Loss of T-type Calcium Current in Sensory Neurons of Rats with Neuropathic Pain

As previously reported in detail, we used a CCI model of rat peripheral neuropathy for our studies. 19Briefly, after Animal Resource Center approval, the right sciatic nerves of 175- to 200-g male rats anesthetized with halothane were ligated with four loosely tied 4-0 chromic gut ligatures under halothane anesthesia. 23Sham rats received the same surgical exposure of sciatic nerve, but no ligatures were placed. The left leg of both groups remained intact for comparison to the operated side. To allow for determination of which neuronal somata projected axons to the sciatic nerve operative site, fluorescing DiI (1,1′-dioctadecyl-3,3,3′,3′-tetramethylindocarbocyanine perchlorate) crystals were placed on the sciatic nerve and enclosed in a silicone rubber membrane (0.005 in; Technical Products, Inc., Decatur, GA) to prevent absorption of the dye into surrounding muscle. Ten days after surgery, we tested the rats for behavioral symptoms of hyperalgesia and allodynia. Briefly, rats were placed on a prewarmed glass surface, and their feet were exposed to a focused beam of incandescent heat with an infrared timer to detect paw withdrawal. Only rats with hind paw withdrawal latency less than 1.5 s shorter on the ipsilateral side compared to the contralateral side were included in the study. In addition, response to nociceptive mechanical stimulation in the plantar region of the hind paw was measured by indenting the skin with the point of a 22-gauge spinal needle. A normal response was a brief (nominally 0.5 s) withdrawal, whereas neuropathic response involved sustained elevation of the foot. In all, a neuropathic pain state was confirmed in approximately 50% of injured animals that exhibited a positive response to both these behavioral tests.

Our previous findings 19and other published reports 16suggest that pronounced T-type currents are primarily found in the medium-sized cells (33–42 μm) representing the nociceptors with Aδ lightly myelinated axons. 24Therefore, only medium cells were used in this study.

The L4 and L5 DRG were removed from rats after halothane anesthesia and decapitation. To retard necrotic degradation, the spinal cord was perfused with cold, oxygenated rodent Ringer's solution consisting of 146 mm NaCl, 2 mm CaCl², 1 mm MgCl², and 1 mm HEPES through a spinal transection at the midsacral region. Ganglia were quickly removed and bathed in piperazine-N,N ′-bis (2-ethanesulfonic acid) (PIPES)–buffered saline composed of 120 mm NaCl, 5 mm KCl, 1 mm CaCl², 1 mm MgCl², 25 mm d-glucose, and 20 mm PIPES. Excised ganglia were segmented with iris scissors to assist enzymatic digestion. Minced ganglia were transferred with silicone-coated Pasteur pipettes into 25-ml tissue culture flasks for shaking in enzyme solution containing 0.016% liberase blendzyme 2 (Roche Diagnostics Corp., Indianapolis, IN), 0.04% trypsin (Boehringer Mannheim, Mannheim, Germany), and 0.01% deoxyribonuclease 1 (150,000 U; Sigma, St. Louis, MO) in 5 ml oxygenated PIPES-buffered saline at a pH of 7.4 for 35–40 min at 150 rev/min. After complete dissociation was confirmed microscopically, cells were centrifuged at 778 g  for 6 min. The supernatant was removed, and cells were resuspended in a culture medium consisting of 0.5 mm glutamine, 0.02 mg/ml gentamicin, 100 ng/ml nerve growth factor 7S (Alomone Labs, Jerusalem, Israel), 2% (v/v) B-27 supplement (Life Technologies, Rockville, MD), and 98% (v/v) neurobasal medium A 1X (Life Technologies) before plating onto 12-mm poly-l-lysine–coated glass coverslips (Deutsche Spiegelglas; Carolina Biologic Supply, Burlington, NC). Coverslips were placed in a 12-well cell culture cluster and stored in a water-jacketed, humidified incubator at 37°C with 95% air and 5% CO². All cells were studied 6–8 h after dissociation.

