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Department of Pharmacology, Southern Illinois University School of Medicine, Springfield, Illinois (R.M.P., D.-S.C., A.P., L.S.P.); and Department of Medical Microbiology, Immunology and Cell Biology & Department of Internal Medicine/Endocrinology, Metabolism and Molecular Medicine, Southern Illinois University School of Medicine Springfield, Illinois (M.E.P.)
Received for publication September 7, 2007.
Accepted for publication December 14, 2007.
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; Sugimoto et al., 2000). A direct effect of hyperglycemia contributing to hyperalgesia has been suggested by various studies (Courteix et al., 1996; Chen and Pan, 2002). However, treatment with insulin-like growth factor or direct neuronal delivery of low doses of insulin, insufficient to reduce hyperglycemia, ameliorated diabetic neuropathy suggesting that mechanisms other than hyperglycemia may be involved in the pathogenesis of hyperalgesia (Zhuang et al., 1997; Brussee et al., 2004).
STZ is transported into β cells of the pancreas through glucose transporter GLUT2 and causes DNA damage either by alkylation, by the generation of nitric oxide (NO), or by the generation of peroxynitrite (Turk et al., 1993; Schnedl et al., 1994; Kröncke et al., 1995). The DNA strand breaks lead to the activation of the nuclear enzyme poly(ADP-ribose) polymerase (PARP), which synthesizes large amounts of the ADP-ribose polymer, using cellular nicotinamide adenine dinucleotide (NAD+) as a substrate (LeDoux et al., 1988; Delaney et al., 1995; Pieper et al., 1999). A decrease in the intracellular NAD+ levels causes a depletion of ATP, a mechanism that can induce cell death (Berger, 1985). Failure of STZ to induce diabetes in PARP-deficient mice suggests the important role played by PARP in STZ-induced β cell necrosis (Burkart et al., 1999). Another proposed mechanism of STZ-induced cytotoxic effect is its selective inhibition of N-acetyl-β-D-glucoaminidase, which is highly expressed in pancreatic β cells compared with neurons (Konrad et al., 2001). Inhibition of N-acetyl-β-D-glucoaminidase results in increased glycosylation of proteins, altering their structure and function and leading to the death of β cells (Konrad et al., 2001). In pancreatic β cells, STZ has been shown to produce superoxide anion by inhibiting the Krebs cycle, which will limit the generation of ATP, promoting the death of β cells (Nukatsuka et al., 1990; Sofue et al., 1991; Turk et al., 1993). However, although the effects of STZ are believed to be specific to pancreatic β cells, STZ administration has been shown to adversely affect renal, hepatic, and muscle tissues (Petzold and Swenberg, 1978; Brambilla et al., 1987; Johnston et al., 2007).
Topical application of capsaicin, a transient receptor potential vanilloid 1 (TRPV1) agonist, improves sensory perception in humans with DPN (Forst et al., 2002). In animal models of diabetes, in which DPN manifests as hyperalgesia, TRPV1 expression and function have been shown to be increased (Kamei et al., 2001; Hong and Wiley, 2005). The TRP family of ion channels is involved in sensing physical and chemical stimuli (Clapham, 2003). TRPV1 is a Ca2+-permeant nonselective cation channel expressed predominantly in unmyelinated C fibers and thinly myelinated A
fibers (Julius and Basbaum, 2001). TRPV1 is activated by heat (>42°C), capsaicin, protons, N-arachidonyl dopamine, anandamide, and leukotrienes (Caterina et al., 1997; Julius and Basbaum, 2001). A role of TRPV1 beyond that of a simple temperature sensor is suggested by the diversity of its agonists and its expression in areas that are not subjected to higher temperatures (Mezey et al., 2000; Birder et al., 2001; Huang et al., 2002). Sensitization of TRPV1 by phosphorylation is mediated by numerous agents, including prostaglandins, bradykinin, glutamate, serotonin, histamine, ATP, and trypsin (Julius and Basbaum, 2001). Modulation by substances associated with inflammation suggests that TRPV1 functions as a sensor of nociceptive inflammatory thermal pain, and the finding is corroborated by a lack of thermal hyperalgesia in TRPV1 knockout mice (Caterina et al., 2000; Davis et al., 2000).
