supplementary Fig. S1b) in db/db mice.

Regarding post-prandial and fasting blood glucose levels, mice that were administered d-allulose in powder and liquid formations (5.16 g/kg, high dose) for 8 weeks demonstrated a significant reduction in glucose levels compared with mice who received low and medium doses (Supplementary Fig. S2a, b). d-Allulose treatment demonstrated improvements in post-bolus glucose clearance than those observed in the vehicle-treated db/db mice when analyzed at several predetermined time points during the oral glucose tolerance test (OGTT) (Fig. 1a and Supplementary Fig. S2c). Similarly, during the insulin tolerance test (ITT), d-allulose treatment showed reduced blood glucose levels compared with treatment with vehicle (Fig. 1b and Supplementary Fig. S2d).

FIG. 1.

Effects of d-allulose on blood glucose levels and activation of the PI3K/Akt signaling pathways in db/db mice. Male db/db mice received vehicle or d-allulose (1.02, 3.07, or 5.16 g/kg) once daily for 8 weeks via oral gavage. At the end of 7th week, OGTT (a) and ITT (b) were performed. (c, d) At the end of the study, insulin and HbA1c concentrations were analyzed. (e) Tibialis anterior muscle lysates were immunoprecipitated with p-Tyrosine antibody and immunoblotted against IRS1/2. (f) Quantitative analysis of protein expression was also performed. (g) Immunoblotting was performed by using p-Akt. (Ser473), Akt, GLUT4, and β-actin antibodies. (h) Quantitative analysis of protein expression was also performed. (i) Immunohistochemical staining was performed by using antibodies against GLUT4 (green) and the plasma membrane marker dystrophin (red). DAPI nuclear staining is shown in blue. Representative images (merged) showing colocalization of GLUT4 with the plasma membrane marker dystrophin. (j) The quantification of colocalization of GLUT4 with dystrophin was performed in the Materials and Methods section. Scale bar: 100 μm. Data are represented as mean ± SEM, n = 8 animals/group. *p < 0.05 versus db/+, ^# p < 0.05 versus db/db+vehicle. Akt, protein kinase B; AL, allulose liquid; AP, allulose powder; GLUT4, glucose transporter 4; H, high; HbA1c, hemoglobin A1c; IRS, insulin receptor substrate; ITT, insulin tolerance test; L, low; M, medium; OGTT, oral glucose tolerance test; PI3K, phosphatidylinositol-3 kinase; SEM, standard error of the mean.

Insulin levels and hemoglobin A1c (HbA1c) scores were markedly decreased in the d-allulose-treated db/db mice compared with those in the vehicle-treated db/db mice (Fig. 1c, d). d-Allulose-treated db/db mice demonstrated significantly increased serum adiponectin levels (Supplementary Fig. S2e). Further, d-allulose improved insulin signaling in the skeletal muscles of db/db mice, as indicated by increased tyrosine phosphorylation of insulin receptor substrate (IRS)-1 and IRS-2 (Fig. 1e, f).

Akt phosphorylation and its downstream GLUT4 expression were consistently increased in the d-allulose-treated db/db mice than those in the vehicle-treated db/db mice (Fig. 1g, h). Increased cytoplasm membrane localization of GLUT4 was also observed (Fig. 1i, j). These observations suggest that d-allulose improves systemic and peripheral insulin resistance, also recovering glucose disposal in the skeletal muscles under diabetic conditions.

d-Allulose regulates skeletal muscle oxidative stress and ER redox imbalance under hyperglycemic conditions

Accumulating evidence has demonstrated a significant association of the overproduction of ROS (primarily superoxide anions) with diabetes and associated complications (8, 43). We first examined ROS levels in the skeletal muscles of d-allulose-treated diabetic mice. Compared with non-diabetic mice, diabetic mice showed higher ROS levels in the skeletal muscles, which were reduced on administration of d-allulose in powder and liquid forms (Supplementary Fig. S3a, b). In addition, AP and AL treatment reduced the increase in lipid peroxidation and protein carbonylation observed in diabetic mice (Supplementary Fig. S3c, d).

Glucose metabolism-related ROS production is highly affected by the conversion of NADPH to NADP⁺, which is controlled by NADPH oxidase (55). Therefore, the NADP⁺/NADPH ratio and NADPH oxidase activity were examined in d-allulose-treated diabetic mice. As expected, the NADP⁺/NADPH ratio and NADPH oxidase activity were reduced on treatment with d-allulose (Fig. 2a, b). Nox2 and 4 were the most highly expressed Nox family proteins in diabetic mice (Fig. 2c).

FIG. 2.

d-Allulose regulates oxidative stress and ER redox imbalance in hyperglycemic conditions. Male db/db mice received vehicle or d-allulose (1.02, 3.07, or 5.16 g/kg) once daily for 8 weeks by oral gavage. Tibialis anterior muscle lysates were analyzed for NADP/NADPH ratio (a) and NADPH oxidase activity (b). (c) Expression of Nox1, Nox2, Nox3, Nox4, and β-actin in tibialis anterior muscle tissues. (d) Expression of Nox4 and β-actin in tibialis anterior muscle. (e) Quantitative analysis of protein expression was also performed. (f) Representative images of Nox4 staining in tibialis anterior muscle. Scale bar: 100 μm. (g) Subcellular fractions of lysates were analyzed by immunoblotting. Tibialis anterior muscle lysates were analyzed for lipid peroxidation level (h) and oxidized proteins by OxyBlot analysis (i) in the ER fraction. (j) Lysates were incubated with 1 mM diamide or left untreated for 15 min. Reduced and oxidized forms of PDI were detected by immunoblotting at predetermined time points. (k) Quantification of the levels of reduced PDI is shown. (l) Lysates were immunoprecipitated and immunoblotted with anti-Nox4, and IRE1α antibodies. (m) Quantitative analysis of protein expression was also performed. Data are represented as mean ± SEM, n = 8 animals/group. Input is the normalized IP band intensities (separately run). Scale bars, 20 μm. *p < 0.05 versus db/+, ^# p < 0.05 versus db/db+vehicle. ER, endoplasmic reticulum, IP, immunoprecipitation; Nox, NADPH-dependent oxidoreductase; PDI, protein disulfide isomerase.

