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Department of Anesthesiology, Yale University School of Medicine, New Haven, Connecticut
Submitted 8 August 2005; accepted in final form 5 November 2005
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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; Kim et al. 1998; Moore et al. 2002; Song et al. 2003a,b; Zhang et al. 1999).
The ionic mechanisms contributing to the hyperexcitability of DRG neurons after peripheral inflammation or neuronal injury have been widely studied, but results have differed between laboratories and models. For instance, after transection of the sciatic nerve, there is an up-regulation of rapidly repriming TTX-sensitive (TTX-S) Na+ channels and a down-regulation of two isoforms of TTX-resistant (TTX-R) Na+ channels (Abdulla and Smith 2002; Black et al. 1999; Dib-Hajj et al. 1999; Sleeper et al. 2000; Waxman et al. 1994). In addition, voltage-gated K+ currents and Ca2+ current are down-regulated as well (Abdulla and Smith 2001b; Everill and Kocsis 1999; Ishikawa et al. 1999; Yang et al. 2004). After inflammation of peripheral tissues by carrageenan injection, there is an up-regulation of mRNA and currents for both TTX-S and TTX-R Na+ channels (Black et al. 2004; Tanaka et al. 1998). In addition, gastric ulcers in rats and ileitis in guinea pigs are associated with a reduction of K+ currents in DRG neurons (Dang et al. 2004; Stewart et al. 2003).
In our laboratory, chronic compression of the DRG (CCD) is used in the rat as a model of the ganglion compression that can occur in humans after, for instance, foraminal or spinal stenosis, disk herniations, degenerative disorders, tumors, or spinal injuries. In this model, a rod is implanted into the lumbar intervertebral foramen, causing a chronic compression of the DRG (Hu and Xing 1998). These rats develop a cutaneous hyperalgesia and tactile allodynia on the ipsilateral foot. Electrophysiologic recordings of DRG neurons from CCD rats evidence an enhanced excitability including a decrease in the current and voltage thresholds required to elicit APs and an increase in the incidence of spontaneous activity (Hu and Xing 1998; Song et al. 2003a,b; Zhang et al. 1999).
The purpose of this study was to study the effects of CCD on both AP properties and voltage-gated Na+ and K+ currents in cutaneous, medium-sized, dissociated DRG neurons. Our findings suggest that a negative shift of activation for voltage-gated Na+ channels, an increase in TTX-R Na+ current, and a decrease in fast-inactivating K+ currents could contribute to neuronal hyperexcitability in these neurons after CCD, including the decreased current and voltage threshold for APs and an increased capability to produce multiple APs during a sustained depolarization.
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METHODS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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Female Sprague-Dawley rats (140–160 g) were anesthetized with pentobarbital sodium (40 mg/kg, ip). A fluorogold solution in distilled water (1%, 0.05 ml; Molecular Probes, Eugene, OR) was injected into the right lateral plantar region at several points (Oyelese and Kocsis 1996; Yao et al. 2003) to retrogradely label the cell bodies of cutaneous afferent neurons.
CCD surgery
Immediately after fluorogold labeling, one group of rats received no surgery and served as unoperated controls. In the other group of rats, the ipsilateral, right transverse process, and intervertebral foramina of L4 and L5 were exposed as described previously (Hu and Xing 1998; Song et al. 1999). A stainless steel L-shaped rod (0.63 mm diam and 4 mm in length) was inserted into each foramen, one at L4 and the other at the L5 ganglion. Another group of rats received no additional surgery and were used as controls. Sham operations (surgery before and without implantation) were not considered necessary because of the absence of behavioral effects of such surgery or electrophysiological differences between neurons from previously unoperated versus sham-operated controls (Zhang et al. 1999).
