Early Painful Diabetic Neuropathy Is Associated with Differential Changes in Tetrodotoxin-sensitive and -resistant Sodium Channels in Dorsal Root Ganglion Neurons in the Rat
2004; Elsevier BV; Volume: 279; Issue: 28 Linguagem: Inglês
10.1074/jbc.m404167200
ISSN1083-351X
AutoresShuangsong Hong, Thomas Morrow, Pamela E. Paulson, Lori L. Isom, John Wiley,
Tópico(s)Botulinum Toxin and Related Neurological Disorders
ResumoDiabetic neuropathy is a common form of peripheral neuropathy, yet the mechanisms responsible for pain in this disease are poorly understood. Alterations in the expression and function of voltage-gated tetrodotoxin-resistant (TTX-R) sodium channels have been implicated in animal models of neuropathic pain, including models of diabetic neuropathy. We investigated the expression and function of TTX-sensitive (TTX-S) and TTX-R sodium channels in dorsal root ganglion (DRG) neurons and the responses to thermal hyperalgesia and mechanical allodynia in streptozotocin-treated rats between 4–8 weeks after onset of diabetes. Diabetic rats demonstrated a significant reduction in the threshold for escape from innocuous mechanical pressure (allodynia) and a reduction in the latency to withdrawal from a noxious thermal stimulus (hyperalgesia). Both TTX-S and TTX-R sodium currents increased significantly in small DRG neurons isolated from diabetic rats. The voltage-dependent activation and steady-state inactivation curves for these currents were shifted negatively. TTX-S currents induced by fast or slow voltage ramps increased markedly in neurons from diabetic rats. Immunoblots and immunofluorescence staining demonstrated significant increases in the expression of Nav1.3 (TTX-S) and Nav 1.7 (TTX-S) and decreases in the expression of Nav 1.6 (TTX-S) and Nav1.8 (TTX-R) in diabetic rats. The level of serine/threonine phosphorylation of Nav 1.6 and In Nav1.8 increased in response to diabetes. addition, increased tyrosine phosphorylation of Nav1.6 and Nav1.7 was observed in DRGs from diabetic rats. These results suggest that both TTX-S and TTX-R sodium channels play important roles and that differential phosphorylation of sodium channels involving both serine/threonine and tyrosine sites contributes to painful diabetic neuropathy. Diabetic neuropathy is a common form of peripheral neuropathy, yet the mechanisms responsible for pain in this disease are poorly understood. Alterations in the expression and function of voltage-gated tetrodotoxin-resistant (TTX-R) sodium channels have been implicated in animal models of neuropathic pain, including models of diabetic neuropathy. We investigated the expression and function of TTX-sensitive (TTX-S) and TTX-R sodium channels in dorsal root ganglion (DRG) neurons and the responses to thermal hyperalgesia and mechanical allodynia in streptozotocin-treated rats between 4–8 weeks after onset of diabetes. Diabetic rats demonstrated a significant reduction in the threshold for escape from innocuous mechanical pressure (allodynia) and a reduction in the latency to withdrawal from a noxious thermal stimulus (hyperalgesia). Both TTX-S and TTX-R sodium currents increased significantly in small DRG neurons isolated from diabetic rats. The voltage-dependent activation and steady-state inactivation curves for these currents were shifted negatively. TTX-S currents induced by fast or slow voltage ramps increased markedly in neurons from diabetic rats. Immunoblots and immunofluorescence staining demonstrated significant increases in the expression of Nav1.3 (TTX-S) and Nav 1.7 (TTX-S) and decreases in the expression of Nav 1.6 (TTX-S) and Nav1.8 (TTX-R) in diabetic rats. The level of serine/threonine phosphorylation of Nav 1.6 and In Nav1.8 increased in response to diabetes. addition, increased tyrosine phosphorylation