The levels of these proteins were significantly increased in the vehicle-treated db/db mice than those in db/+ mice, whereas supplementation with d-allulose significantly reduced the expression of these proteins (Fig. 2d–f and Supplementary Fig. S3e, f). Subcellular fractional analysis revealed that Nox4 was prominently expressed in the ER fractions (Fig. 2g). In the ER fractions, lipid peroxidation and protein carboxylation were significantly decreased on supplementation with d-allulose (Fig. 2h, i).

The intra-ER redox state was examined by confirming the redox status of protein disulfide isomerase (PDI). The oxidized form of PDI continued to exist under diabetic conditions, whereas it was converted to a reduced form with ease under conditions involving washout in the control group (Fig. 2j, k). Notably, the conversion of PDI was recovered with d-allulose treatment. Since ER-localized Nox4 controls the ER redox balance with the primary ER stress-sensing protein IRE1α (33), we next examined the interaction of Nox enzymes with IRE1α.

As shown in Figure 2l and m, Nox4 is bound to IRE1α, and this interaction was inhibited on treatment with d-allulose, indicating that the Nox4 redox imbalance contributes to diabetic condition whereas d-allulose controls the phenotypes.

d-Allulose regulates the IRE1α sulfonation-RIDD-Sirt1 decay axis in skeletal muscles under hyperglycemic conditions

The ER stress response is a potential therapeutic target for chronic metabolic disorders, including diabetes (16). First, we determined the levels of ER stress markers (Fig. 3a, b). The levels of ER stress markers (p-IRE1α, GRP78, and CHOP) were markedly elevated in the vehicle-treated db/db mice than those in the vehicle-treated db/+ mice. We observed that phosphorylation of IRE1α, subsequent Xbp1 splicing, GRP78, and CHOP were inhibited in d-allulose-treated diabetic mice.

FIG. 3.

d-Allulose regulates the IRE1α sulfonation-RIDD-Sirt1 decay axis in hyperglycemic conditions. Male db/db mice received vehicle or d-allulose (1.02, 3.07, or 5.16 g/kg) once daily for 8 weeks through oral gavage. (a) Immunoblotting of p-IRE1α, IRE1α, GRP78, CHOP, and β-actin and RT-PCR of uXbp1, sXbp1, and GAPDH mRNA expression in tibialis anterior muscle tissue. (b) Quantitative analysis of protein and RNA expression were also performed. (c) Representative images showing irreversible sulfonation of IRE1α in tibialis anterior muscle tissue. (d) Quantitative analysis of protein expression was also performed. (e) Duolink PLA was used to identify protein interactions. Slides with tibialis anterior muscle tissues were incubated with primary antibodies (anti-mouse IRE1, 1:100; Santa Cruz, anti-rabbit cysteine sulfonate, 1:100; Abcam, , Cambridge, MA) in a humidity chamber overnight at 4°C, followed by hybridization with PLA probes (1 h/37°C), ligation (30 min/37°C), and amplification (2 h/37°C). Red dots indicate the location and extent of the interaction. Nuclei were stained with DAPI. Representative images showing protein–protein interactions in the tibialis anterior muscle tissue. HSkM were treated with H₂O₂ for 6 h and then used as a positive control. Negative control was developed without treatment with primary antibodies. (f) Quantification of the levels of IRE1α sulfonation is shown. (g) Heatmap depicting mRNA expression of the genes identified as RIDD substrates in tibialis anterior muscle tissue. (h) Predicted secondary structure of the IRE1α cleavage site in Sirt1 mRNA. The cleavage site is indicated by an arrowhead. In vitro cleavage assay and 1.5% denaturing agarose gel electrophoresis of Sirt1 substrate cleaved by IRE1α (i) and mutant Sirt1 mRNA (j). (k) The mRNA level of Sirt1 was assessed by RT-PCR. Data are expressed as mean ± SEM, n = 8 animals/group. Bar = 100 μm *p < 0.05 versus db/+, ^# p < 0.05 versus db/db+vehicle. HSkM, human skeletal myoblasts; mRNA, messenger RNA; PLA, proximity ligation assay; RIDD, regulated IRE1-dependent decay; RT-PCR, real-time polymerase chain reaction.

Among the ER stress response arms, IRE1α post-translational modifications, including phosphorylation, s-nitrosylation, sulfenylation, and sulfonation, play a crucial role in metabolic disorders (5). Especially, IRE1α sulfonation attributed to ROS production was reported to originate from an imbalance in the ratio of NADP⁺/NADPH and the associated activation of NADPH oxidase linked to glucose dysmetabolism (55). In addition, immunoprecipitation (IP) was performed with an anti-sulfonate antibody to further detect the oxidized form. IRE1α sulfonation (IRE1α:SO₃) was also increased under diabetic conditions, whereas d-allulose treatment significantly inhibited this modification (Fig. 3c, d).

Next, to assess the status of IRE1α sulfonation, we conducted a proximity ligation assay (PLA) using IRE1α antibody and cysteine-sulfonate with skeletal muscle samples. The interaction between IRE1α and cysteine-sulfonate observed in vehicle-treated db/db mice was attenuated in d-allulose-treated db/db mice (Fig. 3e, f). Next, the expression of IRE1α-RIDD target genes was analyzed. The expression of Blos1, Hgsnat, Col6, and Scara3, which are well-known RIDD genes, was reduced under diabetic conditions, whereas the messenger RNA (mRNA) levels were significantly recovered by treatment with d-allulose (Fig. 3g).