Cell preparation
Five to 9 days after Flurogold injection, the rats were deeply anesthetized with pentobarbital sodium (40 mg/kg, ip), and the L4 and L5 lumbar DRGs were exposed. In CCD rats, the correct placement of each implanted rod was confirmed. DRGs were removed from control or CCD rats and placed in complete saline solution (CSS) for cleaning and mincing. The CSS contained the following (in mM): 137 NaCl, 5.3 KCl, 1 MgCl2, 3 CaCl2, 25 sorbitol, and 10 HEPES. The connective tissue of the DRGs was digested for 25 min with collagenase A (1 mg/ml; Boehringer Mannheim, Mannheim, Germany) and for another 25 min with collagenase D (1 mg/ml; Boehringer Mannheim) and papain (30 U/ml; Worthington, Lakewood, NJ) in CSS containing 0.5 mM EDTA and 2 µg of cysteine at 37°C as described previously (Rizzo et al. 1995; Yao et al. 2003). The cells were dissociated by trituration in culture medium containing 1 mg/ml bovine serum albumin and 1 mg/ml trypsin inhibitor (Boehringer Mannheim) and plated on glass coverslips coated with polyornithine and laminin. The culture medium contained equal amounts of DMEM (Invitrogen, San Diego, CA) and F-12 (Invitrogen) with 10% fetal calf serum (HyClone, Logan, UT) and 1% penicillin and streptomycin (Invitrogen). The cells were incubated at 37°C (5% CO2 balance air) for 1 h, after which culture medium without the inhibitor was added. Cells were cultured for 14–24 h before recording.
Electrophysiological recording
Coverslips were transferred to a recording chamber that was mounted on the stage of an upright microscope (BX50-WI, Olympus Optical, Tokyo. Japan). Neurons selected for recording were 35–45 µm in diameter and were positive for Fluorogold fluorescence. Patch pipettes (0.8–2 M
) were fabricated from borosilicate glass (World Precision Instruments, Sarasota, FL). Neurons were recorded at room temperature in whole cell mode (Multiclamp 700A, Axon Instruments, Union City, CA) using pClamp 8.01 software (Axon Instruments). Data were filtered at 3 kHz and digitized at 10–50 kHz. The voltage drop across the access resistance was compensated at 50–80%. The chamber was filled with normal bath solution. Chemicals and other bath solutions were applied locally through a perfusion pipette (100 µm diam) using a drug application system (ALA Scientific Instruments, Long Island, NY). All chemicals were purchased from Sigma (St. Louis, MO) unless otherwise indicated.
Three stimulus protocols were used to compare the excitability of DRG neurons from CCD and control. A series of 400-ms depolarizing currents (0.1–2.5 nA in increments of 0.05–0.1 nA) were injected to induce one or more APs. The AP rheobase was defined as the minimum current required evoking an AP. Repetitive discharge of each cell was measured by counting the spikes evoked by injection of a 1,000-ms depolarizing current at 2.5 times of rheobase (Song et al. 2003a,b). The discharge patterns of the DRG neurons were classified into two types: 1) cells firing either one or two APs and 2) cells firing more than two APs. Other AP properties were measured from the first AP evoked by a series of 2-ms depolarizing currents injected (0–4.0 nA in increments of 0.1 nA). The AP voltage threshold was defined as the first point on the upstroke of an AP where the rising rate exceeded 50 mV/ms (Anderson et al. 1987). The AP amplitude was measured from the baseline to peak of the AP. The AP duration was measured at one-half of the peak amplitude. The afterhyperpolarization (AHP) amplitude was measured from the baseline to the valley peak, and the AHP duration was measured as an interval from the onset of the AHP to the point of 50% decay of the AHP.
Solutions for recording under current clamp
The bath solution contained (in mM) 145 NaCl, 3 KCl, 1 CaCl2, 2 MgCl2, 10 HEPES, and 10 glucose. The pH was adjusted to 7.4 with NaOH, and the osmolarity adjusted to 300
310 mOsm. The pipette solution contained (in mM) 138 KCl, 1 CaCl2, 2 MgCl2, 10 EGTA, and 10 Na-HEPES. The pH was adjusted to 7.2, and the osmolarity was 290300 mOsm.