of Nav1.6 and Nav1.7 was observed in DRGs from diabetic rats. These results suggest that both TTX-S and TTX-R sodium channels play important roles and that differential phosphorylation of sodium channels involving both serine/threonine and tyrosine sites contributes to painful diabetic neuropathy. Diabetes mellitus is one of the most common chronic medical problems, affecting over 100 million people world-wide (1Spruce M.C. Potter J. Coppini D.V. Diabet. Med. 2003; 20: 88-98Crossref PubMed Scopus (107) Google Scholar). Diabetic patients frequently exhibit one or more types of stimulus-evoked pain, including increased responsiveness to noxious stimuli (hyperalgesia) as well as hyper-responsiveness to normally innocuous stimuli (allodynia). The underlying mechanisms of persistent pain in diabetic patients remain poorly understood. In animal models of diabetes, hyperalgesia to nonnoxious thermal stimulation as well as tactile allodynia have been observed (2Calcutt N.A. Jorge M.C. Yaksh T.L. Chaplan S.R. Pain. 1996; 68: 293-299Abstract Full Text Full Text PDF PubMed Scopus (202) Google Scholar, 3Fox A. Eastwood C. Gentry C. Manning D. Urban L. Pain. 1999; 81: 307-316Abstract Full Text Full Text PDF PubMed Scopus (157) Google Scholar, 4Malcangio M. Tomlinson D.R. Pain. 1998; 76: 151-157Abstract Full Text Full Text PDF PubMed Scopus (209) Google Scholar). The streptozotocin (STZ) 1The abbreviations used are: STZ, streptozotocin; TTX-R, voltage-gated tetrodotoxin-resistant sodium channels; TTX-S, voltage-gated tetrodotoxin-sensitive sodium channels; DRG, dorsal root ganglion; F, farad.1The abbreviations used are: STZ, streptozotocin; TTX-R, voltage-gated tetrodotoxin-resistant sodium channels; TTX-S, voltage-gated tetrodotoxin-sensitive sodium channels; DRG, dorsal root ganglion; F, farad.-induced diabetic rat model demonstrates many of the abnormalities observed in humans (5Stevens M.J. Dananberg J. Feldman E.L. Lattimer S.A. Kamijo M. Thomas T.P. Shindo H. Sima A.A.F. Greene D.A. J. Clin. Investig. 1994; 94: 853-859Crossref PubMed Google Scholar). Treatment with insulin prevents development or reverses many of the abnormalities observed in early painful diabetic neuropathy (6Srinivasan S. Stevens M. Wiley J.W. Diabetes. 2000; 49: 1932-1938Crossref PubMed Scopus (225) Google Scholar, 7Barber A.J. Lieth E. Khin S.A. Antonetti D.A. Buchanan A.G. Gardner T.W. J. Clin. Investig. 1998; 102: 783-791Crossref PubMed Scopus (1038) Google Scholar). In diabetic rats with hyperalgesia, dorsal root ganglion (DRG) neurons display increased frequency of action potential generation in response to sustained suprathreshold mechanical stimulation (3Fox A. Eastwood C. Gentry C. Manning D. Urban L. Pain. 1999; 81: 307-316Abstract Full Text Full Text PDF PubMed Scopus (157) Google Scholar, 4Malcangio M. Tomlinson D.R. Pain. 1998; 76: 151-157Abstract Full Text Full Text PDF PubMed Scopus (209) Google Scholar, 8Ahlgren S.C. Wang J.F. Levine J.D. Neuroscience. 1997; 76: 285-290Crossref PubMed Scopus (64) Google Scholar, 9Ahlgren S.C. Levine J.D. Neuroscience. 1993; 52: 1049-1055Crossref PubMed Scopus (114) Google Scholar, 10Ahlgren S.C. Levine J.D. J. Neurophysiol. 1994; 72: 684-692Crossref PubMed Scopus (109) Google Scholar) and increased spontaneous activity (11Said G. J. Neurol. 1996; 243: 431-440Crossref PubMed Scopus (56) Google Scholar). Both effects are thought to contribute to the sensation of pain. Voltage-gated sodium channels generate and propagate action potentials in excitable cells. Based on differential sensitivity to tetrodotoxin (TTX), sodium currents in DRG neurons are classified into TTX-sensitive (TTX-S) and TTX-resistant (TTX-R) components (12Caffrey J.M. Eng D.L. Black J.A. Waxman S.G. Kocsis J.D. Brain Res. 1992; 592: 283-297Crossref PubMed Scopus (268) Google Scholar, 13Kostyuk P.G. Veselovsky N.S. Tsyndrenko A.Y. Neuroscience. 