This study was undertaken based on the observation that a proportion of STZ-injected mice did not become diabetic but exhibited hyperalgesia. Here, we sought to determine whether STZ can exert a direct effect on neurons contributing to the hyperalgesia by altering the expression and function of TRPV1.
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Age-matched single transgenic mice (Ins-HA.D2 mice), which express influenza hemagglutinin (HA) peptide, were used and were kindly provided by Dr. Lo (Lo et al., 1992). Freshly prepared STZ (50-200 mg/kg) in saline (pH 4.5 with 0.1 N citrate buffer) was injected intraperitoneally as described previously (Kamei et al., 1991). Control mice received citrate buffered saline alone. Although Ins-HA.D2 mice are genetically altered, HA expression does not seem to affect glucose metabolism or neuronal function as measured by blood glucose levels and hot plate tests compared with normal wild-type mice.
Glucose levels were determined with an OneTouch Ultra blood glucose monitoring system (LifeScan, Milpitas, CA) using whole blood obtained from the tail. Diabetes was defined as blood glucose concentrations greater than 299 mg/dl (16.7 mM) (Kamei et al., 1991).
Determination of Thermal Pain Sensitivity. All of the mice used in this study were housed in the barrier facility. Mice were tested in the barrier facility on the days the cages were not cleaned to avoid factors that might influence the test measurements. Mice were placed individually on a Hot Plate Analgesia Meter (Harvard Apparatus, Boston, MA) maintained at a constant temperature of 52 ± 0.3°C. The paw withdrawal latency (PWL) is defined as the time taken for the animal to exhibit a distinct pain behavior either by a hind paw lick or a characteristic hind paw flick (whichever occurs first). Mice that did not respond within 20 s were removed from the hot plate to prevent tissue damage. We did not find a significant difference in PWL with either the duration or habituation (0.5-2 h). The experiments were conducted in a blind fashion by measuring the PWL in randomly chosen animals from diabetic or control groups. After completing the test, the ear tags were read to place them in the appropriate groups. There were slight gender differences, but they were not statistically significant, therefore male and female mice were grouped together for further analyses.
In Vitro STZ Treatment. To assess the effect of STZ on TRPV1 expression, reactive oxygen species (ROS) production, or cell viability, cultured DRG neurons and stably TRPV1-expressing HEK 293T cells (Puntambekar et al., 2005) were exposed to 10, 40, 100, 300, 400, and 1000 µM STZ. STZ was prepared as a fresh stock solution by dissolving it in 0.1 N citrate buffer, pH 4.5, before adding it to the neuronal medium and incubated for 24 to 72 h. In the experiments that required the exposure of STZ longer than 24 h, the medium was replaced with appropriated doses of STZ. The experimental observations were compared with sister cultures that were treated with 0.1 N citrate buffer.
Electrophysiology. Primary DRG neuronal cultures were prepared from embryonic day 18 rat embryos. Adult pregnant rats were killed with an overdose of isoflurane. DRG were dissected, and the neurons were dissociated by trituration with a fire-polished glass pipette in Hanks' balanced salt solution (Ca2+- and Mg2+-free). Neurons were cultured in neurobasal medium (Invitrogen, Carlsbad, CA), supplemented with L-glutamine and B27 supplement (Invitrogen, Grand Island, NY) and grown on poly(D-lysine)-coated glass coverslips in 24-well plates. Neurons were incubated at 37°C in a humidified atmosphere of 5% CO2. Neurons were used from 3 days after plating.
For whole-cell patch clamp current recordings, the bath solution contained 140 mM sodium gluconate, 2.5 mM KCl, 10 mM HEPES, 1 mM MgCl2, and 1.5 mM EGTA, with pH adjusted to 7.35 with NaOH, and the pipette solution contained 140 mM potassium gluconate, 5 mM KCl, 10 mM HEPES, 2 mM MgCl2, 10 mM EGTA, 2 mM ATP, and 0.25 mM GTP, with pH adjusted to 7.35 with NaOH. Ca2+-free extracellular solution was used to avoid desensitization and tachyphylaxis of capsaicin-induced currents. The junction potential between the patch pipette and the bath solutions was cancelled before the gigaohm seal was formed. The tip of the drug application pipettes was placed within 100 µm of the neurons. Currents were recorded using a WPC-100 patch-clamp amplifier (E.S.F. Electronic, Goettingan, Germany). Data were digitized (VR-10B; InstruTech, Great Neck, NY) and stored in videotapes or directly stored in the computer using a Lab View (National Instruments, Austin, TX) interface. For analysis, data were filtered at 2.5 kHz (-3 dB frequency with an eight-pole low-pass Bessel filter, LPF-8; Warner Instruments, Hamden, CT) and digitized at 5 kHz. Current amplitudes were measured using Channel 2 Software (kindly provided by Michael Smith, Australian National University, Canberra, Australia). The traces and graphs were plotted using Origin Software (OriginLab Corp., Northampton, MA). Capacitance of the cell was measured manually by using the readout in the WPC-100 amplifier.