Solutions for recording sodium currents
Sodium currents were measured under a reduced sodium gradient to reduce space clamp problems. The bath solution contained (in mM) 35 NaCl, 105 TEA Cl, 4 4-aminopyridine (4-AP), 2 BaCl2, 0.1 CaCl2, 0.1 CdCl2, 10 HEPES, and 10 glucose. The pH was adjusted to 7.4 with HEPES, and the osmolarity was adjusted to 300310 mOsm. The pipette solution contained (in mM) 140 CsF, 10 NaCl, 10 EGTA, and 10 HEPES. The pH was adjusted to 7.2 with CsOH, and the osmolarity to 290300 mOsm. To isolate TTX-R Na+ currents, TTX (300 nM) was added to the bath solution.
Solutions for recording potassium current
The K+ bath solution contained (in mM) 140 choline Cl, 3 KCl, 1 MgCl2, 1 CaCl2, 0.1 CdCl2, and 10 HEPES. The pH was adjusted to 7.4 with Tris-base, and the osmolarity was adjusted to 300310 mOsm. The pipette solution contained (in mM) 100 choline Cl, 40 KCl, 1 MgCl2, 10 EGTA, and 10 HEPES. The pH was adjusted to 7.2 with Tris-base, and osmolarity to 290300 mOsm.
Data analysis
Data were analyzed using pClampfit 8.1 (Axon Instruments) and SigmaPlot 8.0 (SPSS, Chicago, IL). Data are expressed as means ± SE. Statistical differences between CCD and control neurons were analyzed by Student's t-test and 
2 tests. In the case of multiple comparisons, for instance, over a voltage range, data were initially analyzed using an ANOVA with repeated measures. If this revealed a significant difference, Student's t-tests were applied for post hoc testing with a Bonferroni correction applied to the probability criterion for significance that was initially set at P < 0.05.
For the analysis of steady-state activation, the peak sodium conductance (gNa) at different potentials was calculated, as a chord conductance, from the corresponding peak current, INa
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where Vg is the voltage for half-maximum activation and kg is the slope factor.
The steady-state inactivation parameter (h
) was calculated at each prepulse potential by dividing the current evoked after a given prepulse potential by the current evoked after the maximal hyperpolarization potential. The inactivation parameter was fitted to a Boltzman function for determination of the half inactivation potential, Vh, and the slope factor
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For the analysis of kinetics, the time constants for activation (
m) and inactivation (h) for TTX-S and TTX-R sodium currents were measured at each potential over the voltage range of –40 to 10 mV.
A two-term exponential function was used to fit the falling phase of 4-AP–sensitive Ka currents. The current amplitude (Ifast and Islow) and time constant (fast and slow) for each cell were determined.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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Fifty-five cutaneous DRG neurons of medium size were recorded by whole cell patch clamp. Thirty-one were from control rats and 24 from CCD. In addition, 26 control and 23 CCD cells were used to compare the capability of repetitive firing induced by the injection of current at 2.5 times rheobase.
The mean rheobase current was significantly smaller for CCD than for control neurons (4.1 ± 0.7 vs. 7.9 ± 1.4 pA/pF; P < 0.05, Student's t-test). In addition, the voltage threshold was significantly more negative for CCD than for control neurons (–33.8 ± 1.9 vs. –26.0 ± 1.9 mV; P < 0.01, Student's t-test; Fig. 1). No significant difference was found between control and CCD group in cell capacitance, resting membrane potential, input resistance, AP amplitude, AP duration, AHP amplitude, or AHP duration (Table 1; Student's t-test). The percentage of cells discharging multiple (>2) APs was significantly higher in CCD than in control neurons (13/23 vs. 5/26; P < 0.01, 2 test; Fig. 1).