1981; 6: 2423-2430Crossref PubMed Scopus (314) Google Scholar, 14Roy M.L. Narahashi T. J. Neurosci. 1992; 12: 2104-2111Crossref PubMed Google Scholar). At least two TTX-S sodium channel α-subunits, Nav1.6, and Nav1.7, are constitutively expressed in the peripheral nervous system (15Ogata N. Ohishi Y. Jpn. J. Pharmacol. 2002; 88: 365-377Crossref PubMed Scopus (149) Google Scholar). In addition, Nav1.3, a TTX-S sodium channel that is normally expressed during embryonic development, is up-regulated in the peripheral nervous system following nerve injury (16Black J.A. Cummins T.R. Plumpton C. Chen Y.H. Hormuzdiar W. Clare J.J. Waxman S.G. J. Neurophysiol. 1999; 82: 2776-2785Crossref PubMed Scopus (266) Google Scholar). Two TTX-R sodium channels, Nav1.8 (17Akopian A.N. Sivilotti L. Wood J.N. Nature. 1996; 379: 257-262Crossref PubMed Scopus (905) Google Scholar) and Nav1.9 (18Tate S. Benn S. Hick C. Trezise D. John V. Mannion R.J. Costigan M. Plumpton C. Grose D. Gladwell Z. Kendall G. Dale K. Bountra C. Woolf C.J. Nat. Neurosci. 1998; 1: 653-655Crossref PubMed Scopus (261) Google Scholar, 19Dib-Hajj S.D. Tyrrell L. Black J.A. Waxman S.G. Proc. Natl. Acad. Sci. U. S. A. 1998; 95: 8963-8968Crossref PubMed Scopus (455) Google Scholar) have been identified in DRG neurons and changes in their expression levels have been implicated in painful diabetic neuropathy (20Coward K. Plumpton C. Facer P. Birch R. Carlstedt T. Tate S. Bountra C. Anand P. Pain. 2000; 85: 41-50Abstract Full Text Full Text PDF PubMed Scopus (214) Google Scholar, 21Hirade M. Yasuda H. Omatsu-Kanbe M. Kikkawa R. Kitasato H. Neuroscience. 1999; 90: 933-939Crossref PubMed Scopus (66) Google Scholar, 22Novakovic S.D. Tzoumaka E. McGivern J.G. Haraguchi M. Sangameswaran L. Gogas K.R. Eglen R.M. Hunter J.C. J. Neurosci. 1998; 18: 2174-2187Crossref PubMed Google Scholar, 23Waxman S.G. Pain. 1999; 6: S133-S140Abstract Full Text PDF PubMed Scopus (149) Google Scholar). TTX-S sodium channels in brain are composed of a pore-forming α-subunit and one or two auxiliary β-subunits (24Yu F.H. Catterall W.A. Genome Biol. 2003; 4: 207Crossref PubMed Scopus (496) Google Scholar, 25Isom L.L. Sod. Channel Neur. Hyperexc. 2002; 241: 124-143Google Scholar, 26Catterall W.A. Neuron. 2000; 26: 13-25Abstract Full Text Full Text PDF PubMed Scopus (1670) Google Scholar, 27Blackburn-Munro G. Fleetwood-Walker S.M. Neuroscience. 1999; 90: 153-164Crossref PubMed Scopus (47) Google Scholar). The subunit composition of TTX-R sodium channels, however, is not clear. Increased TTX-R sodium current (21Hirade M. Yasuda H. Omatsu-Kanbe M. Kikkawa R. Kitasato H. Neuroscience. 1999; 90: 933-939Crossref PubMed Scopus (66) Google Scholar), but decreased expression levels of Nav1.8 mRNA and protein have been reported in models of diabetic neuropathy (28Craner M.J. Klein J.P. Renganathan M. Black J.A. Waxman S.G. Ann. Neurol. 2002; 52: 786-792Crossref PubMed Scopus (154) Google Scholar). However, a systematic analysis of the relative contributions of TTX-S and TTX-R sodium channels, including their phosphorylation status, has not been performed in animal models with documented painful diabetic neuropathy. In the present study, we investigated the expression and functional properties of TTX-S and TTX-R sodium channels in acutely dissociated small to medium sized (nociceptive) DRG neurons isolated from diabetic rats with documented painful neuropathy. We demonstrate that TTX-S and TTX-R sodium currents increased significantly and the voltage-dependent activation and steady-state inactivation curves were negatively shifted in these DRG somas. TTX-S currents induced by both fast and slow voltage ramps increased significantly in diabetic