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Originally published In Press as doi:10.1074/jbc.M404167200 on May 3, 2004

J. Biol. Chem., Vol. 279, Issue 28, 29341-29350, July 9, 2004
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Early Painful Diabetic Neuropathy Is Associated with Differential Changes in Tetrodotoxin-sensitive and -resistant Sodium Channels in Dorsal Root Ganglion Neurons in the Rat*

Shuangsong Hong{ddagger}, Thomas J. Morrow§, Pamela E. Paulson§, Lori L. Isom¶, and John W. Wiley{ddagger}||

From the Departments of {ddagger}Internal Medicine and Pharmacology, University of Michigan, Ann Arbor, Michigan 48109 and the §Department of Neurology, Veterans Affairs Medical Center, Ann Arbor, Michigan 48109

Received for publication, April 14, 2004


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
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.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Diabetes mellitus is one of the most common chronic medical problems, affecting over 100 million people world-wide (1). 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 (2–4). The streptozotocin (STZ)1-induced diabetic rat model demonstrates many of the abnormalities observed in humans (5). Treatment with insulin prevents development or reverses many of the abnormalities observed in early painful diabetic neuropathy (6, 7).

In diabetic rats with hyperalgesia, dorsal root ganglion (DRG) neurons display increased frequency of action potential generation in response to sustained suprathreshold mechanical stimulation (3, 4, 8–10) and increased spontaneous activity (11). 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 (12–14). At least two TTX-S sodium channel {alpha}-subunits, Nav1.6, and Nav1.7, are constitutively expressed in the peripheral nervous system (15). 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 (16). Two TTX-R sodium channels, Nav1.8 (17) and Nav1.9 (18, 19) have been identified in DRG neurons and changes in their expression levels have been implicated in painful diabetic neuropathy (20–23). TTX-S sodium channels in brain are composed of a pore-forming {alpha}-subunit and one or two auxiliary {beta}-subunits (24–27). The subunit composition of TTX-R sodium channels, however, is not clear. Increased TTX-R sodium current (21), but decreased expression levels of Nav1.8 mRNA and protein have been reported in models of diabetic neuropathy (28). 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.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
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 (6, 7).

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 (30). 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 x 20 cm x 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 (31).

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 (32). 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 M{Omega}) 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 (33). 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 (34). 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 G{Omega}) 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 M{Omega}. 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 (21). 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 (35). 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 2x 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).


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
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 (32). 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.



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  		FIG. 1.
Line graphs show the development of mechanical allodynia (A) and thermal hyperalgesia (B) following the onset of diabetes in STZ-treated rats as compared with saline treated controls. Note that significant differences in pain-related behaviors in response to both mechanical (Von Frey test) and thermal (Hargreaves test) stimuli are not significant until at least 4 weeks after the administration of STZ, a time when peripheral signs of neuropathy are clearly evident. Data shown are mean values ± S.E. of 6 rats tested each group. The significance (p < 0.05) are indicated as asterisks (*).

 
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).



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  		FIG. 2.
Changes in amplitude and properties of TTX-R INa in small-sized DRG neurons from diabetic rats. A, typical traces of TTX-R INa conducted in voltage-clamp mode in control (top) and diabetic (bottom) rats. Cells were voltage-clamped at –80 mV, and currents were elicited by stepping voltage from –50 mV in the presence of 200 nM TTX. B, single traces of TTX-R INa elicited by depolarizing cells to 0 mV from the holding potential in neurons from control and diabetic rats. C, line graph showing the current-voltage relationship using current density as the indicator for INa in DRG neurons from control and diabetic rats. This I-V curve shifted leftward in DRG cells from diabetic animals. D, normalized Na+ conductance-voltage curves obtained after 6 min of cell rapture. Conductance was calculated according to the equation G = I/(V–Vres), where V is the test potential and Vres is the reversal potential of sodium. Data are fit with a single Boltzmann equation: GNa/(GNa)max = 1/(1 + exp(E50–E)/),k where E50 is the potential (E) at which G is half of Gmax. E, normalized current-voltage curves of steady-state inactivation. Inlet shows typical traces of INa recorded after a long pre-pulse to –100 mV (1000 ms) followed by stepping voltage from –90 mV to +10 mV in 5 mV increments. Data are fit with a single Boltzmann equation. Error bars indicate S.E. {circ}, control; , diabetic.