To isolate LVA from HVA I^Ca, cells were preincubated for at least 30 min in an external Tyrode's solution of the following composition: 140 mm NaCl, 4 mm KCl, 2 mm CaCl², 10 mm d-glucose, 2 mm MgCl², 10 mm HEPES, 0.001 mm ω-conotoxin (CTx) GVIA, and 0.002 mm ω-CTx MVIIC at a pH of 7.4 with NaOH and an osmolarity of 300 mOsm. ω-CTx GVIA irreversibly blocks N-type calcium channels, and ω-CTx MVIIC irreversibly blocks P-/Q-type calcium channels. 25–27The concentrations used were saturating in preliminary experiments in this laboratory. Antagonist specificity was an issue in this study because no selective T-type Ca channel antagonist exists. Dihydropyridines were not used because they are known to block LVA Ca channels. 28In addition to the irreversible block of HVA, the ω-CTxs used in this study reversibly block substantial amounts of LVA I^Ca. This was evident in control studies in which preincubated cells were subsequently exposed to either ω-CTx GVIA or MVIIC applied in the bath through a multibarreled microperfusion pipette placed 200 μm from the cell to determine sensitivity of the remaining current to acute administration. ω-CTx GVIA reduced current remaining after preincubation by 74%, while ω-CTx MVIIC reduced current by 86%, and these reductions were completely reversible. For this reason, ω-CTx was not used in the perfusion chamber during patch clamp studies. Any residual HVA I^Cafollowing incubation was eliminated by using fluoride in the internal pipette solution, which runs down HVA without reducing LVA I^Ca. 21,29,30The efficacy of HVA rundown from fluoride was tested in six cells after preincubation in ω-CTx GVIA and equilibration with fluoride in the pipette. Application of 200 nm nisoldipine, a potent L-type channel blocker, in the bath had no effect on g^max(baseline = 1.5 ± 0.3 pS/pF vs.  nisoldipine = 1.3 ± 0.2 pS/pF). Nickel sensitivity has been demonstrated for both R- and T-type LVA Ca channels. 27In cells preincubated with ω-CTx, 10 μm Ni²⁺reduced peak Ca channel conductance by 58.2 ± 1.6% in eight cells from neuropathic rats and 66.2 ± 0.02% in eight cells from sham rats (not significant). Ni²⁺was used as a marker for LVA I^Cain current clamp studies only. The efficacy of pharmacological isolation of LVA I^Cacan be determined by comparison to our previous report of whole cell I^Ca. 19In cells without ω-CTx or fluoride, peak I^Cavalues were 3.06 ± 0.3 and 2.22 ± 0.26 pS/pF for cells from sham and neuropathic rats, respectively, more than three times the peak current reported in this study after pharmacological blockade.

The internal pipette solution consisted of 10 mm EGTA, 40 mm HEPES, 2 mm MgCl, an d135–140 mm tetramethylammonium hydroxide (TMA-OH) adjusted to a pH of 7.15–7.2 with hydrofluoric acid at an osmolarity of 290–300 mOsm. After preincubation, coverslips with their attached cells were transferred to the tissue chamber and perfused with an external solution designed to block potassium currents consisting of 152 mm TEA-Cl, 1 mm 4-aminopyridine (4-AP), 10 mm HEPES, and 5 mm CaCl²adjusted to a pH of 7.4 with TEA-OH at an osmolarity of 310–315 mOsm. The final TEA concentration was calculated at 165 mm. All solutions contained cytochrome c  (0.1 mg/ml) to inhibit nonspecific protein binding.

Only medium-sized cells demonstrating DiI fluorescence after excitation at 528–553 nm and emission between 577–632 nm were studied. Experiments were conducted at room temperature in the voltage or current clamp mode of the whole cell configuration of the patch clamp method using a List EP7 patch clamp amplifier (Adams List, Westbury, NY) connected to a personal computer through a Digidata 1200B digitizer (Axon Instruments, Union City, CA). Data were acquired at 5–50 kHz and filtered at 3 kHz with an eight-pole Bessel filter. The pClamp8 software suite (Axon Instruments) was used for data acquisition and analysis.