neurons. The protein expression levels of Nav1.3 and Nav1.7 increased in DRG homogenates from diabetic animals. In contrast, the protein expression levels of Nav1.6 and Nav1.8 decreased in DRG homogenates. Interestingly, serine/threonine phosphorylation of Nav1.6 and Nav1.8 and tyrosine phosphorylation of Nav1.6 and Nav1.7 increased in neurons from diabetic rats. We propose that Nav1.8 phosphorylation may, in part, be responsible for the increased TTX-R current observed in diabetic neurons. Using immunofluorescence staining, we observed that the expression of Nav1.7 increased while the expression of Nav1.8 decreased in small C-fiber neurons of diabetic rats compared with controls, in agreement with our Western blot results. These data support that the abnormal function of nociceptive fibers observed in early painful diabetic neuropathy involves both TTX-S and TTX-R sodium channels. All experiments were approved by the University of Michigan Committee on Use and Care of Animals according to National Institutes of Health guidelines. Animal Model—Male Sprague-Dawley (Harlan, Indianapolis, IN) rats weighing 180–200 g were fasted overnight to maximize the effectiveness of STZ treatment. Diabetes mellitus was induced by a single injection of STZ (Sigma) at a dose of 45 mg/kg body weight in a citrate buffer (pH 4.5). Age-matched rats in the control group received injections of saline vehicle. STZ-injected animals were given 10% sucrose water for 48 h after injection to prevent hypoglycemia. Tail vein blood glucose levels were measured 48 h after injection and the onset of the diabetic condition was defined as glucose levels greater than 300 mg/dl. Animals were euthanized for study 4 to 8 weeks after induction of diabetes. Our previous studies with this model indicated that rats with diabetes for 4–8 weeks demonstrate a variety of functional abnormalities including delayed nerve conduction velocity, increased calcium influx, impaired inhibitory G protein function, impaired mitochondrial function, and activation of the apoptosis cascade in acutely dissociated DRG neurons that were reversible after 2 weeks of insulin-mediated euglycemia (6Srinivasan S. Stevens M. Wiley J.W. Diabetes. 2000; 49: 1932-1938Crossref PubMed Scopus (225) Google Scholar, 7Barber A.J. Lieth E. Khin S.A. Antonetti D.A. Buchanan A.G. Gardner T.W. J. Clin. Investig. 1998; 102: 783-791Crossref PubMed Scopus (1038) Google Scholar). Behavioral Tests—Prior to electrophysiological studies, a subset of STZ-treated diabetic rats and age-matched healthy controls were evaluated for changes in sensory perception as described below. All animals were acclimated for 1 week prior to testing. During this period the animals were handled extensively and habituated to the behavioral testing procedures. Mechanical Allodynia—To quantify mechanical sensitivity of the foot, brisk foot withdrawal in response to a normally innocuous mechanical stimulus was measured as described previously (30Moller K.A. Johansson B. Berge O.G. J. Neurosci. Methods. 1998; 84: 41-47Crossref PubMed Scopus (89) Google Scholar). Response to the mechanical stimulus was measured with a calibrated electronic von Frey pressure algometer (Somedic Sales AB). This system consists of a hand-held electronic von Frey probe with a circular probe tip of 0.5 mm in diameter. The algometer is connected to a computerized data collection system, allowing on-line display of the applied force as well as rate of stimulus application. The rat was placed in a hanging cage with a metal mesh floor and acclimated for 10 min. A 0.5-mm diameter von Frey probe was manually applied to the plantar surface of the hind foot with a pressure increasing by ∼0.05 Newtons/s and the pressure at which a paw withdrawal occurred was recorded. For each hind