 
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 most of the action potential free of the effect of injected current (36). The majority of the action potentials recorded displayed a characteristic shoulder on the falling phase, indicating the activation of nociceptive neurons corresponding to C-type fibers, as previously described (37, 38). The mean amplitude of the action potential was significantly larger in diabetic neurons (130.3 ± 1.3 mV) compared with controls (115.3 ± 0.9 mV) (p < 0.05, n = 8 for each cell type). The 10–90% rise time of the action potential was significantly shorter in diabetic neurons (0.27 ± 0.01 ms) than in control cells (0.58 ± 0.05 ms), as demonstrated in Fig. 3B (p < 0.001, n = 9 for each cell type). These results suggest that diabetic DRG neurons are more sensitive to action potential stimulation than control cells under the same conditions.



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  		FIG. 3.
Enhanced total INa in small-size DRG neurons from diabetic rats. A, increased current density of total INa in small-size DRGs from diabetic rats compared with the control. The difference in the maximum of peak current density was significant between control and diabetic rats (p < 0.05). B, original action potential traces recorded in current-clamp mode without adjusting the membrane potential after cell rupture. The average amplitude of action potential was significantly larger in diabetic (130.3 ± 1.3 mV) than that in control (115.3 ± 0.9 mV) neurons (p < 0.05). Error bars indicate S.E.

 
Fig. 4A shows original traces of total and TTX-R INa evoked by voltage step to 0 mV. The TTX-S current was obtained by digitally subtracting the currents recorded after applying 200 nM TTX from the currents recorded in the absence of TTX. The rising phase of the TTX-S current precedes the peak of TTX-R current, suggesting that the activation of TTX-S current is more rapid. The current-voltage relationship for TTX-S INa in small-sized DRG neurons from control and diabetic rats is shown in Fig. 4B. The peak current density for TTX-S INa was –35.1 ± 3.1 mV in cells from diabetic rats, which is significantly more negative than that for control neurons (–25.5 ± 2.7 mV, p < 0.05).



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  		FIG. 4.
Increased amplitude and altered properties of TTX-S INa in small-sized DRG neurons from diabetic rats. A, typical recording traces showing the isolation of TTX-S sodium current (dotted trace) from total sodium current and TTX-R current. Cells were clamped at –80 mV and depolarized to 0 mV after a pre-pulse to –120 mV for 50 ms to obtain total INa as indicated. TTX of 200 nM was delivered by puffer on top of the cell, and TTX-R current trace was recorded. TTX-S current trace was obtained by subtraction of TTX-R INa trace from total INa trace. B, the maximum current density of TTX-S INa enhanced about 40% in DRG neurons from diabetic rats when compared with the control. The I-V curve shifted leftward around 5 mV in DRG neurons of diabetic rats. C, conductance-voltage relationship of TTX-S INa in DRG neurons from control and diabetic rats. The half-activation voltage was –16.1 ± 0.1 mV for TTX-S INa in diabetics and –12.8 ± 0.4 mV in controls, respectively. The difference was significant (p < 0.05). The slope factor was also significant larger in neurons from diabetic rats (5.4 ± 0.3) than that from control rats (2.7 ± 0.1). Error bars indicate S.E.

 
The kinetics of activation of TTX-S INa were calculated and fit with a single Boltzmann equation as shown in Fig. 4C. The midpoint of voltage-dependent activation for TTX-S INa was –12.8 ± 0.4 mV in cells from control rats and –16.1 ± 0.1 mV in cells from diabetic rats. The difference between these values was significant (p < 0.05). The slope factors for the TTX-S INa activation curves were of 2.7 ± 0.1 for diabetic neurons and 5.4 ± 0.3 for control neurons, and the difference between these values was significant (n = 8; p < 0.01).

