Schwann cells (SCs) are the main glial cells of the peripheral nerve and are responsible for the development, function, and regeneration of peripheral nerves by ensheathment of unmyelinated axons, myelination of myelinated axons and secretion of neurotrophic factor [1]. In the process of peripheral nerve injury, SCs proliferate and phagocytize the disintegrated axons and myelin and then migrate to guide the regeneration direction of the newborn axon [2]. However, SCs are very sensitive to glucose and insulin levels and closely involved in the occurrence and development of type 1 diabetes mellitus, as well as diabetic peripheral neuropathy (DPN) which is a common diabetes’ complication [3]. The accumulating data demonstrated that the damage of SCs in diabetes mellitus caused neurotoxic intermediates accumulation and neurotrophic factors deficiency, contributing in axonal degeneration, finally leading to DPN [4,5]. However, the exact underlying mechanism regarding the involvement of SCs in the DPN pathogenesis is still not clear.

Autophagy is the process of continuously breaking down modified proteins or damaged organelles in lysosomes to maintain cell homeostasis and function [6]. It is characterized by the sequestering of organelles/aging or damaged proteins and the formation of autophagosomes, protecting cells from a variety of factors including oxidative stress and daily wear and tear [7]. Reactive oxygen species (ROS)-induced activation of PARP-1 promotes autophagy by activating AMP-activated protein kinase, likely via inhibition of mTOR [8]. Modulation of autophagy under conditions of oxidative stress has been implicated in pathologies such as cancer, myopathy, neurodegenerative diseases, heart disease, liver disease, gastrointestinal disorders, and complications of diabetes mellitus [9-14]. The expression of autophagy markers in high-glucose-treated SCs decreased with increasing glucose concentration. This indicates that the level of autophagy in SCs decreases in a high-glucose environment and promotes apoptosis [15]. Autophagy in neural tissue eliminates damage caused by cellular stressors. It has been implicated in the pathophysiology of DPN, is related to oxidative stress and a variety of molecular mechanisms, and its dysfunction triggers apoptosis [16]. Therefore, it is important to explore the autophagy signal transduction pathways linked to high glucose-induced damage to SCs.

The ShcA family includes p46shc, p52shc, and p66shc; of these, p66shc are redox enzymes that regulates the level of intracellular ROS under cellular stress [17]. In diabetes, the increase in p66Shc levels have been reported in the peripheral blood mononuclear cells [18]. Moreover, numerous studies have shown that diabetic p66Shc-/- mice are protected from the onset of both micro- and macro-vascular complications compared to their wild-type counterpart [19-22]. It has been shown that p66Shc gene deletion increases the expression of antioxidant enzymes in endothelial cells, alleviates hyperglycemia-induced oxidative stress, and prevents diabetes-induced endothelial dysfunction in streptozotocin (STZ)-treated mice [22]. Besides, a hyperglycemia-induced increase in p66Shc expression was also found in human retinal endothelial cells and retinal pigment epithelial cells. Increased p66Shc expression resulted in functional damage and/or apoptosis in these cells, thus contributing to the development of diabetic retinopathy [23,24]. Interestingly, p66Shc has been reported to interact with LC3-II through the binding between a distinct N-terminal CH2 domain and three putative LC3 interacting region motifs, which is associated with p66shc-dependent autophagy [25]. The ability of p66shc to promote intracellular accumulation of ROS likely triggers metabolic imbalance and thus also autophagy.

We hypothesized that p66shc mediates high glucose-induced ROS generation and autophagy dysfunction. Therefore, we evaluated the role of SCs and related mechanisms in diabetic neuropathy.

Reagents and antibodies

Anti-Beclin1 antibody was purchased from Abgent. Anti-LC3-II and -LC3-II antibodies were from Novus Biologicals. Anti-p66shc was obtained from BD Biosciences. Anti-GRP78, anti-phospho-eIF2α, anti-total-eIF2α, and anti-CHOP were from Cell Signaling. Anti-p62 and the loading control anti-β actin were purchased from Sigma-Aldrich.