paw, the procedure was repeated 5 times and the average pressure to produce a withdrawal computed. Successive stimuli were applied to alternating feet at ∼30-s intervals. A significant decrease in the pressure necessary to elicit a brisk foot withdrawal in response to this mechanical stimulus was interpreted as mechanical allodynia. Thermal Hyperalgesia (Hargreaves Test)—To quantify thermal sensitivity, rats were placed in a clear a Plexiglas chamber (10 cm × 20 cm × 10 cm) located on an elevated floor of clear glass (2-mm thick) and given 5–10 min to habituate. The glass floor was maintained at 30 ± 1 °C. A radiant heat source delivered a thermal stimulus to the plantar surface of each hind foot. The latency to foot withdrawal (escape) served as the behavioral measure of thermal nociception. If the foot was not withdrawn within 20 s, the stimulus was automatically terminated to avoid tissue damage. Each foot was tested 5 times with 3 min between stimulations of either foot to avoid peripheral sensitization effects. The mean withdrawal latency for each foot was computed by averaging the 5 measurements. As compared with the baseline (control) latency, a significant decrease in the latency of foot withdrawal in response to the thermal stimulus was interpreted as indicating the presence of thermal hyperalgesia (31Hargreaves K. Dubner R. Brown F. Flores C. Joris J. Pain. 1988; 32: 77-88Abstract Full Text PDF PubMed Scopus (4215) Google Scholar). Cell Preparation—DRGs were isolated from acutely dissociated thoracic and lumbar regions of the spinal column, and neurons were prepared according to the methods described previously (32Hall K.E. Liu J. Sima A.A. Wiley J.W. J. Neurophysiol. 2001; 86: 760-770Crossref PubMed Scopus (82) Google Scholar). Briefly, ganglia were digested with 0.3% collagenase (Worthington Type 2) in a minimal essential medium (MEM, Invitrogen) supplemented with 16.5 mm NaHCO3 and 28.2 mm glucose (M-MEM) for 50 min and then 0.1% trypsin (type 1, Sigma) for 10 min. The digested DRGs were centrifuged in 2% bovine serum albumin in M-MEM at 4 °C for 5 min and washed twice with M-MEM solution. After titration in M-MEM with additional 10% fetal bovine serum (Invitrogen), DRG neurons were plated on 35-mm sterile culture dishes coated with calf collagen. Isolated neurons were incubated in 95% air + 5% CO2 at 37 °C for 2–7 h prior to the recording. Whole Cell Patch Clamp Recording—Sodium currents were recorded in the whole cell patch clamp configuration using an Axopatch 200B amplifier (Axon Instruments, Foster City, CA) at room temperature (20–23 °C). Electrodes (1–2 MΩ) were pulled from standard wall glass pipettes (G150F-4, Warner Instrument) using a horizontal puller P-87 (Sutter Instrument) and filled with (in mm): 100 CsCl, 30 tetraethylammonium-Cl, 5 NaCl, 2 MgCl2, 1 CaCl2, 3 EGTA, 10 HEPES, 2 Mg-ATP, 1 Li-GTP; pH was adjusted to 7.2 by Tris base and osmolarity was adjusted to 285 mOsm. Pipette solutions were filtered at 0.2 μm immediately before use. The bath solution used to record currents contained (in mm): 35 NaCl, 30 tetraethylammonium-Cl, 65 choline-Cl, 0.1 CaCl2, 5 MgCl2, 0.05 CdCl2, 10 HEPES, and 11 glucose. The pH was adjusted to 7.4 and osmolarity was 325 mOsm. Reduced extracellular Na+ was required to reduce the magnitude of sodium currents to improve the fidelity of the voltage clamp (33Kral M.G. Xiong Z. Study R.E. Pain. 1999; 81: 15-24Abstract Full Text Full Text PDF PubMed Scopus (70) Google Scholar). Under these recording conditions, the estimated Nernst reversal potential for Na+ was +33 mV. In some experiments, action potentials were recorded in the current clamp mode using methods described previously (34Renganathan M. Cummins T.R. Waxman S.G. J. Neurophysiol. 