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Voltage-gated Na+ current was measured (Fig. 2) first in the absence of TTX to obtain the total Na+ current and then in the presence of TTX to obtain the TTX-resistant (TTX-R) current. By subtracting the latter from the former, the TTX-sensitive (TTX-S) current was obtained. The activating and inactivating kinetics were faster for TTX-S currents than for TTX-R currents (Fig. 2).
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TTX-S currents were activated at approximately –50 mV for CCD and control neurons and reached peak current values near –20 mV (Fig. 3A). Student's t-tests were used to test for differences in current between CCD and control groups. Peak current values were not significantly different between control and CCD neurons (CCD: –136.4 ± 27.1 pA/pF; n = 16; control: –143.3 ± 18.0 pA/pF; n = 18). However, at –40 mV the mean current amplitude was significantly higher for the CCD group (-111.8 ± 24.4 pA/pF) than for the control group (–44.6 ± 15.0 pA/pF). No significant difference in current amplitude was found between CCD and control groups at higher activation voltages.
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Because some neurons expressed little or no TTX-R current, analyses of TTX-R current magnitude/kinetics were only undertaken on cells whose ratio of peak TTX-R to TTX-S currents was >10% at –20 mV. This criterion reduced the population to nine cells from the control group and nine cells from the CCD group. The relative proportion of cells that met the above criteria was not different between control (39.1%) and CCD (42.9%; P > 0.05, 2 test). The mean diameters of these cells were 39.7 ± 1.2 µm for control and 40.4 ± 1.0 µm for the CCD group.
TTX-R currents activated at around –40 mV for both CCD and control groups (Fig. 3B) and reached maximal levels at –20 mV (Fig. 3B). The current amplitudes at –30 and –20 mV were significantly higher for the CCD group (–89.1 ± 23.1 and –109.7 ± 17.5 pA/pF, n = 9) than for the control group (–37.5 ± 12.8 and –61.9 ± 16.0 pA/pF, n = 9, Student's t-test), respectively.
There was no significant difference between CCD and control groups in activation and steady-state inactivation (Fig. 4B, Student's t-test). The mean mid-points and mean slope factors of activation curves were –25.9 ± 2.9 and 7.5 ± 1.8 mV for CCD (n = 9) and –28.6 ± 2.1 and 5.8 ± 1.1 mV for control (n = 9). The mid-points and slope factors of inactivation curves were –42.0 ± 3.3 and 2.5 ± 0.5 mV for CCD (n = 7) and –43.2 ± 4.0 and 4.5 ± 1.2 mV for control (n = 9).
The time constants of activation and inactivation were measured for voltages within the range of –40 to 10 mV (Fig. 5B). There were no significant differences in the mean activation or mean inactivation time constants between CCD (n = 9) and control (n = 9) (repeated-measures ANOVA).
Voltage-gated K+ currents
Voltage-gated K+ currents were recorded as shown in Fig. 6. The command potential protocol was initially undertaken from a holding potential of –100 mV and repeated from a holding potential of –30 mV. Because the inactivating components inactivated at –30 mV and the sustained current, Kdr, was evoked from –30 mV, a subtraction of the current waveforms elicited at the two holding potentials yielded the inactivating current, Ka (Fig. 6).
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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) and in ganglion neurons after dissociation (Ma and LaMotte 2005). Both of these studies used sharp electrode recording and did not label neurons from their peripheral receptive fields. In this study, we confirmed the enhanced excitability produced by CCD using whole cell patch-clamp recordings and focused on one particular neuronal population—cutaneous neurons of medium size that are labeled after Flurogold injection to the hindpaw.