Experiments designed to characterize I^Cabegan with an I-V protocol from a holding potential (V^H) of −100 mV. Leak current was subtracted post hoc  by applying linear regression analysis to data points below current activation and then subtracting linear leak from the remainder of the I-V profile, provided that the linear fit passed through zero. Peak current amplitude was used in analyzing activation, inactivation, and deactivation protocols as described in relevant figure legends in the text. For determination of the dynamic characteristics of LVA current, two protocols were used. First, standard recovery from inactivation protocols employed a 1,000-ms conditioning pulse to inactivate channels followed by a test pulse of equal amplitude at increasing time intervals from 25 to 1,000 ms in 25-ms increments. Second, a ratio of sustained current (I^S) to peak current amplitude (I^P) was used to measure the rate of current inactivation. Specifically, late sustained current at the end of a 500-ms step protocol ranging from −100 mV to 10 mV was divided by early peak current at the same voltage. To approximate how I^Camight respond to natural voltage stimulation, a voltage command in the form of an action potential (AP) waveform was employed, which was derived from an actual recorded AP in a medium-sized cell lasting 7 ms from a V^Hof −68 mV. Another series of experiments was performed using current clamp mode in Tyrode's solution to determine the effect of T-type calcium channel blockade on excitability. Specifically, the amount of current necessary to evoke a single AP in a 20-ms step protocol was applied for 200 ms, and the number of APs were determined as a measure of neural excitability.

Whole cell I^Cawas normalized to cell capacitance for comparison between cells. To characterize whole cell current, voltage dependence of I^Cafor each cell was fit to the following Boltzmann equation:

where I is current, G^maxis the maximum channel conductance, V^1/2is the voltage at which current is half maximal, k is a slope factor describing voltage dependence of conductance, V^Ris the reversal potential for current, and v is the membrane potential. Steady state activation and inactivation data were normalized to maximum current (I^max) or maximum conductance (g^max) and fit with a Boltzmann function with upper and lower confidence levels using Origin 6.0 (Microcal, Northampton, MA). Tail currents representing deactivation were easily fit to a single exponential function. The resulting time constants (τ) were averaged and described as a function of test potential. Average ratios of I^Sversus  I^Pwere plotted in studies of inactivation rate as a function of membrane potential in Origin and fit to a Boltzmann equation with upper and lower confidence levels. AP waveform data were normalized to cell capacitance and measured for peak inward current, area under the curve, half-width, and duration. Fitted parameters, central values, and AP waveform data were compared using Student t  tests for unequal size between neuropathic and sham groups, and significance was determined at P < 0.05. Values expressed are mean ± SEM, unless otherwise noted.

CCI injury produced a neuropathic state with clearly distinct behavioral characteristics compared to sham-operated rats. Sham rats had normal gait and paw appearance with an average nominal response latency of 0.5 s to punctate mechanical stimulation and an average latency of 7.0 ± 0.6 s to radiant heat. These were significantly different from neuropathic rats that displayed protective behavior and deformed paw appearance accompanied by a delayed response to nociceptive mechanical stimulation and holding the paw aloft for 5.3 ± 0.8 s (P < 0.01) as well as an average heat withdrawal latency of 5.4 ± 0.8 s (P < 0.05). Mean cell diameter was 35.3 ± 0.8 μm with an average cell capacitance of 48.8 ± 3.7 pF. Sixty-nine cells from 26 rats, 15 sham and 11 neuropathic, were used in this study.

Whole cell I^Cadiffered significantly between the two groups of cells (fig. 1and table 1). Inward Ca²⁺current in sham cells achieved peak activation at −10 mV where the conductance (g^max) was 0.99 ± 0.44 pS/pF, while current in neuropathic cells was maximally activated at 0 mV with a mean conductance of 0.36 ± 0.07 pS/pF. Current half-activation (V^1/2) was also significantly lower at −23 ± 3 mV for sham versus −12 ± 2 mV for neuropathic. Reversal potential (V^R) and the slope factors (k) were the same. Typical current traces (fig. 1, inserts) show how HVACC blockade in sham cells revealed a noninactivating current at extremely low voltages which gave way to a current at higher voltages with voltage-dependent rapid inactivation, a characteristic of T-type currents. Visible in neuropathic cells is a faster rate of inactivation at voltages generating peak currents compared to sham cells. Four of 17 neuropathic cells displayed characteristic T-type LVA Ca channels, while 16 of 22 sham cells exhibited currents with these features—a difference of P = 0.004 according to Fisher exact test for two-sided chi-square. Of the neuropathic cells lacking characteristic T current, 11 showed extremely short currents in which inactivation was not voltage dependent.