Measurement of ROS. ROS production was detected using the dye 2',7'-dichlorofluorescein diacetate (DCF-DA; Invitrogen). DCF-DA, a nonfluorescent cell-permeant compound, is cleaved by endogenous esterases and the de-esterified product becomes fluorescent upon oxidation by ROS. Cells were incubated with DCF-DA (20 µM) at 37°C for 20 min and washed twice in Hanks' balanced salt solution to reduce nonspecific fluorescence. Fluorescence measurements were carried out using an inverted microscope (DMIRE2; Leica, Plymouth, MN) equipped with a camera (Retiga Ex; Roper Scientific, Ottobrunn, Germany) and the Lambda DG4 wavelength switcher (Sutter Instrument Company, Novato, CA), and the data were analyzed using Scanalytics software (Scanalytics Inc., Fairfax, VA). The exposure time was kept to <1 s to avoid photo oxidation of the ROS-sensitive dyes, and for all treatments, the exposure time was kept constant. At least three independent fields were chosen for each condition, and 5 to 20 cells in a given field were used for quantification of the fluorescence signals. DCF fluorescence was measured and collected with an excitation (
exc) of 488 nm and emission (emi)of 535 nm.
Ca2+ Imaging. DRG neurons grown on glass coverslips were incubated with Fluo-4 AM (3 µM) (Invitrogen) for 20 min at 37°C and washed with physiological buffer containing 140 mM NaCl, 5 mM HEPES, 2 mM CaCl2, 1 mM MgCl2, 2.5 mM KCl, and 2 mM lido-caine, pH 7.35. The experiments were carried out using a microscope (DMIRE2) attached to a camera (Retiga Ex), and the Lambda DG4 wavelength switcher (Sutter Instruments). Fluo-4 was excited at 488 nm, and the emitted fluorescence was filtered with a 535 ± 25 nm bandpass filter and analyzed using the Scanalytics software. Multiple cells were selected, and the fluorescence of individual cells was tracked. The ratio of the fluorescence change F/Fo was plotted to represent the change in intracellular Ca2+ levels.
Cell Viability Assay. Coverslips containing embryonic DRG neurons were incubated with 15 µl of fluorescein diacetate (FDA; 15 mg/ml) and 15 µl of propidium iodide (PI; 4.6 mg/ml) for 3 min by adding them into 0.5 ml of neurobasal medium. Cells were washed twice with Ca2+-free buffer to prevent nonspecific background fluorescence. Green fluorescence (exc 488 nm and emi 520 nm wavelengths for FDA) for live and red fluorescence (exc 535 nm and emi 590-615 nm wavelengths for PI) for dead cells were observed using a microscope (DMIRE2) equipped with a camera (Retiga Ex) and the Lambda DG4 wavelength switcher (Sutter Instruments) and were analyzed using Scanalytics software. At least five to six independent fields were chosen for analysis for each condition. The percentage of survival or viability of DRG neurons was calculated for different concentrations of STZ.