Cell culture and transfection

Rat SCs (S16) were obtained from the American Type Culture Collection and cultured in Dulbecco’s modified Eagle medium supplemented with 4 mM L-glutamine, 25 mM glucose, 1 mM sodium pyruvate, 1,500 mg/l sodium bicarbonate, 10% fetal bovine serum, 10 U/ml penicillin, and 10 µg/ml streptomycin (Lonza). The cells were incubated at 37°C in a 5% CO₂ atmosphere. In duplicate, cultured cells were treated with 25, 50, 100, and 150 mM glucose for 72 h. S16 rat SCs were transfected with short interfering RNA (siRNA) for 72 h (rat p66shc siRNA: sense-UGAGUCUCUGUCAUCGCUGUU and antisense-CAGCGAUGACAGAGACUCAUU [Bioneer]) and negative control siRNA using the Lipofectamine 2000 reagent (Invitrogen) per the manufacturer’s recommendations.

Diabetic mice model

The Institutional Animal Care and Use Committee of Chungnam National University (CNUH-019-A0058) approved the animal experiments. Six-week-old male C57BL/6 mice (Samtako, Inc.) were divided randomly into control and experimental groups. Diabetes was induced with STZ (2-deoxy-2-3-[methyl-3-nitrosoureido]-D-glucopyranose; Sigma) dissolved in 0.1 M citrate buffer (pH 4.5) and intraperitoneally injected at a dose of 50 mg/kg for 5 consecutive days. The control group was injected with citrate buffer vehicle. A glucometer (LifeScan) was used to measure blood glucose levels in tail venous blood every week. Following injection, the mice were monitored for symptoms of diabetes and body weight loss. The mice were weighed before STZ injection, and weekly thereafter, to monitor changes. All in vivo experiments (von Frey testing and immunofluorescence microscopy) were performed by two different individuals. Development of sensory neuropathy was determined by von Frey filaments and the up-down method. This was performed before STZ injection, and 4 and 8 weeks thereafter. Mice were placed individually in a plastic cage with a wire-mesh bottom and allowed to acclimatize to this environment for 30 min before testing. The mechanical threshold was determined by applying a series of von Frey filaments (0.6, 1, 1.4, 2, 4, 6, 8, 10, and 15 g). The filaments were pressed perpendicular to the median plantar surface of the right hind paw of each mouse. Prolonged hind paw withdrawal and licking or biting of the hind paw were considered positive responses. Each filament was tested ten times per paw. Control and experimental mice were sacrificed 8 weeks after STZ injection for histology and Western blot analysis.

Measurement of ROS levels

S16 rat SCs were stained with 5 µM 2,7-dichlorodihydrofluo-rescein diacetate (H₂-DCFDA) (Invitrogen) for 30 min at 37°C. Equal numbers of cells (10 × 10⁴) were collected and transferred to a black 96-well plate. Fluorescence was analyzed using a fluorometer (Thermo Scientific) with a 485 nm excitation/530 nm emission filter.

Fluorescence-activated cell sorting (FACS) analysis of apoptosis and ROS

The apoptosis of SCs was assessed by flow cytometry with Annexin V-FITC/propidium iodide (PI) staining according to the manufacturer’s instructions. S16 rat SCs were treated with the siRNA p66shc and glucose for 72 h, suspension and adherent cells were collected, washed twice with cold phosphate-buffered saline (PBS), and stained with Annexin V at room temperature for 15 min and PI at room temperature (apoptosis) and 5 µM H₂-DCFDA for 30 min at 37°C (cellular ROS). The cells were analyzed by flow cytometry (BD Biosciences FACS). Annexin V-positive and PI-negative cells were considered early apoptotic and Annexin V-positive and PI-positive cells late apoptotic or dead. For ROS analysis, histogram shift was confirmed by measuring the glucose concentration or the flow cytometry value of S16 rat SCs treated with the siRNA p66shc.