Intact neurons with medium-sized cell bodies and myelinated axons include those that have peripheral nociceptors and others that have low-threshold mechanoreceptors (e.g., Fang et al. 2005; Ma et al. 2003). Nonnociceptive neurons with myelinated axons are thought to contribute to pain or altered pain states after peripheral inflammation or nerve injury by contributing, for example, to the production or maintenance of central sensitization or to the allodynia believed to be caused by the sensitization (Ma and Woolf 1996; Neumann et al. 1996; Noguchi et al. 1995). After CCD, medium-sized neurons can develop abnormal spontaneous activity and/or exhibit AP activity in response to inflammatory mediators (as can subpopulations of neurons with small and large sized cell bodies) (Ma and LaMotte 2005) and an increased density of hyperpolarization-activated cation current (IH) that may contribute to increased repetitive firing (Yao et al. 2003). The hyperexcitability of medium-sized neurons may therefore contribute to the cutaneous hyperalgesia that develops ipsilateral to a CCD injury (Song et al. 1999; Zhang et al. 1999).
The main signs of hyperexcitability in cutaneous, medium-sized neurons were a decrease in the threshold current/voltage required to generate an AP and an increased capability to repetitively fire. Several factors may contribute to the enhanced excitability observed in CCD neurons—a hyperpolarizing shift in the TTX-S activation curve, an enhanced magnitude of TTX-R current, and a reduction in Ka current in CCD compared with control neurons. The shift in TTX-S activation would cause a net increase in inward current at a more hyperpolarized potential and lead to a more negative voltage threshold for a regenerative depolarization to occur. An increase in TTX-R current would also support a more negative threshold voltage, because the threshold for a regenerative depolarization (Table 1) is above the threshold for TTX-R activation (Fig. 3). A decrease in Ka currents could also lead to a decrease in the accommodative properties of neurons after CCD, thereby increasing their capacities to fire repetitively (Yoshimura and de Groat 1999).
A decrease in Ka and an increase in TTX-R current would be expected to prolong the AP duration by reducing the repolarization current and enhancing the duration of Na influx. However, AP duration was not significantly altered in CCD. This was likely caused by the enhanced speed of inactivation of TTX-S current observed after CCD, which may have balanced the prolonging actions of TTX-R and Ka on AP duration.
Changes in Na channel expression or activity have been observed in other models of neuropathic pain, but the direction of change has been variable. A partial or total transection of the sciatic nerve caused an up-regulation of Nav 1.3, a TTX-S isoform that is usually expressed only in the neonatal period (Black et al. 1999; Dib-Hajj et al. 1999; Waxman et al. 1994). Nav1.3 is similarly up-regulated after carrageenan inflammation of the ipsilateral paw (Black et al. 2004; Tanaka et al. 1998). Although we did not identify changes in specific isoforms after CCD, we observed no change in the magnitude or steady-state properties associated with the inactivation of TTX-S current in our neurons. This suggests that there was no major change in TTX-S isoforms expressed in our cells. However, there was a leftward shift in the voltage of activation for the TTX-S current by 8 mV. Although the reason for this shift is not identified, a similar shift in activation is observed in hippocampal cells during exposure to pertussis toxin, suggesting that activation of a G protein–coupled pathway may occur with CCD (Ma et al. 1994). Results on TTX-R currents have been variable. We observed an up-regulation of TTX-R current after CCD—a result also observed in small neurons after carrageenan inflammation (Black et al. 2004; Tanaka et al. 1998). In contrast, the message for TTX-R current isoforms decreased after sciatic nerve transection, but it is unclear whether the membrane current was similarly decreased (Black et al. 1999; Dib-Hajj et al. 1999; Waxman et al. 1994). The variability of expression changes in different models of neuropathic pain suggest that multiple mechanisms may give rise to enhanced excitability of DRG neurons, resulting in a similar behavioral outcome.