Western Blot. Mice were sacrificed 1 week after STZ treatment, and DRG were removed and placed in a lysis buffer (0.1% SDS, 1% Triton X-100, 1% deoxycholate, protease, and phosphatase inhibitor cocktail, 1:100; Sigma, St. Louis, MO), homogenized, and centrifuged. Stably TRPV1-expressing HEK 293T cells were cultured as described previously (Puntambekar et al., 2005). TRPV1-expressing HEK cells were scraped into a 400-µl lysis buffer and centrifuged. The protein concentration was measured by the bicinchoninic acid assay. Proteins were separated by 10% SDS-polyacrylamide gel electrophoresis and transferred to the nitrocellulose membrane (Bio-Rad Laboratories, Hercules, CA). Membranes were probed overnight with rabbit anti-p38, phospho-p38 (1:200; Santa Cruz Biotechnology, Santa Cruz, CA), β-Actin (1:200; Sigma), or goat anti-TRPV1 (1:100; Santa Cruz Biotechnology) antibodies followed by incubation with horseradish peroxidase-conjugated goat anti-rabbit IgG or rabbit anti-goat IgG (1:10,000, Santa Cruz Biotechnology) for 1 h. After incubation with enhanced chemiluminescence reagents (Santa Cruz Biotechnology), membranes were scanned using the Hitachi genetic systems (Hitachi Software Engineering, Tokyo, Japan), and blots were analyzed using GeneTools Analysis Software (SynGene, Frederick, MD).
Data Analysis. For behavioral experiments, mixed model analysis was performed using SAS/STAT software (SAS Institute, Cary, NC), which included both fixed events (age and time after diabetes onset) and random events (number of subjects) with repeated measures of analysis of variance. The comparisons were made between control group and diabetic groups. Data are shown as mean ± S.E.M.. Data are considered significant at p < 0.05. For all other experiments, data are shown as mean ± S.E.M. Significance is tested using unpaired Student's t test, and the data are considered significant at p < 0.05. DCF and Fluo-4 AM were obtained from Invitrogen. B27-supplement was obtained from Invitrogen. p38 MAPK inhibitor SB203580 was obtained from Santa Cruz Biotechnology. All other chemicals used in this study were obtained from Sigma.
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Alterations in Thermal Pain Sensitivity in STZ-Injected Nondiabetic Mice. A proportion (20-25%) of STZ-injected mice did not become diabetic as indicated by the blood sugar levels (<300 mg/dl), but these mice exhibited thermal hyperalgesia. This was further confirmed by a nondiabetogenic dose of STZ (50 mg/kg). The blood glucose levels were slightly higher compared with vehicle-injected mice (STZ-injected nondiabetic mice, 170 ± 8.8; week 2, 222.4 ± 24.3; week 6, 211.4 ± 19.6 mg/dl, n = 13; vehicle-injected mice, 157 ± 6.1; week 2, 163.8 ± 3.9; week 6, 186 ± 1.6 mg/dl, n = 6) (Fig. 2A). Body weights of STZ-injected nondiabetic and vehicle-injected mice increased steadily (STZ-injected nondiabetic mice week 1, 23 ± 1; and week 6, 25.2 ± 0.9 g, n = 13) (vehicle-injected mice week 1, 22.1 ± 0.2; week 6, 26.2 ± 0.3 g, n = 6) (Fig. 2B). STZ-injected nondiabetic mice exhibited a phase of hyperalgesia (STZ-injected nondiabetic mice, 9.6 ± 0.5 s; week 2, 6.1 ± 0.3 s, n = 13; p < 0.001) followed by a phase of normal PWL (STZ-injected nondiabetic mice week 7, 7.9 ± 0.6 s, n = 13; vehicle-injected mice, week 8, 9 ± 0.8 s, n = 6) (Fig. 2C). These results suggest that STZ treatment induces changes in thermal pain sensitivity, which is independent of hyperglycemia.