Immunofluorescence microscopy

For immunofluorescence staining, sciatic nerves were thoroughly washed with 0.01 M PBS, placed in sodium citrate, boiled, and cooled for 5 min. After washing three times with PBS, sections were placed in PBS with 10% goat serum and 0.1% Triton X-100 for 30 min. Samples were incubated overnight at 4°C with primary antibodies against mouse LC3 (1: 1,000), and p62 (1:500). After washing with PBS, the samples were incubated with Cy3 anti-rabbit or anti-mouse secondary antibody (1: 500) for 2 h at room temperature. Nuclei were stained with 4,6-diamidino-2-phenylindole (DAPI). Sections were examined using a digital fluorescence microscope (DMI 6000; Leica Microsystems) equipped with a DAPI filter (365 nm). LC3, and p62 were visualized using a PE filter (546 nm) at 400× magnification, and images were captured with a photometric camera (Cool Snap HQ; Photometrics). S16 rat SCs were cultured on glass coverslips in 12-well plates in medium supplemented with 25 or 150 mM glucose for 72 h. After washing with PBS, cells were fixed with 4% (w/v) paraformaldehyde and permeabilized with 0.1% (v/v) Triton X-100. After blocking for 30 min with PBS containing 5% (w/v) bovine serum albumin and 5% (v/v) horse serum, cells were incubated overnight at 4°C with antibodies against LC3 (1:200), and p62 (1:200). The cells were labeled with a Alexa Fluor 488-conjugated secondary antibody (1:1,000) or Fluor 647-conjugated secondary antibody (1:1,000) for 1 h in the dark at room temperature. Images were obtained using a fluorescence microscope.

Western blot analysis

Sciatic nerve sections were homogenized in lysis buffer (20 mM Tris-HCl, pH 7.5, 150 mM NaCl, 1 mM ethylene glycol tetra acetic acid, 1% NP-40, 1% sodium deoxycholate, 2.5 mM sodium pyrophosphate, 1 mM sodium orthovanadate, 1 mM β-glycerophosphate, and a protease inhibitor cocktail) using a homogenizer (PRO200; PRO Scientific). Cells were harvested and the supernatants were collected. These were separated by sodium dodecyl sulfate polyacrylamide gel electrophoresis and transferred to a nitrocellulose membrane. After blocking with 5% non-fat dry milk in Tris-buffered saline containing 0.05% Tween 20, the membrane was incubated with the primary antibodies overnight at 4°C. The membrane was treated with an appropriate peroxidase-conjugated secondary antibody, and the chemiluminescent signal was developed using the SuperSignal West Pico or Femto substrate (Pierce Biotechnology). Band densities were quantified using the Gel Doc 2000 imaging system and Quantity One software (Bio-Rad). The membrane was reacted with an anti-β-actin antibody to normalize loading.

Mouse genotyping

Chromosomal DNA was isolated from mouse tails (2–3 mm length). Polymerase chain reaction (PCR) was conducted using a KOD-FX (TOYOBO) genotyping kit according to the manufacturer’s instructions. The primer sequences were as follows: reverse (5'-TAA TAC GAC TCA CTA TAG GG-3') and forward (5'-AGG CGA TTA GTT GGG TA-3') for p66shc-RNAi KO. PCR conditions were 95°C for 5 min (one cycle), 95°C for 30 sec, 56°C for 45 sec, 72°C for 45 sec (33 cycles), and 72°C for 10 min.

Statistical analysis

Statistical analysis was performed using SPSS software (ver. 12.0 for Windows; SPSS Inc.). Student’s t-test and analysis of variance (ANOVA) with Tukey’s post-hoc test were used for between-group comparisons. A p-value less than 0.05 was considered indicative of statistical significance. Data are means ± standard error of the mean (SEM) of three independent experiments.

p66shc deficiency attenuates oxidative stress and early apoptosis triggered by high glucose condition in SCs

As the importance of apoptosis in the induction of diabetes and the pathophysiology of different diabetes related complications have been reported in several papers [26-28], we evaluated apoptotic cell death of SCs in high glucose condition by performing Annexin V assay. As shown in Fig. 1A, FACS showed that high glucose (150 mM) in SCs significantly induced early apoptosis compared to the control (25 mM).