Unlike changes in Na channel expression, changes in K+ current have been relatively consistent among models of neuropathic pain. After transection of the sciatic nerve, K+ currents (Ka, Kdr, and calcium-activated K+ currents) decreased in DRG neurons (Abdulla and Smith 2001b; Everill and Kocsis 1999). Ka was decreased in DRG neurons innervating an inflamed stomach wall (Dang et al. 2004) and an inflamed bladder (Yoshimura and de Groat 1999), and both Ka and sustained K+ currents were reduced in DRG neurons supplying an inflamed portion of the ileum (Stewart et al. 2003). In this study, the CCD-induced decrease in Ka currents was not accompanied by a change in sustained K+ currents. A selective down-regulation of Ka currents, which could contribute to AP threshold and repetitive firing of DRG neuron, may increase the response to a nociceptive stimulus and contribute to cutaneous hyperalgesia—i.e., the enhanced withdrawal to noxious stimuli that occurs after CCD (Song et al. 1999).
Different isoforms of voltage-gated K+ currents are expressed in DRG neurons, and several may change after injury. For instance, the mRNAs of Kv1.1, 1.2, 1.3, 1.4 (Ka), kv2.2, and kv4.2 (Ka), 4.3 (Ka) are expressed in DRG neurons and were reduced after sciatic nerve injury (Ishikawa et al. 1999; Yang et al. 2004). Although we have not identified a specific isoform changed in CCD, a likely candidate is one contributing to a fast inactivating current, such as would be expected from Kv1.4 and Kv4.2. Currently, experiments are underway in our laboratory to provide a better molecular identification.
The increased expression of TTX-R and reduced expression of Ka may both contribute to the tendency of CCD neurons to generate multiple APs during a sustained depolarization. TTX-R currents in DRG (primarily Nav.1.8) inactivate at more depolarized potentials than TTX-S currents and thus would be available to mediate a regenerative depolarization during a prolonged stimulus. Previous work from our laboratory also showed an increased expression of hyperpolarization-activated current (Ih) current in CCD neurons. This current is activated during the AHP that follows an AP and leads to a sustained depolarizing current, resulting in repetitive firing (Yao et al. 2003). Thus multiple changes in Na channel expression, Ka channels, and Ih currents may contribute to repetitive AP generation in CCD neurons.
In summary, this study identified several changes in Na+ and K+ current characteristics that may explain the enhanced excitability of DRG neurons after CCD. These are 1) a negative shift in the voltage of activation of TTX-S currents but no change in overall level of expression, 2) an increase in TTX-R current expression, and 3) a decreased expression of a fast inactivating K+ current.
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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ACKNOWLEDGMENTS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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Present address of Z.-Y. Tan: 602 MRC, Department of Internal Medicine, University of Iowa, Iowa City, IA 52242.
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FOOTNOTES |
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Address for reprint requests and other correspondence: R. H. LaMotte, Dept. of Anesthesiology, Yale Univ. School of Medicine, New Haven, CT 06510 (E-mail: robert.lamotte{at}yale.edu)
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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 J.-H. Zheng, E. T. Walters, and X.-J. Song Dissociation of Dorsal Root Ganglion Neurons Induces Hyperexcitability That Is Maintained by Increased Responsiveness to cAMP and cGMP J Neurophysiol, January 1, 2007; 97(1): 15 - 25. [Abstract] [Full Text] [PDF]
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J. H. Sun, B. Yang, D. F. Donnelly, C. Ma, and R. H. LaMotte MCP-1 Enhances Excitability of Nociceptive Neurons in Chronically Compressed Dorsal Root Ganglia J Neurophysiol, November 1, 2006; 96(5): 2189 - 2199. [Abstract] [Full Text] [PDF]
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C. M. Hingtgen A New "Kine" of Pain: MCP-1 and Sensory Neuron Excitability. Focus on "MCP-1 Enhances Excitability of Nociceptive Neurons in Chronically Compressed Dorsal Root Ganglia" J Neurophysiol, November 1, 2006; 96(5): 2171 - 2172. [Full Text] [PDF]
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and 
, respectively). *P < 0.05 (Student's t-test, CCD vs. control).