Increased Slow and Fast Ramp Currents in DRG Neurons from Diabetic Rats—Because threshold ramp currents might influence the excitability of DRG neurons, we also examined sodium currents in small-sized DRG neurons elicited in response to a slow voltage ramp (~0.23 mV/ms) depolarization from –120 mV to +40 mV over a 695 ms time course. To increase INa to more easily measurable levels the Na+ concentration in the external recording solution was increased to from 35 to 120 mM. Sodium currents elicited by this protocol were larger in neurons from diabetic rats than in controls (Fig. 5A). When the peak ramp currents were normalized to cell capacitance, the mean current density was ~130% larger in neurons from diabetic rats compared with the controls (Fig. 5C). In diabetic neurons, the ramp current density was –21.6 ± 3.0 pA/pF whereas it was –6.5 ± 1.3 pA/pF in control neurons (p < 0.01, n = 13 for each cell type).



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  		FIG. 5.
Comparison of ramp currents in small-sized DRG neurons from control and diabetic rats. A, slow ramp currents elicited in DRG neurons by 695 ms voltage ramp extending from –120 to +40 mV (~0.23 mV/ms) from control and diabetic rats. B, fast ramp current traces examined by 69.5-ms voltage ramp from –120 to +40 mV (~2.3 mV/ms) in DRG neurons from control and diabetic rats. C, bar graph depicting the ramp current density (estimated by dividing the peak currents by the cell capacitance) evoked by fast and slow voltage ramps. Both current densities of fast and slow ramps were significantly larger in diabetic rats than controls (n = 13). Error bars indicate S.E., and the * indicates p < 0.05. D, original traces of resurgent current in DRG neurons from control and diabetic rats. The resurgent current was examined by a high depolarization to +30 mV followed by hyperpolarizing to –40 mV. The large DRG neuron (cell capacitance = 106) shows a typical slow inactivating current when hyperpolarized to –40 mV after a deep depolarization.

 
To elicit the fast ramp current, DRG neurons were depolarized to +40 mV from –120 mV at a rate of 2.3 mV/ms. As shown in Fig. 5B, the fast ramp current elicited in diabetic neurons was larger than the control. The mean peak current density elicited by fast voltage ramp was –24.9 ± 5.3 pA/pF in controls and –42.6 ± 5.9 pA/pF in diabetic neurons (Fig. 5C). We also examined the levels of resurgent current in diabetic versus control neurons (39). After a conditioning step to +30 mV for 50 ms to promote inactivation, hyperpolarizing pulses ranging from –50 to 0 mV were used to assay resurgent current. Fig. 5D shows the original traces recorded in neurons depolarized to +30 mV followed by a hyperpolarizing pulse to –40 mV. Large sized DRG neurons elicited a resurgent current using this protocol; however, there were no significant differences in this current observed between diabetic and control neurons. Resurgent currents were barely measurable in small sized DRG neurons from either control or diabetic rats.

Diabetic Neuropathy Is Associated with Differential Changes in Sodium Channel Expression in DRG Neurons—To test whether the observed increases in TTX-S and TTX-R INa in diabetic neurons could be due to altered sodium channel expression, we extracted crude homogenates of DRGs and analyzed the expression of sodium channels by Western blot. Our results show that the expression levels of TTX-S and TTX-R sodium channel {alpha}-subunits were differentially affected in DRGs of diabetic rats (Fig. 6). In agreement with previously published results (28), we observed that the expression of Nav1.8 significantly decreased (40% less) in diabetic DRGs compared with control (p < 0.05, n = 5). In contrast to previous studies (28), we found that the expression of Nav1.6 also decreased in diabetic DRGs (38%, n = 5, Fig. 6, A and B). The expression levels of Nav1.3 and Nav1.7 increased significantly in DRGs from diabetic rats compared with controls (p < 0.05, n = 4 for each group). These increases were ~82 and 45% Nav1.3 and Nav1.7, respectively, in DRGs from diabetic rats as compared with the controls (Fig. 6B).