2001; 86: 629-640Crossref PubMed Scopus (428) Google Scholar). TTX-R sodium currents were isolated from TTX-S currents by adding TTX (200 nm, Sigma) to the bath solution. The TTX stock solution (20 mm) was prepared with distilled water and stored at –20 °C. Working TTX solutions were made with extracellular solution on the day of experiments. After formation of a gigaohm seal (1–5 GΩ) and compensation of pipette capacitance with amplifier circuitry, whole cell access was established. The pipette potential was zeroed before seal formation. Membrane resistance, series resistance, and capacitance were determined from current transients elicited by 5-mV depolarizing steps from a holding potential of –60 mV, via the membrane test application of pClamp 8.2. Series resistance was compensated 75–85% as necessary, and recordings were conducted only when access resistance was lower than 10 mΩ. During recordings, the cells were held at –80 mV, and the junction potentials (about 4–5 mV) were not corrected. To access the changes in the current-voltage (I-V) and conductance-voltage (G-V) relationships, data were collected for an I-V curve 5 min after cell rupture. To assess changes in the steady-state inactivation of TTX-R sodium currents, H-infinity curves were collected after obtaining the I-V curve as described previously (21Hirade M. Yasuda H. Omatsu-Kanbe M. Kikkawa R. Kitasato H. Neuroscience. 1999; 90: 933-939Crossref PubMed Scopus (66) Google Scholar). TTX-R sodium currents were evoked from a holding potential of –80 mV to 0 mV every 30 s. Recording data were acquired using a Digidata 1322A 16-bit data acquisition system (Axon Instruments), digitized at 10 kHz, low-pass filtered at 5 kHz, and stored on a computer that was controlled by pClamp software (v8.2, Axon Instruments). Data Analysis—Activation and steady-state inactivation data were fitted with a Boltzmann equation of the form: G = Gmax/(1 + exp(V50–Vm)/k). Here G equals to I/(Vm-Vres), where Vm is the potential at which current is evoked, and Vres is the reversal potential for the current determined by extrapolating the linear portion of the I-V curve through 0 current, Gmax = the calculated maximal conductance, V50 = the potential of half activation or inactivation, and k = the slope factor. All data were expressed as means ± S.E. Statistical analyses were performed using the Student's t test. Immunofluorescence Labeling of Peripherin and Sodium Channels—DRG sections were prepared and stained according to the method described (35Amaya F. Decosterd I. Samad T.A. Plumpton C. Tate S. Mannion R.J. Costigan M. Woolf C.J. Mol. Cell Neurosci. 2000; 15: 331-342Crossref PubMed Scopus (244) Google Scholar). Briefly, the animals were anesthetized with halothane and transcardially perfused with ice-cold saline solution. DRGs from thoracic and lumbar regions of the spinal column were quickly removed, postfixed for 2–3 h in 4% paraformaldehyde in 0.1 m phosphate buffer (PB), and cryoprotected in 10% sucrose in 0.1 m PB for 24 h at 4 °C. Transverse sections through the DRG (10 μm) were cut on a cryostat and mounted serially onto Superfrost/plus microscope slides (Fisher Scientific). For immunofluorescence labeling of sodium channels, the sections were first washed with 0.1 m PB, permeabilized with 0.3% Triton X-100 in 0.1 m PB (PBST) for 2 h at room temperature, and blocked with 10% normal goat serum in PBST (PBSTG) for at least 4 h. The sections then were incubated with anti-sodium channel antibodies and monoclonal anti-peripherin (1:150, Chemicon) antibody in PBSTG for 24 h at 4 °C. Antibodies for sodium channels used were anti-rabbit Nav1.6 (1:200, Sigma), anti-rabbit Nav1.7 (1:100, Sigma), or anti-rabbit Nav1.8 (1:100) from Dr. S. R. Levinson. After three washes with PBST, the sections were incubated with secondary antibodies Alexa