Figure 1. p66shc-deficient Schwann cells attenuated oxidative stress, and early apoptosis triggered by high glucose concentrations. S16 rat Schwann cells were treated with glucose for 72 h. (A) Percentages of apoptotic S16 rat Schwann cells by flow cytometry. (B) p66shc protein expression was measured by Western blotting. (C) Schwann cells transfected with siCON and sip66shc and treated with glucose (25 or 150 mM). ROS was measured by DCF-DA fluorescence and flow cytometry. (D) Percentages of apoptotic S16 rat Schwann cells by flow cytometry. β-actin was used as the loading control. Data represent means ± SEM, n = 3. ROS, reactive oxygen species; PI, propidium iodide; DCF-DA, 2,7- dichlorofluorescein. *p < 0.05, **p < 0.01, ***p < 0.001, and ****p < 0.0001, siCON + glucose 150 mM compared with siCON + glucose 25 mM; ^#p < 0.05, sip66shc + glucose 150 mM compared with siCON + glucose 150 mM.

p66shc was reported as a pro-apoptotic protein that regulates oxidative stress and induces apoptosis by means of redox activity [29]. Since we observed high glucose-induced apoptosis in SCs, we investigated protein expression of p66shc. Our data showed a dose-dependent increase in protein expression of p66shc in SCs treated with high glucose (Fig. 1B). To further clarify whether the elevated p66shc level produced intracellular ROS causing apoptosis in SCs in high glucose conditions, we first inhibited p66shc protein expression with sip66shc. S16 cells were transfected with sip66shc, following with high glucose treatment for 72 h. p66shc deficiency was confirmed after siRNA transfection (Supplementary Fig. 1A). As expected, both accumulation of ROS and early apoptosis triggered by high glucose concentration was significantly abrogated by knockdown of p66shc (Fig. 1C, D). These results suggest that high glucose condition induces apoptosis via p66shc regulation.

Downregulation of p66shc enhances the expression of autophagy-related proteins in high glucose condition

High glucose inhibits autophagy in chondrocytes and induces autophagy via oxidative stress [30]. We previously demonstrated that increased levels of ROS induced by high glucose may contribute to autophagy dysfunction in SCs [31]. Since p66shc knockdown rescued high glucose-induced oxidative stress, autophagy may also be regulated by p66shc deficiency. To investigate this hypothesis, we examined the expression of autophagy-related proteins in p66shc-deficient S16 cells treated with high glucose. As shown in Fig. 2A, both the expression of p62 and the conversion of LC3 I to LC3 II were reduced by high glucose, which was restored similarly to the control by deletion of p66shc. Immunofluorescence staining confirmed the protein level data (Fig. 2B, C). Furthermore, in order to ensure precise autophagy assessment, lysosomal inhibitor Bafilomycin A1 was employed. Interestingly, with the presence of Bafilomycin A1, while high glucose concentration (150 mM) still decreased the amount of LC3II and p62 compared to control condition (25 mM), the deletion of p66shc partly restored the protein level of LC3II and p62 (Supplementary Fig. 1B).

Figure 2. Downregulation of p66shc enhanced the expression of autophagy-related protein in high glucose condition. Schwann cells transfected with siCON and sip66shc and treated with glucose (25 or 150 mM). (A) Expression of autophagy-related proteins (p62 and LC3-II) and p66shc by Western blotting in S16 rat Schwann cells. (B) Immunofluorescence images showing expression of p62 (red) or LC3 (red), and p66shc (green). (C) p62 and LC3 intensity was quantified per cell. Nuclei were stained with DAPI. Scale bar, 20 μm. β-actin was used as the loading control. Data represent means ± SEM, n = 3. DAPI, 4,6-diamidino-2-phenylindole. **p < 0.01 and ***p < 0.001, siCON + glucose 150 mM compared with siCON + glucose 25 mM; ^#p < 0.05, ^##p < 0.01, and ^###p < 0.001, sip66shc + glucose 150 mM compared with siCON + glucose 150 mM.

p62 deletion impairs autophagosomes, exacerbating cell injury and misfolded protein stress in cardiomyocytes [32]. It was hypothesized that endoplasmic reticulum (ER) stress is activated in SCs treated with high glucose. We evaluated ER stress-related proteins. There was an overexpression of GRP78, p-eIF2α, CHOP in SC-treated high glucose (Supplementary Fig. 2A), and the p66shc deletion alleviated high glucose induced-ER stress (Supplementary Fig. 2B). These data collectively demonstrate that downregulation of p66shc recovers the high glucose-induced impairment of autophagy and ER stress in SCs.