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Incubation of Neurons with STZ Increases TRPV1-Mediated Current Responses. To determine whether STZ has a direct action on neurons, cultured embryonic DRG neurons were incubated for 24 h with different concentrations of STZ (20-400 µM), and capsaicin (1 µM)-induced TRPV1-mediated whole-cell currents were recorded. There was a dose-dependent increase in TRPV1 current at lower concentrations followed by a decrease at higher concentrations. Vehicle-treated sister cultures were used as controls. TRPV1-mediated currents were significantly higher at 20 µM (control, 1 ± 0.1, n = 10; STZ, 2.0 ± 0.1-fold, n = 10, p < 0.004) and 100 µM STZ (control, 1 ± 0.1, n = 9; STZ, 2.5 ± 0.2-fold, n = 9, p < 0.04), but there was no significant change at 200 µM STZ (control, 1 ± 0.3, n = 10; STZ, 1.1 ± 0.2-fold, n = 10), and a decrease in the current amplitude was observed at 400 µM STZ (control, 1 ± 0.2, n = 6; STZ, 0.6 ± 0.2-fold, n = 6) (Fig. 3B). Typically, small-diameter (<25 µm) neurons responded to capsaicin. Average capacitance of DRG neurons in 100 µM STZ-treated neurons (19.2 ± 1.48 pF, n = 9) did not differ from that of control neurons (21.5 ± 1.7 pF, n = 11). Peak current amplitudes were expressed as current densities (in picoamperes per picofarads) using cell capacitance to normalize the differences in cell size, which was found to be significantly higher at 20 and 100 µM STZ concentrations (control, 1 ± 0.2, n = 10; 20 µM STZ, 1.7 ± 0.2-fold, n = 10, p < 0.004) (control, 1 ± 0.2, n = 9; 100 µM STZ, 2.1 ± 0.2-fold, n = 9; p < 0.05) but not at 200 and 400 µM STZ (Fig. 3C). These results demonstrate that lower concentrations of STZ (20 and 100 µM) induce an increase in TRPV1-mediated currents, whereas higher concentrations (>200 µM) cause a decrease, possibly as a result of reaching toxic concentrations. Elevated TRPV1 currents may be one of the mechanisms by which neuronal excitability increases, contributing to hyperalgesia observed in STZ-injected diabetic and nondiabetic mice.
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We noticed a decrease in TRPV1-mediated current when the neurons were exposed to higher concentrations of STZ (>200 µM). Studies have suggested that 1 mM STZ induces pancreatic β cell death in vitro (Nukatsuka et al., 1990; Konrad et al., 2001). Based on these studies, we hypothesized that STZ might alter the viability of DRG neurons. To test whether STZ can alter the viability of neurons, we performed a viability assay using live and dead cell staining dyes FDA and PI, respectively. Results of the viability assay suggest that increasing concentrations of STZ decreased the viability of embryonic DRG neurons (200 µM, 26%, n = 180; 400 µM, 27.4%, n = 114; and 1000 µM STZ, 16.4%, n = 64, viable cells) compared with vehicle-treated sister cultures.
STZ Treatment Induces ROS Production in Cultured DRG Neurons in Vitro. It has been suggested that short- and long-term treatments of STZ induce myopathy through the ROS-mediated mechanism (Johnston et al., 2007). Likewise, nerve growth factor (NGF) induces TRPV1 expression through NADPH oxidase-dependent ROS pathway (Suzukawa et al., 2000; Puntambekar et al., 2005). Therefore, we determined whether ROS is involved in the STZ-induced increase in TRPV1 currents. Initially, we tested whether a known ROS-generating agent, such as hydrogen peroxide, (H2O2, 25 µM) could increase DCF fluorescence intensity in cultured DRG neurons treated for 24 h. As expected, H2O2 significantly increased DCF fluorescence (data not shown). Next, to determine the role of STZ, we pretreated the cultured embryonic DRG neurons with different concentrations of STZ for 24 h, and changes in ROS were measured. We observed a higher DCF fluorescence intensity with increasing concentrations of STZ (citrate buffer, 1 ± 0.03, n = 170 cells; 100 µM STZ, 1.8 ± 0.04, n = 105 cells, p < 0.0001; 400 µM STZ, 1.77 ± 0.05, n = 75 cells, p < 0.0001) (Fig. 5). These results suggest that STZ treatment induces ROS production in cultured DRG neurons, which might play a role in increasing TRPV1-mediated current through ROS-mediated transcriptional or translational regulation. Although ROS levels were higher even at a higher concentration of STZ (400 µM) treatment, TRPV1-mediated currents showed a reduction (Fig. 3), suggesting that higher levels of ROS may become toxic to neurons.