Various features of these currents, including voltage dependence and kinetics, differed between the sham and neuropathic groups. Neuropathic injury shifted the voltage dependence of activation in a depolarized direction (fig. 2). Activation was determined by dividing peak current by the driving force to determine chord conductance and expressed as proportion of peak conductance. Sham cells were half-activated at −25 ± 0.7 mV, while neuropathic cells became half-activated at −11 ± 0.7 mV (P = 0.04).

Steady state inactivation was measured in a conventional way by a 500-ms prepulse from V^Hof −100 to 0 mV in 10-mV increments followed by a second test pulse to −10 mV. Results were normalized to peak current (fig. 2). One half of inward Ca²⁺current was inactivated at −38 ± 0.7 mV in both the sham and neuropathic groups. Though voltage dependence of inactivation was unchanged by injury, the extent and kinetics of current inactivation were significantly different. Since macroscopic inactivation was immeasurably small at hyperpolarized potentials where current did not inactivate, a ratio of current at early peak (I^EP) to sustained current (I^S) was used to examine the extent of LVA Ca²⁺current inactivation (fig. 3). Inactivation was greater in neuropathic cells at voltages of −50 to −20 mV. At −48 (−51 LCL to −46 UCL) mV, sham I^Cawas 50% inactivated at the end of a 500-ms test pulse, while neuropathic I^Cawas 50% inactivated at −38 (−44 LCL to −39 UCL) mV. The extent of inactivation was more steeply voltage dependent in sham versus  neuropathic cells (fig. 3A, fig. 1insert). Voltage differences in the ratio reflect higher peak conductance at lower membrane potentials, but they do not indicate the kinetics of channel inactivation. To estimate these rates, the trace at voltages showing 50% inactivation was fit to a single exponential for each cell. Neuropathic cells were 50% inactivated significantly faster at (τ= 186 ± 32 ms, n = 16) compared to sham cells (τ= 268 ± 36 ms, n = 18, fig. 3B). These significant differences in extent and pace of inactivation did not affect time to recover from inactivation (fig. 4). Recovery from inactivation for both groups was easily fit by a two-exponential nonlinear function with τ^fast= 409 ± 22 ms; τ^slow= 68 ± 4 ms for sham and τ^fast= 447 ± 45 ms; τ^slow= 64 ± 6 ms (not significant) for neuropathic.

Since T-type Ca²⁺currents deactivate (revert to nonconducting state upon repolarization) slowly, tail current kinetics were measured after a 20-ms depolarizing prepulse to a voltage that achieves peak activation followed by a return to test potentials ranging from −10 mV to −100 mV. Figure 5shows a family of tail currents with the typical pattern of T-type Ca channel relaxation. Tail currents were fit by a single exponential, and the time constant of deactivation, τ, was plotted against membrane potential. Deactivation was clearly voltage dependent in sham cells where the decay curve was nicely fit by an exponential function, whereas deactivation was not voltage dependent in neuropathic cells in which time constants for deactivation within the range of test potentials were easily associated with a linear function.

Neuropathy significantly reduced Ca²⁺current in response to an AP voltage command waveform (fig. 6). Peak current density dropped from −45.2 ± 13.4 pA/pF (n = 21) to −20.1 ± 4.6 pA/pF (n = 16) after neuropathy, and the area under the wave representing total Ca²⁺charge flux declined from 199.6 ± 91.4 pA/pF to 40.4 ± 5.7 pA/pF · ms after neuropathy. Likewise, duration of Ca²⁺current flux was reduced from 9.9 ± 1.3 ms to 5.2 ± 0.4 ms by neuropathy, and half-width declined from 2.5 ± 0.2 ms to 1.5 ± 0.1 ms.