p66shc deficiency alleviates diabetes-induced impairment of autophagy in sciatic nerve

p66shc expression was upregulated in SCs in high glucose condition. To confirm whether p66shc also showed increased expression in diabetic mice model, STZ-induced diabetic mice were employed. Six-week-old BL6 mice were consecutively injected with vehicle or 50 mg/kg/day STZ intraperitoneally for 5 days, and diabetes was observed for 8 weeks after onset. Diabetes in the mice was confirmed by monitoring their blood level and body weight before and after STZ injection. Two weeks after injection, the concentration of blood glucose in the diabetes group was higher than that of the control group; non-fasting blood glucose concentration of 250 mg/dl or more was defined as diabetes (Supplementary Fig. 3A). As the disease progressed, the body weight of STZ-treated mice did not change while that of normal mice were increased (Supplementary Fig. 3B). Our data confirmed that p66shc protein expression was significantly increased in the sciatic nerve of diabetic mice compared with control mice in both Western blot (Fig. 3A), and immunofluorescence data (Fig. 3B). DPN is often accompanied with by loss of sensation [33] and deficiency of p66Shc protects against cellular dysfunction caused by high glucose condition. To investigate the potential role of p66shc in DPN, von Frey test to monitor mice pain sensitivity was performed before, 4 weeks, and 8 weeks after STZ injection in p66shc knockout (KO) and wild-type (WT) mice. We proved that there is no significant difference in pain sensitivity between p66shc KO and WT mice. As expected, diabetic WT mice showed a markedly increased threshold upon exposure to von Frey filaments compared with normal WT mice, and diabetic p66shc KO mice showed a significant decrease in threshold compared with diabetic WT mice (Supplementary Fig. 3C).

Figure 3. Expression of p66shc in the sciatic nerve from STZ-induced diabetes mice. (A, B) Expression of p66shc by Western blotting and immunofluorescence staining, respectively in sciatic nerves from CON and T1DM mice. β-actin was used as the loading control. Nuclei were stained with DAPI. Scale bar, 20 μm. Data represent means ± SEM, n = 15. STZ, streptozotocin; CON, control (saline); T1DM, type 1 diabetes; DAPI, 4,6-diamidino-2-phenylindole. **p < 0.01 and ***p < 0.001.

Since downregulation of p66shc recovers the high glucose-induced impairment of autophagy and ER stress in SCs, p66shc may ameliorate diabetes-induced mechanical hypoalgesia via autophagy recovery. To assess this hypothesis, we compared expression of autophagy and ER stress-associated proteins between WT and p66shc KO mice injected vehicle or STZ. There was no difference in blood glucose level between p66shc KO mice and WT mice, and diabetes was induced in p66shc KO mice based on non-fasting blood glucose concentration. However, diabetic p66shc KO mice had lower blood glucose level than the diabetic WT mice (Supplementary Fig. 3A). Our Western blot data figure out that decrease in p62 expression and LC3-II in sciatic nerve from diabetic WT mice were significantly restored in diabetic p66shc KO mice (Fig. 4A). Similarly, immunofluorescent analyses suggested an effect of p66shc downregulation on p62 expression and the conversion of LC3 I to LC3 II, respectively (Fig. 4B, C). ER stress experienced same results, with the activation of ER stress in diabetic mice (Supplementary Fig. 4A) and diabetic p66shc KO mice partly suppressed ER stress in sciatic nerve (Supplementary Fig. 4B). In summary, p66shc ablation alleviates the impairment of autophagy and ER stress in the sciatic nerve of STZ-induced diabetes mice.

Figure 4. Effect of p66shc KO on autophagy in the sciatic nerve of STZ-induced diabetes mice. (A) Expression of autophagy-related proteins (p62, and LC3-II) by Western blotting in sciatic nerves from CON, T1DM, p66shc KO, and p66shc KO + T1DM mice. β-actin was used as the loading control. (B) Immunofluorescence of the expression of the autophagy-related proteins p62 (red), or LC3 (red), and p66shc (green) in sciatic nerves from CON, T1DM, p66shc KO, and p66shc KO + T1DM mice. (C) The fluorescent area of p62 and LC3 was quantified. Nuclei were stained with DAPI. Scale bar, 20 μm. Data represent means ± SEM, n = 15. CON, control (saline); T1DM, type 1 diabetes mice; p66shc KO, p66shc knockout mice; p66shc KO + T1DM, p66shc KO type 1 diabetic mice; DAPI, 4,6-diamidino-2-phenylindole. *p < 0.05 and ***p < 0.001, T1DM compared with CON; ^#p < 0.05, ^##p < 0.01, and ^###p < 0.001, p66shc KO + T1DM compared with T1DM.