Fluor 488 (goat anti-mouse IgG) and Alexa Fluor 594 (goat anti-rabbit IgG) from Molecular Probes (Eugene, OR) for 2 h at room temperature. The sections were then washed, mounted with anti-fade fluorescence mounting medium and stored at 4 °C. All images were captured with a Zeiss Axioplan microscope with a CCD digital camera and processed with Adobe Photoshop 7. Immunoprecipitation and Western Blotting—DRGs in the lumbar and thoracic regions of the spinal column from diabetic and control rats were removed and homogenized in ice-cold lysis buffer containing 50 mm Tris, pH 8.0, 150 mm NaCl, 1 mm EGTA, 50 mm NaF, 1.5 mm MgCl2, 10% v/v glycerol, 1% v/v Triton X-100, 1 mm phenylmethylsulfonyl fluoride, 1 mm NaVO4, and “Complete” protease inhibitor mixture (Roche Diagnostics, Mannheim, Germany). Aliquots of DRG homogenates containing equal amounts of total protein were mixed with the anti-phosphoserine (Chemicon), anti-phosphothreonine (Chemicon), or anti-phosphotyrosine (Sigma) specific antibodies at a 1:40 dilution. The mixtures were incubated and rotated in Eppendorf tubes in the presence of 50 mm NaF and protease inhibitors for 4–14 h at 4 °C. Protein G-agarose beads (Roche Diagnostics) were then added to the samples and incubated overnight at 4 °C. After washing three times with ice-cold lysis buffer, the protein G beads were pelleted and mixed with 2× SDS sample buffer. Proteins were separated on 4–15% gradient Tris-HCl gels and transblotted onto polyvinylidene difluoride membranes (Amersham Biosciences). In some experiments, crude DRG homogenates were loaded on gels for Western blot analysis. The membrane blots were blocked with 10% nonfat dry milk for 12 h and incubated with primary antibodies: anti-Nav1.3 (1:200, Sigma), anti-Nav1.6 (1: 200), anti-Nav1.7 (1:100), or anti-Nav1.8 (1:100) overnight at 4 °C. The membranes were then incubated with horseradish peroxidase-conjugated goat anti-rabbit secondary antibody (Amersham Biosciences) for 2 h at room temperature and developed using the Supersignal West Pico chemiluminescence kit (Pierce). The corresponding bands were scanned at 1200 dpi and semiquantified with Image Quanti-Scan software (Molecular Dynamics, Sunnyvale, CA). Short Term Diabetic Rats Develop Mechanical Allodynia and Thermal Hyperalgesia—Two days after injection of STZ, 70% of the rats developed high levels of blood glucose (mean level = 460 ± 22 mg/dl), whereas untreated rats had normal levels (mean level = 85 ± 6 mg/dl). The elevated level of blood glucose in STZ-injected rats was maintained during the entire experimental period. These results are very similar to our previous studies (32Hall K.E. Liu J. Sima A.A. Wiley J.W. J. Neurophysiol. 2001; 86: 760-770Crossref PubMed Scopus (82) Google Scholar). In the present study, mechanical allodynia was determined by measuring the paw withdrawal threshold in response to application of a von Frey probe (Fig. 1A). Diabetic rats showed a significant decrease in the pressure required to elicit paw withdrawal as compared with their pre-diabetic baseline responses (p < 0.05, n = 6). The paw pressure withdrawal threshold for diabetic rats began to decrease 1 week following STZ-injection, but this change in responsiveness to mechanical stimuli did not reach significance until 4 weeks after the induction of diabetes. Thermal hyperalgesia was determined by measuring the withdrawal latency to a radiant heat stimulus applied to the hind foot. Fig. 1B illustrates the decrease in the withdrawal latency to thermal stimulation in hind paws of diabetic rats. As compared with pre-diabetic baseline responses, diabetic rats began to exhibit a significant reduction in the temperature required to elicit a hind paw withdrawal 4 weeks after STZ injection (p < 0.01, n = 6). Behavioral signs of both