Sirtuins are known to exert a variety of effects on gluconeogenesis, glycolysis, insulin secretion, and sensitivity and could demonstrate therapeutic potential for a variety of metabolic diseases (38). The sirtuin gene family comprising SIRT 1–7 genes was screened as a target for RIDD using RNA structure, a web server for RNA secondary structure prediction. Among the Sirt family members, Sirt1 had a specific RIDD target sequence, “CUGCAG” (Fig. 3h and Supplementary Fig. S4).

To assess whether IRE1α sulfonation affects the decay of Sirt1, we performed an in vitro cleavage assay and found that the cleavage of Sirt1 mRNA by IRE1α peptide increased in a time-dependent manner (Fig. 3i, j and Supplementary Fig. S4), leading to impairment of IRE1α-mediated decay of Sirt1. Sirt1 mRNA expression was decreased under diabetic conditions and was recovered on treatment with d-allulose (Fig. 3k). The expression of SIRT1 protein was also examined in d-allulose-treated diabetic mice.

Treatment of diabetic mice with d-allulose increased SIRT1 protein levels than those observed in the vehicle-treated diabetic mice (Supplementary Fig. S5a). Since SIRT1 is a protein deacetylase with targets including PGC-1α, a primary stimulator of mitochondrial biogenesis (40), we further examined the expression of PGC-1α. Our results showed that PGC-1α acetylation was significantly increased under diabetic conditions, whereas treatment with d-allulose resulted in decreased acetylation (Supplementary Fig. S5b), indicating that d-allulose is linked to the SIRT1/PGC1-α pathway.

Next, the expression of the downstream target genes of PGC-1α, such as nuclear respiratory factor (NRF)-1, NRF-2, and mitochondrial transcription factor A (TFAM) genes, was examined (17, 47). The protein expression of the mitochondrial function-enhancing proteins NRF-1, NRF-2, and TFAM was significantly reduced in the skeletal muscles of vehicle-treated db/db mice than those observed in the skeletal muscles of db/+ mice; the expression was recovered on treatment with d-allulose (Supplementary Fig. S5c). In addition, hepatic IRE1α sulfonation, SIRT1 expression, PGC-1α deacetylation, and expression of downstream transcription factors such as NRF-1, NRF-2, and TFAM were recovered by d-allulose treatment (Supplementary Fig. S6a–d).

Further, d-allulose improved insulin signaling in the liver of db/db mice, as indicated by increased tyrosine phosphorylation of IRS-1 and IRS-2, Akt phosphorylation, and the expression of downstream GLUT4 (Supplementary Fig. S6e–g). These observations suggest that d-allulose regulates the IRE1α sulfonation-RIDD-Sirt1 decay axis and associated mitochondrial function under hyperglycemic conditions. Recent evidence has shown that a shift in the muscle fiber type from oxidative type I to the more glycolytic type II damages the muscles of obese subjects with impaired glucose tolerance (49). We investigated the effect of d-allulose on the change in skeletal muscle fiber types. However, the proportions of type I and type II fibers did not differ between those observed in the d-allulose-treated mice and vehicle-treated db/db mice (Supplementary Fig. S7a–c).

d-Allulose controls Nox4-IRE1α-RIDD-Sirt1 decay, ameliorating the diabetic state under conditions involving administration of HFD

We next examined another metabolic stress mouse model. The HFD-fed mice treated with d-allulose exhibited improvements in post-bolus glucose clearance compared with the vehicle-treated HFD-fed mice at all timepoints of the OGTT (Supplementary Fig. S8a, b). During ITT, d-allulose-treated mice showed a substantial reduction in blood glucose levels (Supplementary Fig. S8c, d). In addition, d-allulose treatment abrogated the glucose dysmetabolism-associated NADP⁺/NADPH ratio (Fig. 4a). Increases in NADPH oxidase activity, ROS, lipid peroxidation, and protein carbonylation were observed in HFD-fed mice, which were reduced by treatment with d-allulose (Fig. 4b and Supplementary Fig. S9a–d).

FIG. 4.

d-Allulose controls Nox4-IRE1α sulfonation-RIDD-Sirt1 decay, ameliorating the diabetic state in HFD conditions. Mice were fed an NCD or HFD with vehicle or d-allulose (1.02, 3.07, or 5.16 g/kg) once daily for 10 weeks through oral gavage. Tibialis anterior muscle lysates were analyzed for the NADP/NADPH ratio (a) and NADPH oxidase activity (b). (c) Expression of Nox4 and β-actin in tibialis anterior muscle tissues. (d) Quantitative analysis of protein expression was also performed. (e) Representative images of Nox4 staining in tibialis anterior muscle tissues. (f) Immunoblotting of p-IRE1α, IRE1α, GRP78, CHOP, and β-actin and RT-PCR of uXbp-1, sXbp1, and Gapdh mRNA expression in tibialis anterior muscle tissue. (g) Quantitative analysis of protein and RNA expression was also performed. (h) Representative images showing irreversible sulfonation of IRE1α in tibialis anterior muscle tissue. (i) Quantitative analysis of protein expression was also performed. (j) Heatmap depicting mRNA expression of the genes identified as RIDD substrates. (k) The mRNA level of Sirt1 was assessed by RT-PCR. Data are expressed as mean ± SEM, n = 8 animals/group. Scale bar: 100 μm. *p < 0.05 versus NCD, ^# p < 0.05 versus HFD+vehicle. HFD, high-fat diet; NCD, normal chow diet.

This finding indicates that d-allulose also regulates oxidative stress in the skeletal muscles under conditions involving the administration of HFD. Nox2 and 4 expression was remarkably increased in HFD-fed mice than those in normal chow diet (NCD)-fed mice, whereas d-allulose supplementation markedly reduced Nox4 expression than that observed under conditions involving administration of HFD (Fig. 4c–e and Supplementary Fig. S9e). IRE1α phosphorylation and subsequent Xbp1 splicing were inhibited by d-allulose (Fig. 4f, g). d-allulose supplementation substantially inhibited IRE1α sulfonation while enhancing under conditions involving the administration of HFD (Fig. 4h, i).