SCs are the fundamental component of peripheral nervous system as well as the neuronal regulatory network, specific in the process of peripheral nerve injury in diabetes. Excessive oxidative stress in DPN is considered a common pathway for hyperglycemia-induced SC dysfunction [34]. Oxidative stress in DPN patients is higher compared to healthy individuals and increases proportionally with disease severity. p66shc regulates oxidative stress and is implicated in diabetic diseases [35]. However, the function of p66shc in SC dysfunction is still unclear. In this study, our results revealed a vital role of p66shc in SCs in diabetes. While the impairment of autophagy was observed, p66shc was upregulated in diabetic mice’s sciatic nerve. In addition, the deletion of p66shc restored autophagy function in diabetic mice.

Sera from DPN patients induces the accumulation of autophagosomes by increasing LC3-II immune reactivity when exposed to neuroblastoma cells [36]. Autophagy was impaired in cerebellar Purkinje neurons in mice induced by STZ-induced DPN [37]. Therefore, regulation of autophagy by hyperglycemia in DPN is important for neuronal function and health. In this study, we investigated the effect of DPN-induced hyperglycemia on autophagy of SCs. In SCs, expression of p62 and LC3-II was decreased, while the presence of lysosomal inhibitor could not raise the protein level of both p62 and LC3-II. Generally, by using lysosomal inhibitors such as bafilomycin A1, the autophagic flux could be demonstrated by the raise in LC3II and p62 level in Western blot. p62 participates in the conversion of LC3-I to LC3-II and induces autophagosome maturation and autophagosome entry of damaged organelles [32]. Moreover, p62 deletion results in the impaired formation of the LC3-II, aggresome, and autophagosome, lowering cell viability under basal conditions and misfolded protein stress in cardiomyocytes [32]. LC3-I is activated by Atg7 and modified by Atg3 to bind the membrane and generate LC3-II. The LC3-II level is proportional to the number of autophagic vacuoles, and expression of LC3-II is a direct indicator of autophagy induction [38]. From our results, we suggest the downregulation of p62 caused by hyperglycemia condition was the result behind the decrease in LC3-II due to the impairment of LC3-II formation. Therefore, autophagy process was inhibited in hyperglycemia condition. Interestingly, hyperglycemia induced decrease in p62 expression and LC3-II/LC3-I ratio were reversed by p66shc knockdown in SCs.

Our data suggest a correlation with autophagy signaling pathways in the presence of oxidative stresses, including ROS. Persistent hyperglycemia during diabetes causes a redox imbalance due to overproduction of ROS. Moderate ROS levels increase autophagosome formation by converting LC3-I to LC3-II to the reduced form of Atg4 via thiol modification of Cys81 of Atg4. However, excessive ROS levels inhibit autophagy—the reduced form of Atg4 delipidates LC3 and inhibits autophagosome membrane elongation [39]. Moderate ROS generation stimulates activation of hypoxia-inducing factor 1α, p53, FoxO3, and nuclear factor erythroid 2-related factor 2, and these transcription factors promote the expression of genes required for autophagy [40]. Several of the transcription factors increased by ROS bind p62 (S405/409), which is phosphorylated to induce autophagy. Furthermore, an appropriate level of H₂O₂ promotes autophagosome formation prior to lipidated LC3-II deconjugation. However, excessive ROS production impairs autophagy. The autophagy-associated proteins ATG7, ATG10, and ATG3 have a sulfhydryl group that is sensitive to ROS oxidation and inactivates autophagy-related genes [41]. Inactivation of these key enzymes leads to autophagy dysfunction. Our data suggest that high glucose inhibits autophagy during autophagosome formation. Also, p66shc downregulation restores the hyperglycemia-impaired autophagy of SCs.