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  		FIG. 6.
Western blot analysis of sodium channel {alpha}-subunits in DRG extracts from control and diabetic rats. A, expression of the {alpha}-subunits of sodium channels shown by Western blot using crude DRG membrane homogenates and polyclonal antibodies directly against the TTX-S (Nav1.3, Nav1.6, Nav1.7) and TTX-R (Nav1.8) {alpha}-subunits of sodium channels. These antibodies labeled predominantly a high molecular mass protein (>200 kDa). The transblots were representatives of 4–5 independent analyses. B, histogram depicting relative intensities of Western blot analysis corresponding to the sodium channel {alpha}-subunits. The protein expression of Nav1.3 and Nav 1.7 were found to be significantly increased (p < 0.05, Student's t test) in diabetic rats compared with controls (n = 4, each group), whereas the total protein of Nav 1.6 (TTX-S) and Nav1.8 (TTX-R) decreased significantly in DRG neurons of diabetic rats compared with the control (p < 0.05, n = 5). CT, control; DM, diabetic. Error bars indicate S.E., and the * indicates p < 0.05.

 
Immunolocalization of Sodium Channels in DRG Neurons—As shown in the Fig. 7A, we observed that Nav1.6 was expressed primarily in small and medium sized DRG neurons in both control and diabetic rats. We found that Nav1.6 immunoreactivity was similar in control and in diabetic DRGs. As expected, the 38% decrease in total Nav1.6 protein expression observed in Fig. 6 was not detectable using immunofluorescence. Of the total DRG neurons, 63.6 ± 1.9% were immunoreactive for Nav1.6 in control rats and 59.7 ± 4.7% showed immunoreactivity in diabetic rats (Table I). This difference was not significant (p > 0.05). The distribution of Nav1.6 expression among small and large diameter neurons also did not appear to change in diabetes. For example, 44.7 ± 1.1% versus 44.8 ± 5.1% of the neurons that were Nav1.6-positive were also positive for peripherin (a marker of unmyelinated C-fibers) (35, 41) in control and diabetic DRGs, respectively.



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  		FIG. 7.
Double immunofluoresence staining of DRG neurons with C-type fiber marker peripherin (green) and sodium channel (red) antibodies. The neurons of double immunoreactivity positive are shown in yellow. Left panel, control; right panel, diabetic. A, decreased expression of TTX-S Nav1.6 in DRG neurons from diabetic rats (right panel) compared with the control (left panel). B, increased protein expression TTX-S Na 1.7 was detected in DRG neurons from vdiabetic (right panel) rats compared with the control (left panel). Both the percentage of Nav1.7-positive and the percentage of neurons double positive for Nav 1.7 and peripherin were increased significantly for DRGs from diabetic rats (p < 0.05). C, protein expression of TTX-R Nav 1.8 decreased in DRG neurons from diabetic rats. The percentage of DRG neurons positive for both peripherin (green) and Nav1.8 (red) decreased in DRG neurons from diabetic rats. Scale bar, 50 µm.

 


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  		TABLE I
The proportion of neurons that are immunoreactive-positive for sodium channel {alpha}-subunits and peripherin in DRGs from control and diabetic rats

 
Nav1.7 is highly expressed in small DRG neurons (42, 43), has slowly repriming kinetics, and a slow onset of closedstate inactivation (44). In agreement with our Western blot results (Fig. 6), we found that the level of Nav1.7 immunoreactivity increased in small and medium sized DRG neurons from diabetic rats compared with controls (Fig. 7B). 55.6 ± 2.4% of total DRG neurons were immunoreactive for Nav1.7 in control rats compared with 65.6 ± 1.1% in diabetic rats (Table I), representing an increase of ~18% (p < 0.01). The percentage of the neurons that stained positively for both Nav1.7 and peripherin was 24.1 ± 1.3% in control rats and 28.2 ± 1.4% in diabetic rats. While this difference was significant (p < 0.05) (Table I), it also indicates that the greatest increase in Nav1.7 expression was in non-C-fiber neurons. These data are the first demonstration of changes in Nav1.7 expression in painful diabetic neuropathy.