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Activation of STZ-ROS-p38 MAPK Pathway Promotes TRPV1 Expression. Previous studies have suggested that JNK and p38 MAPK are strongly activated by ROS or by a mild oxidative shift of the intracellular thiol/disulfide redox state (Abe et al., 1996; Hehner et al., 2000). A similar mechanism is also involved in the NGF-induced increase in TRPV1 expression, in which p38 MAPK is activated by ROS, thereby contributing to hyperalgesia (Ji et al., 2002; Puntambekar et al., 2005). Experiments were conducted using stably TRPV1-expressing HEK 293T cells to determine whether the increase in TRPV1 expression induced by long-term STZ treatment is mediated by the ROS-p38 MAPK pathway. Cells were treated with STZ, ROS-generating and -scavenging agents, and then probed for TRPV1 and p-p38 MAPK. Consistent with our previous results, STZ (100 µM)-treated cells exhibited an elevated TRPV1 expression (1.52 ± 0.13-fold, n = 4, p < 0.01) (Fig. 8B). Removal of ROS by catalase (200 U/ml) abolished this increase (1.08 ± 0.02-fold, n = 4, p < 0.05). p-p38 MAPK was also found to be increased in STZ-treated cells (1.25 ± 0.04-fold, n = 4, p < 0.01) but did not increase when treated simultaneously with catalase (0.98 ± 0.04-fold, n = 4, p < 0.05) (Fig. 8B). In support of our findings, we successfully reproduced these results with a ROS-generating agent, H2O2 (25 µM) (TRPV1: H2O2, 1.4 ± 0.1-fold, n = 4, p < 0.01; H2O2 + catalase, 1.01 ± 0.09-fold, n = 4, p < 0.05) (p-p38 MAPK: H2O2, 1.48 ± 0.09-fold, n = 4, p < 0.01; H2O2 + catalase, 0.92 ± 0.06-fold, n = 4, p < 0.05) (Fig. 8C), suggesting that STZ-induced TRPV1 expression is mediated by the ROS-p38 MAPK pathway. To confirm that the effect is mediated by p38 MAPK, we incubated the cells with a p38 MAPK inhibitor, SB203580 (20 µM). The increase in TRPV1 and p-p38 MAPK levels was significantly reduced after treatment with the inhibitor (TRPV1, 1.08 ± 0.11-fold, n = 3, p < 0.05; p-p38 MAPK, 0.95 ± 0.08-fold, n = 3, p < 0.05) (Fig. 8D).
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20-25%) of STZ-treated mice did not become diabetic but became hyperalgesic compared with the vehicle-injected group. It was further confirmed that hyperalgesia was observed in nondiabetogenic doses of STZ. However, we could not find a correlation between the degree of hyperalgesia and the dose of STZ. Possible reasons for the failure of STZ to induce diabetes are 1) inability to damage all pancreatic β cells as a result of insufficient concentration; 2) faster recovery of damaged pancreatic β cells; and 3) decreased bioavailability as a result of rapid breakdown by the liver enzymes or rapid excretion by the kidneys. A normal gain of body weight in STZ-injected nondiabetic mice and a decrease in the body weight of diabetic mice suggest that STZ-injected nondiabetic mice do not suffer from metabolic derangement. Both time course and the degree of thermal hyperalgesia were similar in all STZ-treated mice, regardless of their blood glucose levels after STZ injection. Other investigators have not reported this phenomenon, perhaps because of the common practice of excluding nondiabetic mice from studies. However, it has been reported in a study that 40 to 60% of rats injected with STZ were nondiabetic but exhibited mechanical hyperalgesia and insulinopenia (Romanovsky et al., 2004). It was suggested that insulinopenia itself or consequences of insulinopenia that is independent of hyperglycemia might contribute to mechanical hyperalgesia. A direct effect of STZ was ruled out as a possible causative factor of mechanical hyperalgesia because insulin-treated STZ-injected diabetic rats became normoglycemic and exhibited attenuated pain sensation (Courteix et al., 1996).
TRPV1 plays a role in both diabetic and inflammatory hyperalgesia (Caterina et al., 2000; Hong and Wiley, 2005). STZ destroys pancreatic β cells by elevating the ROS levels (Nukatsuka et al., 1990; Sofue et al., 1991; Turk et al., 1993). In another study, moderate increases in ROS levels have been shown to function as secondary messengers that can influence redox-sensitive signal transduction pathways (Suzukawa et al., 2000). In our in vitro studies, STZ-treated neurons exhibited an increase in TRPV1-mediated currents and an increase in the levels of ROS. By recording TRPV1-mediated whole-cell currents in cultured embryonic DRG neurons treated with ROS-generating and -scavenging agents, we have shown that STZ induces TRPV1 expression through the ROS-mediated pathway. We further observed that STZ also caused an increase in the phosphorylated form of p38 MAPK, suggesting that the increase in the TRPV1 protein expression may involve the ROS-p38 MAPK pathway. However, at higher concentrations of STZ and higher levels of ROS, the effect of toxicity is seen as a decrease in TRPV1 expression and a reduction in cell viability.