IRE1α-RIDD target genes, including Sirt1, were regulated by d-allulose (Fig. 4j, k). Next, the expression of SIRT1 protein was examined in vehicle- and d-allulose-treated HFD-fed mice. The HFD mice showed inhibition of muscular SIRT1 expression, PGC-1α deacetylation, and expression of downstream transcription factors such as NRF-1, NRF-2, and TFAM, which were recovered by d-allulose treatment (Supplementary Fig. S10a–c). Enhanced tyrosine phosphorylation of IRS-1 and IRS-2 was observed in d-allulose supplemented HFD-fed mice, suggesting that d-allulose supplementation improves insulin signaling in the skeletal muscle.

Moreover, increased Akt phosphorylation was observed in d-allulose-treated HFD-fed mice than that in the vehicle-treated HFD-fed mice. Further, d-allulose supplementation ameliorated the recovery of decreased GLUT4 expression and its translocation to the plasma membrane (Supplementary Fig. S10d–g). These data showed a consistent mechanism underlying IRE1α sulfonation-RIDD-Sirt1 decay observed in the aforementioned diabetic db/db model, further supporting the role of d-allulose in controlling glucose dysmetabolism signaling.

d-Allulose controls Nox4-IRE1α-RIDD-Sirt1 decay, enhancing glucose intake in HSkM cultured under high glucose conditions

Cells grown with a medium containing high levels of glucose mimic the diabetic condition and are linked to glucose dysmetabolism-based oxidative stress (34, 45). In this study, HSkM cultured with physiological glucose (5.5 mM) were incubated with a 30 mM glucose medium to mimic the diabetic glucose concentration. Glucose enhanced cellular ROS production and Nox4 expression in a time-dependent manner (Supplementary Fig. S11a–c). IRE1α sulfonation and the resultant RIDD were significantly increased under high glucose-culture conditions (Supplementary Fig. S11d–h).

The expression of the novel RIDD target gene, Sirt1, was also decreased under high glucose-culture conditions (Supplementary Fig. S11i, j). d-allulose recovered the NADP⁺/NADPH ratio and ROS production (Fig. 5a–e). Due to certain limitations of dihydroethidium (DHE) staining, including instability of the probes, potential interference (3) by heme enzymes (59), and incomplete coverage of H₂O₂, the N-acetyl-3, 7-dihydroxyphenoxazine (Amplex Red) assay was used to measure ROS levels, including those of H₂O₂, as it is based on horseradish peroxidase (HRP)-catalyzed oxidation, which reacts with H₂O₂.

FIG. 5.

d-Allulose controls oxidative stress and IRE1α sulfonation-RIDD-Sirt1 decay, ameliorating the diabetic state in high glucose-cultured HSkM. HSkM were treated with 2.5, 5, or 10 mM AP or AL, 10 μM GKT137831, 2 mM NAC, and 200 μg/mL catalase with or without 30 mM glucose for 7 days. (a) Cell lysates were analyzed for NADP/NADPH ratio. Representative DHE-stained images depicting ROS production (b), quantification (c), and lipid peroxidation (d) in each condition. Scale bars, 100 μm. (e) H₂O₂ concentration contents were determined with the Amplex^® Red Hydrogen Peroxide/Peroxidase Assay Kit. (f) Representative images from the PLA showing protein–protein interactions in vitro. Cells were incubated with primary antibodies (anti-mouse IRE1, 1:100; Santa Cruz, anti-rabbit cysteine sulfonate, 1:100; Abcam) in a humidity chamber overnight at 4°C, followed by hybridization with PLA probes (1 h/37°C), ligation (30 min/37°C), and amplification (2 h/37°C). (g) Quantification of the levels of IRE1α sulfonation is shown. (h) Irreversible sulfonation of IRE1α was analyzed as described in the Materials and Methods section. (i) Heatmap depicting mRNA expression of the genes identified as RIDD substrates. (j) The mRNA level of Sirt1 was assessed by RT-PCR. (k) Immunoblotting of SIRT1 and β-actin expression. (l) Immunoblotting of p-Akt, Akt, GLUT4, and Na⁺/K⁺ ATPase in cells. Data are represented as mean ± SEM. Scale bars, 100 μm. *p < 0.05 versus Con, ^# p < 0.05 versus 30 mM glucose. CAT, catalase; DHE, dihydroethidium; GKT, GKT137831; Glu, glucose; NAC, N-acetylcysteine; ROS, reactive oxygen species.

IRE1α sulfonation and the resultant expression of RIDD target genes including Blos1, Hgsnat, Col6, Scara3, and that of the Sirt1 gene were significantly inhibited by d-allulose (Fig. 5f–j). SIRT1 protein expression was also recovered in the presence of d-allulose (Fig. 5k). Akt phosphorylation and the expression of its downstream target GLUT4 in the plasma membrane were consistently increased under conditions involving treatment with d-allulose (Fig. 5l). These observations suggest that d-allulose improves systemic and peripheral insulin resistance and also recovers the glucose disposal in the skeletal muscles under diabetic conditions.

In in vitro experiments, inhibitors of Nox4 (GKT137831 and GLX351322) showed effects similar to those of d-allulose (Fig. 5 and Supplementary Figs. S12 and S13a–c). Consistently, N-acetylcysteine (Fig. 5a–d) and catalase (Fig. 5e) showed similar effects. In Nox4 small interfering RNA (siRNA)-treated cells showing decayed Nox4 expression (Supplementary Fig. S14), NADP⁺/NADPH ratio, ROS production, lipid peroxidation, IRE1α sulfonation, and the resultant RIDD were significantly inhibited (Fig. 6a–h).

FIG. 6.