Nav1.8 is expressed primarily in small- and medium-sized DRGs and has been implicated in the transmission of neuropathic pain (45, 46). As demonstrated in Fig. 7C, the level of Nav1.8 staining decreased in DRGs from diabetic rats compared with controls, in agreement with our Western blot results (Fig. 6) and with the results of previous studies (28). The majority of peripherin-positive neurons also expressed Nav1.8 in control animals. Based on the total neurons counted (Table I), ~77.6 ± 3.0% of control neurons were immunopositive for Nav1.8 in control rats. This value dropped significantly (20%), to 62.6 ± 2.1% in diabetic rats (p < 0.01). Neurons that stained positively for both Nav1.8 and peripherin were 37.2 ± 4.3% of the total in control rats and 29.5 ± 0.4% of the total in diabetic rats (p > 0.05) (Table I). Thus, both the overall levels of Nav1.8 protein expression and the number of neurons expressing Nav1.8 decreased with the onset of diabetic neuropathy despite increased TTX-R current.

Enhanced Phosphorylation of Both TTX-S and TTX-R Sodium Channels in DRG Neurons from Diabetic Rats—Phosphorylation of TTX-R sodium channels results in increased sodium current and this may play a role in nociceptive processing (26, 45, 47). To test whether nociceptive signaling in diabetic neuropathy also involves changes in sodium channel phosphorylation, we immunoprecipitated solubilized DRG homogenates with anti-phosphoserine-, anti-phosphothreonine-, or anti-phosphotyrosine-specific antibodies, and then analyzed the immunoprecipitated proteins by Western blot using anti-sodium channel antibodies. We also performed reciprocal immunoprecipitation experiments using the antibodies in the reverse order. The results of both sets of experiments were indistinguishable and thus the data were pooled. As demonstrated in Fig. 8A, an increased level of anti-phosphoserine immunoreactivity (~60%) for Nav1.8 protein was measured in DRG homogenates from diabetic rats compared with controls. Nav1.8 anti-phosphothreonine immunoreactivity increased ~82% in DRGs from diabetic rats compared with the controls (n = 4, p < 0.01). No evidence for tyrosine phosphorylation of Nav1.8 was observed in DRGs from either control or diabetic rats. In contrast, anti-phosphotyrosine immunoreactivity was observed for Nav1.6 (Fig. 8B) and Nav1.7 (Fig. 8C), and this apparent tyrosine phosphorylation appeared to be increased in diabetic rats. Threonine-phosphorylated forms of Nav1.6 and Nav1.7 were measured and the level of anti-phosphothreonine immunoreactivity increased ~80% for both isoforms in diabetic rats compared with controls (n = 5 for both groups, p < 0.05). In addition, an increased level of serine phosphorylation of Nav1.6 was observed in diabetic rats (Fig. 8B). Very low levels of serine-phosphorylated Nav1.7 were observed in DRGs from control rats and a clear corresponding band was detected in DRGs from diabetic rats (Fig. 8C). Overall, it appears that phosphorylation of Nav1.6, Nav1.7, and Nav1.8 is increased in DRGs of diabetic rats, suggesting that changes in serine/threonine and tyrosine phosphorylation of voltage-gated sodium channels may play a role in transmission of painful stimuli in diabetic neuropathy.



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  		FIG. 8.
Phosphorylation states of sodium channels in DRG neurons from control and diabetic rats. A, increased phosphorylation of Nav1.8 in serine residue (p-Ser) and threonine residue (p-Thr) in diabetic rats. Crude DRG homogenates were used for immunoprecipitation followed by Western blot analysis. A high molecular mass band of protein (>200 kDa) was labeled in immunoblots (left panel), and the average of pixel intensity corresponding to the labeled band was shown in the bar graph (right panel). B, increased phosphorylation of Nav 1.6 in serine, threonine, and tyrosine (p-Tyr) residues in diabetic rats. C, increased phosphorylation of Nav1.6 in serine, threonine, and tyrosine residues in diabetic rats. CT, control; DM, diabetic.