ER stress occurs when the ER processing capacity is exceeded, limits its ability to fold or assemble proteins, or after calcium depletion [42]. ER stress is activated in the peripheral nervous system of DPN mice, and chemical inducers of ER stress trigger neuralgia [43]. In this study, ER stress was increased in the sciatic nerve and high glucose-treated SCs in hyperglycemia-induced DPN mice. The increase in ER stress induced by hyperglycemia was restored by suppression of p66shc expression. This indicates that ER stress is detrimental to SC survival after DPN development, whereas inhibition of ER stress enhances SC survival. DPN-induced hyperglycemia significantly increased ER stress-related gene expression, whereas expression of basic fibroblast growth factor and nerve growth factor decreased ER stress-related gene expression. ER stress is strongly associated with autophagy [44]. To address ER stress, cells induce the unfolded protein response (UPR), which is an adaptive signaling process that triggers a variety of mechanisms to clear misfolded proteins, resolve protein folding, and enhance and maintain ER homeostasis. Autophagy under pathological ER stress, particularly neurodegeneration, is regulated by ERN1 and related pathways of the UPR. The ER degradation enhancer, mannosidase α-like 1, a target gene of XBP1 that encodes an essential component of ERAD substrate signaling and recognition, induces ER stress and thereby interferes with autophagy activation [45].

The autophagy receptor p62 prevents the effects of misfolded proteins on proteins or organelles by sequestering aggregated proteins [32,46]. Whether the UPR generates p62 bodies as a cytoprotective mechanism under ER stress is unknown. We plan to evaluate the association between decreased p62 expression and ER stress. In SCs, suppression of p66shc expression restored high glucose-induced eIF2a phosphorylation and CHOP expression but not GRP78 expression. In sciatic nerves, hyperglycemia-induced p66shc expression, but not eIF2a phosphorylation, was restored. ER stress has long been known to be linked to oxidative stress. Because protein folding is influenced by redox homeostasis, oxidative stress induces ER stress by disrupting protein folding and enhancing the production of misfolded proteins [47]. In this study, ER stress-related gene expression, which is induced by hyperglycemia, was restored by suppression of p66shc expression, albeit differently in cell culture and the mouse model.

Many stress-induced signaling pathways regulate both autophagy and apoptosis. In this study, early apoptosis of SCs was induced by high glucose, and reversed by suppression of p66shc expression. High glucose inhibits axon growth and induces apoptosis of SCs. In addition, exposure to high glucose impairs the morphology and function of SCs [48]. Oxidative stress induced by high glucose can cause DNA damage and trigger apoptosis via p53-related mechanisms. It also activates the calcium-dependent protease calpain, which triggers apoptosis by caspase-dependent and independent mechanisms [49,50]. Many stimuli that normally lead to apoptosis also trigger autophagy; in these cases, autophagy is activated before induction of apoptosis. Autophagy induction is exacerbated by inhibition of apoptosis by removing BCL-2 related X protein and pro-apoptotic proteins such as BCL-2 antagonists or killers or by caspase inhibitors [51]. Excessive response to external signals, such as ionizing radiation, chemotherapy, depletion of essential nutrients, and inhibition of growth factor receptors, causes loss of autophagy control, eventually leading to apoptosis [52]. This suggests that autophagy dysfunction caused by hyperglycemia initiates apoptosis. Numerous questions remain about the relative contributions of stressors to the cascade in which autophagy and apoptosis interact with other cellular signals. We plan to address these questions in further studies.

Overall, we demonstrate that SC dysfunction and p66shc upregulation occurred in diabetes. In addition, regulation of p66shc expression restored impaired autophagy and the functions of SCs and sciatic nerves. Therefore, agents that modulate SC autophagy may have therapeutic potential for sciatic nerve protection in diabetes.

This research was supported by the Basic Science Research Program through the National Research Foundation of Korea (NRF) funded by the Ministry of Education (NRF-2018R1A2B6007425, NRF-2018H1A2A1062309 and NRF-2014R1A6A1029617), the Korea Health Technology R&D Project through the Korea Health Industry Development Institute (KHIDI) funded by the Ministry of Health & Welfare (grant number: HI21C1019) and by the research fund of Chungnam National University. Kaikobad Irani was supported by funding from R01HL147545, R01HL115955, VA Merit Award I01BX002940, and ONR N0001142012147.

None.

The authors declare no conflicts of interest.

Supplementary data including four figures can be found with this article online at https://doi.org/10.4196/kjpp.24.155