Nox4 deficiency improves the diabetic state in d-allulose-treated cells in the 30 mM glucose-treated HSkM. Non-specific-siRNA and Nox4 siRNA were transiently transfected into HSkM. (a) Cell lysates were analyzed for NADP/NADPH ratio. Representative DHE-stained images depicting ROS production (b), quantification (c), and lipid peroxidation (d) in each condition. Scale bars, 100 μm. (e) Representative images from the PLA depicting protein–protein interaction in vitro. Cells incubated with primary antibodies (anti-mouse IRE1, 1:100; Santa Cruz, anti-rabbit cysteine sulfonate, 1:100; Abcam) in a humidity chamber overnight at 4°C. This was followed by hybridization of PLA probes (1 h/37°C), ligation (30 min/37°C), and amplification (2 h/37°C). (f) Quantification of the levels of sulfonation of IRE1α is shown. (g) Irreversible sulfonation of IRE1α was analyzed as described in the Materials and Methods section. (h) Heatmap depicting mRNA expression of the genes identified as RIDD substrates. (i) The mRNA level of Sirt1 was assessed by RT-PCR. (j) Expression of SIRT1 and β-actin. (k) Immunoblotting of p-Akt and Akt expression in cells. (l) Expression of GLUT4 and Na⁺/K⁺ ATPase in cells. Data are represented as mean ± SEM. Scale bar: 100 μm. *p < 0.05 versus non-specific-siRNA, ^# p < 0.05 versus 30 mM glucose-cultured cells with non-specific-siRNA. siRNA, small interfering RNA.

The decay of Sirt1 RNA leading to decreased protein expression was significantly recovered under conditions involving Nox4-knockdown (Fig. 6i, j). Akt phosphorylation and the expression of its downstream target GLUT4 in the plasma membrane were consistently increased under conditions involving treatment with Nox4 siRNA (Fig. 6k, l). Nox4 silencing showed similar effects as d-allulose, indicating that the Nox4-IRE1α-RIDD-Sirt1 decay axis contributes to glucose dysmetabolism, whereas d-allulose can control the signaling and recover muscular cell glucose disposal. In Figure 7, the schematic illustration indicates how d-allulose controls Nox4-linked IRE1α post-translational modification-induced GLUT4 disturbance and associated insulin resistance.

FIG. 7.

Schematic illustration of how d-allulose alleviates insulin resistance in skeletal muscle by Nox4-IRE1α-RIDD-Sirt1 decay.

Discussion

This study demonstrated that diabetes-induced ROS production mediated by Nox4 and upregulation of IRE1α sulfonation, a master ER stress response signal, and associated RIDD-Sirt1 degradation could be controlled by d-allulose treatment. Our findings suggest that the involvement of IRE1α sulfonation-Sirt1 degradation axis is a novel mechanism underlying the anti-diabetic effects of d-allulose. Moreover, d-allulose contributes to muscular glucose disposal under diabetic conditions in vitro and in vivo.

In this animal and cell-based study, Nox4 and associated ER stress were studied. This study demonstrated upregulation of Nox4 expression under diabetic conditions (Figs. 2 and 4). Under conditions involving glucose dysmetabolism and overnutrition, ER stress is associated with hyper-activated intra-ER electron uncoupling and electron leak with delayed ER oxidative folding process (19). Among the membrane-bound enzymes, Nox family proteins and ER-localized Nox4 (33) play a major role in the ER stress-linked ROS accumulation (46).

Phagocyte-like NAD(P)H oxidases are also a source of ROS in diabetes (1, 18). High glucose levels mediate ROS production via Nox4 in the rat kidney glomerular mesangial cells (18). In db/db and HFD-fed mice, the expression of Nox2 and Nox4 was high (Fig. 2d, e and Supplementary Figs. S3e and S9e). Nox2 expressed by plasma membrane is considered a typical ROS signal but not an ER-specific redox transducer. Nox4, specifically expressed by ER, clearly explains the ER stress-based GLUT4 disturbance, and it is highly associated with ER-localized NADPH oxidase-ROS signaling.

This phenomenon is linked to IRE1α sulfonation-RIDD-Sirt1 decay. Under physiologically stress conditions involving low ROS levels, the expression of GLUT4 is enhanced, indicating hormesis between ROS and GLUT4. Under hyperglycemic conditions, Nox4-induced ROS production and linked IRE1α sulfonation were reversed by d-allulose treatment (Figs. 3 and 4), indicating that the ER stress-associated ROS-Nox4-IRE1α axis contributes to the diabetic state.

Sirt1, a major metabolic enzyme, was shown to be one of the IRE1α sulfonation-linked RIDD targets. Among the Sirt family, Sirt1 harbors a target sequence for RIDD (Fig. 3h–j and Supplementary Fig. S4). The effect of d-allulose against Sirt1 decay may have important implications, considering the critical role of SIRT1 in diabetes (31). Similarly, recent studies revealed the negative association between the gene expression of Sirt1 in hepatic and muscular cells with insulin resistance (12).

Moreover, Sirt1 gene expression is decreased in patients with glucose intolerance (31). Hence, there is a direct relationship between Sirt1 transcription and measurements of insulin sensitivity (13). In this study, we found that Nox4 is controlled by d-allulose. The regulation of Nox4-IRE1α sulfonation leads to the inhibition of the substrate, Sirt1, which is one of the mechanisms underlying the anti-diabetic effect observed in the d-allulose-treated db/db mice and HFD-fed models. Previous reports demonstrated that SIRT1 expression is decreased in the skeletal muscles in diabetes (10).

Thus, we assessed Sirt1 levels in the skeletal muscles of db/db mice and HFD-fed mice. In response to d-allulose treatment, the diminished SIRT1 expression and deacetylation of its substrate PGC-1α were markedly recovered in the diabetic models (Supplementary Figs. S5a, b and S10a, b). Muscle cells rely on PGC-1α to stimulate mitochondrial biogenesis and oxidative phosphorylation during their glucose metabolic state. Indeed, PGC-1α suppression significantly impaired mitochondrial biogenesis and oxidative phosphorylation and induced muscular insulin resistance (35).