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
Increased TTX-R sodium currents have been implicated in chronic pain, including diabetic neuropathy. We believe that our results represent one of the first systematic analyses of TTX-S and TTX-R sodium channel expression and function, including phosphorylation status, in diabetic neuropathy. Our data demonstrate that both TTX-S and TTX-R sodium currents increased in small (nociceptive) DRG neurons from diabetic rats and that these increases in current correlated with the development of hyperalgesia and allodynia in early diabetes. The expression levels of TTX-S and TTX-R channel subtypes were differentially affected in diabetic rats. Expression of Nav1.8 decreased significantly as assessed by both Western blot analysis of DRG homogenates and immunocytochemical analysis of peripherin-positive DRG neurons. Our data suggest that the development of diabetes in this model does not involve alterations in the percentage of C-type neurons that express Nav1.8, but does involve decreases in the level of Nav1.8 expression in individual C-type neurons compared with healthy controls. In contrast to Nav1.8 expression, we observed a robust increase in the expression of Nav1.7 that paralleled the development of hyperalgesia and allodynia. We observed an increase in the number of Nav1.7-positive neurons in the DRG, an increase in the strength of the Nav1.7 immune signal in individual neurons, and an increase in Nav1.7 protein expression in Western blot analysis of DRG homogenates. These results support our hypothesis that both TTX-S and TTX-R sodium channels play important roles in the development of early painful diabetic neuropathy.

We observed a significant increase in TTX-R sodium current, decreased protein expression, and increased phosphorylation of Nav1.8 in DRG neurons isolated from diabetic rats. These data, together with the results from other groups (21, 28), support a role for Nav1.8 channel in painful diabetic neuropathy. The level of serine/threonine phosphorylation of Nav1.8 increased at DRGs from diabetic rats, strengthening the evidence that phosphorylation of sodium channels is involved in modifying channel properties in pain-related animal models and showing for the first time that sodium channel phosphorylation contributes to painful diabetic neuropathy. The observed phosphorylation in our diabetic model may be the cause of the increase in TTX-R current observed in diabetic rats since cAMP-dependent phosphorylation of Nav1.8 increased TTX-R currents in both oocytes and transfected COS-7 cells (48, 49). We propose that phosphorylation of Nav1.8, possibly at sites on the DI-DII linker domain, alters the voltage-dependent activation and inactivation or open probability of the channel but not the incorporation of additional channels into the plasma membrane, and consequently increases the TTX-R current. In our experiments, based on the inactivation phase of the TTX-R current, 80–90% of the TTX-R current observed in DRG neurons can be attributed to Nav1.8 because Nav1.9 shows much slower inactivation kinetics compared with Nav1.8. The role of Nav1.9 remains to be examined in diabetic neuropathy although a recent study shows that knockdown of this channel produced no effects on thermal hyperalgesia or tactile hypersensitivity in the neuropathic rat (50).

Other studies support the interpretation that TTX-R channels are not the only subtypes that are involved in painful diabetic neuropathy. A recent study reports that TTX application to injured nerve only partially reverses neuropathic pain (51). In Nav1.8-null mutant mice, nerve injury elicits thermal hyperalgesia and tactile hypersensitivity after day 3, suggesting that neuropathic pain is developed and maintained despite the absence of Nav1.8 (46, 52). Taken together, these results suggest that TTX-S channels also contribute to neuropathic pain. We observed that TTX-S current contributes 40–50% of the total current in small DRG neurons from both control and diabetic rats. In diabetic rats, the TTX-S currents increased significantly and the midpoint voltage of activation shifted modestly in the hyperpolarizing direction. Since the contribution of TTX-S currents to the threshold and upstroke of nociceptor action potentials is likely to be sensitive to small changes in the resting potential (36), the increased amplitude and negative shift of the activation of TTX-S current (see Fig. 4) could result in a larger contribution to the electrogenesis of action potentials in cells with a depolarizing resting potential near the –52 mV observed in diabetic rats. The faster activation kinetics of TTX-S currents in diabetic rats are also predicted to allow channels to inactivate more rapidly, thereby maintaining higher excitability.