mechanical allodynia and thermal hyperalgesia persisted for up to 8 weeks after the onset of diabetes, the maximum duration of our observations. We found no significant differences between baseline and post-injection responses to either mechanical or thermal stimuli of similarly tested, age matched, non-diabetic controls injected with saline instead of STZ. Diabetic Neuropathy Is Associated with Increased Amplitude and Altered Properties of TTX-R Sodium Current—To record TTX-R sodium currents (INa), small-sized DRG neurons (soma diameter < 25 μm, cell capacitance < 35 pF) were isolated and voltage-clamped at –80 mV in the presence of 200 nm TTX in the bath solution. The average resting membrane potential was –60.3 ± 0.3 mV in DRG neurons from control rats and –52.7 ± 0.7 mV in diabetic rats (n = 110 for each group). The difference between these two groups was highly significant (p < 0.0001). The average whole cell capacitance was 22.3 ± 1.2 pF in the control group and 23.8 ± 2.1 pF in the diabetic group. Fig. 2A shows original current traces of TTX-R INa in DRG neurons prepared from a diabetic rat 6 weeks after the onset of diabetes compared with an age-matched control. The amplitude of outward INa was significantly larger in neurons from diabetic rats compared with controls (Fig. 2B). The mean peak current density in diabetic neurons was 39.2 ± 1.5 pA/pF (n = 12) compared with 27.8 ± 2.3 pA/pF in control neurons (n = 15) and the difference between these two values was significant (p < 0.01). As shown in Fig. 2C, the current-voltage relationship calculated for DRG neurons isolated from diabetic rats was shifted ∼8 mV in the hyperpolarizing direction compared with that calculated for controls. Peak current density was elicited by depolarizing control neurons to +0 mV or by depolarizing diabetic neurons to –10 mV. The kinetics of activation was determined by curve fitting the rising phase of the conductance using a single exponential function Boltzmann equation (Fig. 2D). The midpoint of the voltage-dependence of activation was –14.8 ± 0.4 mV in cells from diabetic rats. This value was significantly more negative than the value obtained for control neurons (–8.5 ± 0.3 mV, p < 0.001). The voltage dependence of steady-state inactivation was best fit to a modified Boltzmann function: (INa)/(INa)max = [1 + exp(V–Vh)/k]–1, where Vh is the midpoint of steady-state inactivation and k is the slope factor. The midpoints of steady-state inactivation were –29.1 ± 0.3 mV in neurons from diabetic rats and –23.8 ± 0.3 mV in control neurons. The difference between these values was significant (p < 0.05). Thus, the steady-state inactivation of TTX-R currents was also negatively shifted in small-sized DRG neurons from diabetic rats compared with controls. DRG Neurons from Diabetic Rats Demonstrate Increases in TTX-S and Total Sodium Currents—To elicit total INa, small-sized DRG neurons were depolarized by a pre-pulse to –120 mV for 50 ms and then stepped to potentials ranging from –50 mV to +50 mV in 5-mV increments every 10 s. Fig. 3A demonstrates the current-voltage relationships for the total INa of DRG neurons isolated from control and diabetic rats. The peak current density of total INa was –59.7 ± 8.0 mV in cells from control rats and –73.4 ± 5.0 mV in diabetic rats (p < 0.05, n = 11 for each cell type). Maximal currents for diabetic neurons were measured at –10 mV compared with –5 mV for control neurons. We also observed increased action potential amplitudes in DRG neurons isolated from diabetic rats compared with controls. In current clamp whole cell configuration, the membrane potential was not adjusted, and the action potential was elicited by delivery of 1.5 nA current for 0.5 ms to the patched cell through the amplifier, leaving
Referência(s)