SIRT1 in the skeletal muscles is essential for the PGC-1α-mediated induction of the expression of fatty acid oxidation-associated genes in response to fasting and exercise (9, 15). This specific observation suggests that during energy-demanding processes such as fasting and physical activity, PGC-1α is activated by SIRT1-mediated deacetylation.

d-Allulose augmented insulin action-dependent glucose uptake accompanied by enhanced GLUT4 translocation to the cell surface (Fig. 1i, j and Supplementary Figs. S3f and S10g). In the cells of the skeletal muscles, GLUT4 is prominent glucose transporter activated in response to insulin, whereas GLUT1 is primarily responsible for glucose transport during the basal condition (58). Collectively, the SIRT1-PGC-1α-GLUT4 axis also explains the d-allulose-induced anti-diabetic mechanism. Our novel findings demonstrating direct muscular effects of d-allulose observed under diabetic conditions may provide a valuable therapeutic option for patients with diabetes.

The study dosage range seems to match the reported human equivalent dose range, “5 and 10 g” in diabetic or prediabetic patients (22, 41). With respect to the two formulations, liquid and powder, no differences concerning the anti-diabetic effects were observed, indicating that d-allulose itself contributes to controlling glucose metabolism regardless of the form of d-allulose.

In summary, the decay of Sirt1 as an RIDD target gene is suggested to be one of the mechanisms involved in the attenuation of diabetic conditions along with a well-established SIRT1-PGC-1α-linked GLUT4-associated glucose controlling mechanism. In addition, d-allulose contributes to muscular glucose homeostasis by controlling the Nox-IRE1α sulfonation-RIDD axis.

Materials and Methods

d-Allulose products

d-allulose powder (AP, purity >98%) and syrup (AL, purity >95%) were provided by Samyang Corp. (Seoul, South Korea). d-Allulose products were prepared by using Microbacterium foliorum (non-GMO; genetically modified organism).

Cell culture and transfection

HSkM (A12555; Thermo Fisher Scientific) were grown in a growth medium containing 10% (v/v) fetal bovine serum in Dulbecco's modified Eagle's medium (DMEM) supplemented with 1% penicillin/streptomycin under a standard humidified atmosphere. On reaching 85%–95% confluence, cells were stimulated with DMEM containing 2% horse serum for 2 days. HSkM were treated with 2.5, 5, or 10 mM AP or AL, 10 μM GKT137831, 2 mM N-acetylcysteine (NAC; Sigma-Aldrich, St. Louis, MO), and 10 μM GLX-351322 (Abcam, Cambridge, MA) with or without 30 mM glucose for 7 days. Transfections were performed by using Lipofectamine 3000 (L3000001; Invitrogen) according to the manufacturer's recommendations.

Animal and experimental design

Diabetic db/db mice (C57BLK-Leprdb/Leprdb) and non-diabetic db/m mice (C57BLK-Leprdb/+) were purchased from the Jackson Laboratory (Bar Harbor, ME), and they were housed in the laboratory animal care facilities of Jeonbuk National University Hospital under standard SPF conditions. After acclimatization, 7-week-old male mice were randomly divided into eight groups (n = 8/group): normal control (db/m; water), negative control (db/db; water), three test groups (db/db; AP-low, -medium, and -high), and three test groups (db/db; AL-low, -medium, and -high).

d-Allulose was administered at doses of 1.02 g/kg (low), 3.07 g/kg (middle), or 5.16 g/kg (high) to the appropriate test groups for 8 weeks. The oral doses of d-allulose were determined based on human studies (7, 22, 41). Reagents (water and d-allulose) were administered daily via oral gavage, and weekly food consumption, body weight, and post-prandial blood glucose levels were recorded. At the end of the seventh week, OGTT and an ITT were performed. At the end of the eighth week, mice were anesthetized and euthanized. Blood samples from were collected the truncal vein, tibialis anterior muscle sections, and liver as per the guidelines set by the Institutional Animal Care and Use Committee of Jeonbuk National University Hospital (JBUH-IACUC-2020-04). All animal protocols are implemented as per the Guide for the Care and Use of Laboratory Animals.

Similarly, for developing another metabolic stress animal model, 7-week-old male mice were fed with HFD or a normal diet. All mice were randomly divided into eight groups (n = 8/group): normal control (normal diet; water), negative control (HFD; water), three test groups (HFD; AP-low, -medium, and -high), and three test groups (HFD; AL-low, -medium, and -high). d-Allulose was administered at doses of 1.02 g/kg (low), 3.07 g/kg (middle), or 5.16 g/kg (high) to the appropriate test groups for 10 weeks. Reagents (water and d-allulose) were administered daily via oral gavage. At the end of the 10th week, the mice were anesthetized and euthanized.

Blood samples were collected from the truncal vein, tibialis anterior muscle sections, and liver. The tibialis anterior muscle and the liver sections were used for Western blotting, real-time polymerase chain reaction (RT-PCR), and histological analysis. All dissected tissues were immediately frozen in liquid nitrogen and stored at −80°C until further analysis.

Glucose and ITT

For OGTT, mice fasted overnight were administered glucose at 1 g/kg via oral gavage. For ITT, mice fasted for 6 h were intraperitoneally injected with insulin (0.75 IU/kg BW: Actrapid Penfill Insulin human; Novo Nordisk, Bagsværd, Denmark). After administration of glucose/insulin, blood samples (∼30 μL) were derived from the tail vein at 0, 15, 30, 45, 60, 90, and 120 min, and blood glucose levels were determined by using a fresh capillary tube (Accu-Check glucometer; Roche Diagnostics). For checking glucose tolerance and sensitivity, we measured the total area under the curve (AUC) to show glucose tolerance test (GTT) and insulin tolerance test (ITT).

Biochemical analysis

The collected serum was used to measure the biochemical parameters. Insulin and HbA1c levels were measured with commercially available assay kits (Insulin: K4271; HbA1c: E4657; Biovision, San Francisco, CA). All protocols were performed according to the manufacturer's guidelines.