Among the TTX-S channels, Nav1.7 displays slow activation and inactivation kinetics and slow repriming kinetics (44). In diabetic rats, the increased protein expression of Nav1.7, particularly in peripherin-positive (C-fiber) neurons, may lower the threshold in these neurons (34) that previously expressed low levels of Nav1.7. Consequently, nociceptive C-fiber neurons may generate more robust responses to slowly depolarizing inputs in diabetic rats compared with healthy controls because of the increased slowly inactivating ramp currents (Fig. 7). The increased expression of Nav1.7 in diabetic rats may also increase the duration of the action potential and decrease the conduction velocity (43). Previous studies showed that phosphorylation of Nav1.7 decreased current amplitude in oocytes either via a protein kinase A (PKA)-mediated pathway without altering voltage sensitivity and gating kinetics or via a protein kinase C (PKC)-dependent pathway that is accompanied by a depolarizing shift in the steady-state activation (48). In adrenal chromaffin cells, activation of PKC down-regulated Nav1.7 via either promoting endocytic internalization of Nav1.7 channels or destabilizing the mRNA (55). In contrast to these results, we observed increased Nav1.7 protein, currents and increased phosphorylation of Nav1.7 at Ser/Thrresidues and significantly at Tyr residues in DRGs isolated from diabetic rats. We propose that modulation of Nav1.7 through phosphorylation may affect channel function and expression differently in DRG neurons in vivo compared with adrenal chromaffin cells or oocytes. For example, forskolin decreased Nav1.7 currents at low concentrations but failed to alter Nav1.7 currents at high concentrations, suggesting a biphasic effect of forskolin on channel function (48). Our data also raise the possibility that tyrosine kinase-mediated pathways modulate Nav1.7 in addition to PKA and PKC. It has been reported that tyrosine phosphorylation produced by stimulating receptor tyrosine kinases inhibits sodium channel currents in PC12 cells (54) and dephosphorylation provides opposite regulation of sodium channel function (40). However, this inhibition of tyrosine phosphorylation may be tissue-specific since inhibition of protein tyrosine kinase inhibits fast sodium currents in rabbit cardiac myocytes (53). Therefore, the role of tyrosine phosphorylation in regulating individual sodium channel subtype, i.e. Nav1.7 in diabetic neuropathy, remains to be investigated.

For Nav1.6, our data indicate that it is highly expressed in both small and large DRG neurons (Fig. 7). The overall expression of Nav1.6 channel decreased in homogenates of diabetic DRG neurons but there was no difference in either the percentage of neurons expressing Nav1.6 or in its expression in C-fibers as assessed by immunocytochemistry. We observed phosphorylation of Nav1.6 at Tyr residues and the level of tyrosine phosphorylation increased in DRGs isolated from diabetic rats. Tyrosine phosphorylation of cerebellar Purkinje cell Nav1.6 channels has been implicated in the maintenance of resurgent current (29, 39). However, we did not detect resurgent currents in small DRG neurons from either control or diabetic rats despite this phosphorylation, suggesting that this effect may be cell-specific.

In conclusion, this study demonstrates that the function of TTX-S and TTX-R sodium channels are increased in early diabetic neuropathy and, therefore, may contribute to the pathophysiology of painful diabetic neuropathy. These enhanced sodium currents are correlated with increased phosphorylation at both serine/threonine and tyrosine sites. A new generation of specific inhibitors or blockers for TTX-S sodium channels would help to elucidate the contribution of different subtypes of TTX-S channels to the development of thermal hyperalgesia and mechanical allodynia in painful diabetic neuropathy, setting the foundation for improved targeted therapeutic interventions.


   FOOTNOTES
 
* This work was supported by Grants RO1 DK52387 and DK 45820 from the National Institutes of Health (to J. W. W.) and Grant RG2882 from the National Multiple Sclerosis Society (to L. L. I.). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

|| To whom correspondence should be addressed: Dept. of Internal Medicine, University of Michigan Hospital, A7007 UH, Box 0108, 1500 East Medical Center Dr., Ann Arbor, MI 48109-0108. Tel.: 734-936-8080; Fax: 734-936-4024; E-mail: jwiley{at}med.umich.edu.

1 The 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. Back



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