Peripheral nerve injury leads to an increase in the expression of brain-derived neurotrophic factor and the 
2 Ca2+ channel subunit through activation of p38 MAPK (Zhou et al., 1999; Luo et al., 2001). A similar mechanism is also involved in NGF-induced increase in TRPV1 expression, in which p38 MAPK is activated by ROS, thereby contributing to hyperalgesia (Ji et al., 2002; Puntambekar et al., 2005). Ji et al. (2002) have proposed that p38 MAPK activates the translational factor eIF4E via MAPK-interacting kinases 1 and 2 and the phosphorylation of which results in an increased affinity of eIF4E for capped RNA influencing translation. Different oxidative stress-inducing stimuli, such as relatively low concentrations of hydrogen peroxide, UV light, 
-irradiation, and interleukin-1, have been shown to activate the activator protein-1, a transcription factor (Angel and Karin, 1991; Devary et al., 1991; Meyer et al., 1993). Likewise, STZ-ROS-p38 MAPK pathway could increase the activity of transcription factors like activator protein-1, inducing transcription of TRPV1, or it may increase the translation of TRPV1 through elF4E. This could lead to an increase in membrane expression and function of TRPV1, thereby contributing to the hyperalgesia observed in STZ-injected nondiabetic mice. Nevertheless, a similar mechanism cannot be ruled out in STZ-injected diabetic mice. In HEK cells stably transfected with TRPV1, treatment of STZ caused a similar increase in TRPV1 and p-p38 MAPK expression suggesting that the effect we observe is post-translational rather than post-transcriptional.
It has been suggested that STZ is transported into β cells through the glucose transporter GLUT2 (Schnedl et al., 1994). It is intriguing that the predominant glucose transporter expressed in the peripheral nervous system is GLUT3. STZ treatment may result in PARP activation, possibly through its direct damaging effects of DNA. PARP activation results in an increased production of poly(ADP-ribose), from which adenine 5'-diphosphoribose is generated using cellular NAD+ as a substrate (LeDoux et al., 1988; Delaney et al., 1995; Pieper et al., 1999). ADP-ribose is a potent activator of TRPMelastatin 2, which has been suggested to play a role in sensing oxidative stress (Fonfria et al., 2004).
Together, our results suggest that STZ exerts a direct action on neurons altering the expression and function of TRPV1 via the ROS-p38 MAPK pathway-mediated translational regulation. Of course, we cannot rule out the possibility that STZ might exert its effects on other nociceptive ion channels such as the voltage-gated sodium channels and the mechanosensitive channels. Therefore, future studies are needed to address the extent of modulation of other ion channels by direct action of STZ (Hayase et al., 2007). To our knowledge, this is the first report suggesting that STZ might have a direct effect on neurons; thus, caution should be exercised in interpreting data as diabetes- or hypoglycemia-induced while using STZ as a diabetogenic agent.
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Acknowledgements |
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R.M.P. and D.-S.C. contributed equally to this work.
ABBREVIATIONS: STZ, streptozotocin; DPN, diabetic peripheral neuropathy; DRG, dorsal root ganglion; FDA, fluorescein diacetate; HA, hemagglutinin; HEK, human embryonic kidney; PWL, paw withdrawal latency; PI, propidium iodide; PARP, poly(ADP-ribose) polymerase; ROS, reactive oxygen species; MAPK, mitogen-activated protein kinase; AM, acetoxymethyl ester; TRPV1, transient receptor potential vanilloid 1; DCF-DA, 2',7'-dichlorofluorescein diacetate; NGF, nerve growth factor; SB203580, 4-(4-fluorophenyl)-2-(4-methylsulfinylphenyl)-5-(4-pyridyl)1H-imidazole.
Address correspondence to: Dr. Louis S. Premkumar, Department of Pharmacology, Southern Illinois University School of Medicine, Springfield, IL 62702. E-mail: lpremkumar{at}siumed.edu
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References |
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) compared with control mice (
). B, the body weight of diabetic mice (