Histological analysis and immunohistochemistry

The tibialis anterior muscle was excised and frozen in liquid nitrogen-cooled isopentane (Nacalai Tesque). Transverse sections (5 μm) were cut by using cryostats (CM3050S; Leica Microsystems, Wetzlar, Germany) and collected onto MAS-coated glass slides (Matsunami). Hematoxylin and eosin (H&E) staining was carried out by using these sections. Immunohistochemistry (IHC) was performed as previously described (32). Briefly, hydrogen peroxide-fixed islets were embedded and cut into sections with a thickness of 4 μm.

To prevent nonspecific binding, sections were incubated with a blocking reagent (UltraTech HRP; Beckman Coulter, Brea, CA). The sections were subsequently incubated with antibodies against Nox4, Troponin T type I (Novus Biologicals, Littleton, CO), and MY-32 (Abcam). HRP-conjugated biotin-streptavidin complexes were used to assess antibody reactivity, and staining color was developed by substrate diaminobenzidine tetrahydrochloride.

Immunofluorescence staining

The immunofluorescence staining protocol was optimized for the analysis of GLUT4 (6). Briefly, the tibialis anterior muscle sections were fixed and washed with phosphate-buffered saline (PBS). The sections were exposed to a primary antibody against GLUT4 (MA5-17176; dilution: 1:200; Invitrogen) in combination with a primary antibody against dystrophin (PAS-32388; dilution: 1:400; Invitrogen) for 2 h at room temperature. After incubation with a primary antibody, the sections were washed thrice for 5 min in PBS and were then incubated in 1:200 dilution of secondary antibodies for 30 min at room temperature.

The sections were washed thrice for 5 min in PBS, and glass coverslips were mounted with 20 μL aqueous-mount solution (Scytek Laboratories). Images were captured by using an inverted confocal microscope (Leica DMIRE2; Leica Microsystems) with a 63× oil immersion objective lens. AlexaFluor 488-conjugated goat anti-rabbit IgG secondary antibody (F0382; Sigma) was used to detect the GLUT4 primary antibody, whereas AlexaFluor 594-conjugated goat anti-mouse IgG2b (T5393; Sigma) was used to detect the dystrophin primary antibody. Images were captured by using the Zeiss LSM 880 inverted confocal microscope (Carl Zeiss, Inc.). All the images were captured with the same laser intensities. For quantitating, colocalization of GLUT4 and dystrophin was calculated from each image by using ImageJ.

Immunoblotting and IP

Immunoblotting was performed as previously described, with minor modifications (32). Briefly, 20–40 μg of protein samples (tibialis anterior muscle, liver, and cell lysates) was separated on a polyacrylamide gel and transferred onto PVDF membranes. Later, blocked membranes were incubated with primary antibodies against p-IRE1α (No. ab124945; Abcam), IRE1α (No. 3294; Cell Signaling), p-eIF2α (No. 3597; Cell Signaling), eIF2α (No. 9722; Cell Signaling), ATF4 (No. 11815; Cell Signaling), CHOP (No. 2895; Cell Signaling), GRP78 (No. 13539; Santa Cruz), β-actin (No. sc-47778; Santa Cruz), Nox4 (No. PA5-72816; Invitrogen), Nox1 (No. PA5-38031; Invitrogen), Nox2 (No. ab129068; Abcam), and Nox3 (No. ab81864; Abcam). The blots were then washed, and corresponding species-specific secondary antibodies were added.

The signals were detected with enhanced chemiluminescence reagents. For IP, 100 μg of protein samples was incubated with corresponding antibodies overnight at 4°C. To pull the antibody/antigen complex, samples were incubated with protein A/G Sepharose beads. Then, the immunoprecipitates were washed with PBS-T and analyzed by immunoblotting with corresponding antibodies.

DHE staining

DHE was used to evaluate intracellular ROS production in the tibialis anterior muscle and HSkM cells. Fixed tibialis anterior muscles and cells were exposed to DHE (ThermoFisher Scientific, MA) at a concentration of 10 and 20 μM, respectively, for 30 min/37°C, followed by washing with PBS. The images were captured by using confocal microscopy. Relative fluorescence intensity was analyzed with ImageJ (National Institutes of Health, Bethesda, MD).

ROS measurement

Quantitative measurement of H₂O₂ levels was performed by using the Amplex^® Red hydrogen peroxide/peroxidase assay kit (A22188; Molecular Probes, Thermo Fisher Scientific, CA) according to the manufacturer's instructions. Briefly, 50 μL of the lysate was incubated with 50 μL of a solution containing 0.05 M NaH₂PO₄ (pH 7.4), 0.2 U/mL of HRP, and 25.7 mg/mL of the Amplex Red (10-acetyl-3,7-dihydroxyphenoxazine) reagent for 2 h at 37°C.

After incubation, the absorbance of the reaction mixture was measured at 560 nm. Absorbance readings were compared with those obtained to develop a standard curve for H₂O₂ (0–40 μM). Results are expressed as H₂O₂ concentrations in μM.

Oxyblot assay

The levels of carbonylated protein in the tibialis anterior muscle were measured via OxyBlot Protein Detection Kit (Millipore, Billerica, MA) according to guidelines provided by the manufacturer. Briefly, the skeletal muscle lysate samples were incubated in a 1:1 ratio in 2,4-dinitrophenylhydrazine (DNPH), followed by the addition of 2-mercaptoethanol for protein denaturing. Later, a neutralization reagent was added to stop the reaction, and samples were loaded for immunoblotting as described in previous sections.

IRE1α sulfonation

Sulfonation was performed as previously described (30). The total tibialis anterior muscle and cell lysates (∼500 μg) were prepared by using lysis buffer (Cell Signaling) comprising protease and phosphatase inhibitor cocktail (Sigma). To detect sulfonation of IRE1α, IP was performed by using anti-cysteine-sulfonate (ab176487; Abcam) derived from lysates, followed by application of the anti-IRE1α antibody (3294; Cell Signaling). Protein A/G Sepharose beads (Sigma) were added and incubated for an additional 1 h.

Immunoprecipitates were washed five times with PBS-T buffer or PBS before being resolved by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) and were immunoblotted with the indicated antibodies.

Proximity ligation assay

Duolink in situ reagents (Sigma-Aldrich) were used to perform the PLA. The protocol was performed according to the manufacturer's instructions. Briefly, the tibialis anterior muscle sections (5 μm) and the cells were fixed with 4% paraformaldehyde (pH 7.4) in PBS for 10 min at room temperature. Permeabilization was performed by using 0.1% Triton X-100 in PBS for 10 min at room temperature. Blocking was realized with 0.1% Triton X-100 in PBS with 5% bovine serum albumin (BSA) for 1 h at room temperature. Incubations with primary antibodies (cysteine sulfonate and IRE1α, 1:100 in 0.1% Triton X-100 in PBS with 5% BSA) were performed overnight at 4°C in a humidity chamber.

Then, the cells were rinsed with Wash Buffer A. Next, PLUS and MINUS secondary PLA probes comprising both rabbit and mouse immunoglobulins diluted in 0.1% Triton X-100 in PBS with 5% BSA were added for 1 h at 37°C. The incubation was followed by washing for 5 min with two changes of Wash Buffer A. Subsequently, the slides were incubated with the Duolink ligation mix for 30 min at 37°C and thereafter washed with two changes of Wash Buffer A for 5 min each.

The Duolink amplification mix was then applied to the slides for 100 min at 37°C. Subsequently, the slides were washed twice for 10 min each time with Wash Buffer B. The nuclei were stained with DAPI and mounted on microscope slides, sealed with an anti-fading solution, and examined under a Widefield microscope Zeiss Axiovert 200M. HSkM were treated with H₂O₂ for 6 h and used as a positive control. Negative control was developed without incubation with primary antibodies. Quantification of the PLA red fluorescent dots was performed by using ImageJ.

NADPH-dependent oxidoreductase activity assay

NADPH oxidase activity was evaluated by analyzing superoxide production via lucigenin-enhanced chemiluminescence (28). Tibialis anterior muscle tissues or cells were washed with ice-cold PBS and homogenized in cold lysis buffer (20 mM sodium phosphate buffer [pH 7.0], 1 mM EDTA, 1 mM PMSF, 0.5 mM leupeptin, and 0.5 mM pepstatin) by sonication on ice. The homogenates (5 μL) were added to 0.5 mL of assay buffer (50 mM sodium phosphate buffer [pH 7.0], 1 mM EDTA, 150 mM sucrose, and 50 μM lucigenin).

After the addition of 0.1 mM NADPH, luminescence was measured as relative light units (RLUs) for 20 s at 1 min intervals in a Skanlt RE 6.1 (Thermo Fisher Scientific, Waltham, MA). The results were expressed as RLUs/mg protein and compared between groups.

Quantitative RT-PCR

The tibialis anterior muscle tissue was homogenized with the POLYTRON^® homogenizer Kinematica Polytron^™ PT2100) in 700 μL of QIAzol^® Lysis Reagent. Total RNA was purified from the homogenate by using the miRNeasy Mini Kit (Qiagen) according to the manufacturer's instructions. Reverse transcription was performed by using 1 μg of RNA following the instructions of the iScript^™ complementary DNA (cDNA) synthesis kit (Bio-Rad).

Experimental transcript levels were analyzed by using the iQTM SYBR Green Supermix (Bio-Rad) on StepOnePlus (Applied Biosystems). All quantitative RT-PCR data were normalized to β-actin expression analyzed in separate reactions.

In vitro transcription and cleavage assays

The synthetic Sirt1 gene sequence was inserted into pMA-RQ (GeneArt). To produce 5′ mRNA fragments, pMA-RQ Sirt1 plasmid DNA was linearized by BglII (Promega). In vitro transcription of the 5′ region of Sirt1 was performed with a T7 transcription kit (AM1333; Thermo Fisher Scientific). In vitro cleavage assay of Sirt1 mRNA was performed following a protocol previously described (27). The RNA products obtained by in vitro transcription were incubated with or without recombinant IRE1 (E31-11G; SignalChem) at 37°C. The RNA fragments were separated on a 1.5% denaturing agarose gel.

Statistical analysis

All immunoblots for in vitro studies were developed in triplicate. Data analysis was performed by using GraphPad Prism 5.02 (San Diego, CA). One-way analysis of variance performed by using Tukey post hoc comparison was used to compare groups. Basic data were presented as mean ± standard error of the mean. Statistical significance was set at p < 0.05. An electronic laboratory notebook was not used.

Footnotes

Authors' Contributions

H.-Y.L. and G.-H.L. planned the experiments, conducted the studies, analyzed the data, and were primarily responsible for writing the article. T.H.H., S.-A.P., J.W.L., and J.H.L. conducted the studies and analyzed the data. S.S., G.E.K., and J.S.H. reviewed and edited the article. J.K. and H.-J.C. is the guarantor of this work and, as such, had full access to all the data in the study and take responsibility for the integrity of the data and the accuracy of the data analysis.

Author Disclosure Statement

The authors have read the journal's policy and declare conflict of interests. There are no patents, products in development, or marketing products to declare.

Funding Information

This work was supported by Samyang Corp.

Supplementary Material

Supplementary Figure S1

Supplementary Figure S2

Supplementary Figure S3

Supplementary Figure S4

Supplementary Figure S5

Supplementary Figure S6

Supplementary Figure S7

Supplementary Figure S8

Supplementary Figure S9

Supplementary Figure S10

Supplementary Figure S11

Supplementary Figure S12

Supplementary Figure S13

Supplementary Figure S14

Abbreviations Used

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