Blockade of LVA Ca²⁺current with low concentrations of Ni²⁺or mibefradil enhanced neuronal excitability and decreased inward current in uninjured cells (fig. 7). Five micromolar Ni²⁺increased the frequency of APs twofold in response to a 200-ms ramped current injection, and this effect was reversible on washout. The presumption that this was due to Ca²⁺channel blockade is supported by the simultaneous decrease in inward current, which was reversed on washout. It is known that Ni²⁺also blocks Na⁺currents, but decreased I^Nawould be expected to decrease spike initiation rather than increase it. In another series using 300 nm mibefradil, a potent LVA Ca²⁺channel blocker with an EC⁵⁰of 3 μm, the frequency of APs evoked in response to a 200-ms ramp pulse rose significantly from 3.4 ± 0.9 (n = 7) to 6 .5 ± 0.8 (n = 6), while peak inward current significantly declined from 319 ± 49 pS/pF to 160 ± 77 pS/pF at a V^testof −10 mV (data not shown).

This study of the effect of neuropathic injury on LVA I^Cain rat DRG cells shows that low-voltage–activated calcium currents are significantly reduced after CCI of the peripheral nerve. The cause of this reduction appears to be a decrease in current density, a depolarizing shift in the voltage dependence of activation, an increase in the rate of deactivation, and more rapid and extensive inactivation. Even though I^Cain neuropathic cells was activated at more positive membrane potentials, there was no change in voltage dependence of channel availability as shown by the steady state inactivation curves. The shift in activation after neuropathy also diminished the window current seen in the overlap of inactivation and activation curves of sham cells. The combined effect of these changes was observed in response to an AP waveform. Depressed Ca²⁺influx is shown by reduced area under the curve in neuropathic cells as well as diminished peak current, and accelerated deactivation appears in the shorter duration of I^Cain response to a physiologically relevant command. Since LVA activity cannot be isolated during current clamp protocols, the contribution of LVA Ca²⁺current to membrane excitability was mimicked by administration of low-dose Ni²⁺or mibefradil to the cell in current clamp mode. Both mibefradil and Ni²⁺increased the firing frequency in response to a 200-ms current injection.

The origin of neuropathic pain after peripheral trauma or pathology is complex. Ectopic discharge in DRG somata of hyperalgesic rats has been implicated. 7However, the number of spontaneously active cells constitutes a small portion of all cells recorded in DRGs from neuropathic animals. 31Sympathetic sprouting into the DRG, accompanying Wallerian degeneration, also contributes to excitability in models of painful neuropathy, 32–35although the cells in our experiments were free from adrenergic inputs. Another physiologic response to injury is the phenomenon of cross-excitation, whereby spontaneously active or hyperexcitable cells can recruit adjacent cells, possibly through intercellular K⁺accumulation. 36Recent evidence suggests, however, that critical changes take place in pain by which cellular and chemical modifications cause primary sensory neurons to spontaneously discharge or oscillate secondary to mechanical stimulation. 13,37 

The cellular mechanisms of enhanced excitability are poorly understood. Widely reported changes in the expression of various Na channel subtypes suggest that faster spike initiation would presumably increase excitability. It is known that fast-activating, TTX-sensitive Na channels are overexpressed and slower-activating, TTX-resistant channels are underexpressed in sensory neurons after injury. 38Regardless of the initiation of neuronal activity, afferent signaling is ultimately limited by the ability of the sensory neuron to fire repetitively. Inward Ca²⁺current decreases neuronal firing rates by opening Ca-activated K channels, which make the cell less excitable. When Ca²⁺current is reduced, the cell becomes less stable and more excitable. 39–43 

Low-voltage–activated Ca²⁺currents have been implicated in rebound burst firing in thalamic relay neurons and cerebellar Purkinje cells associated with epilepsy 44and pacemaker activity in the upper heart. 45However, the role of LVA Ca²⁺currents in primary afferent neurons is unknown. One study in the DRG disclosed a rebound triggering mechanism, 17in which LVA currents are de-inactivated at very negative potentials and subsequently opened during return to resting membrane potential, admitting an inward current. However, the extreme hyperpolarization used in that study to produce this response is not characteristic of sensory neurons, and the very short (1- to 5-mS) duration of sensory neuron APs results in minimal channel inactivation. It is more likely, rather, that the role of T currents in primary afferent neurons, as a conduit of substantial Ca²⁺entry, 46is due to slow deactivation at the end of an AP, at which time the driving force on Ca²⁺ions is also very large. 46Absence of this prolonged Ca²⁺influx in injured neurons would contribute to increased excitability. 47Of course, in the central nervous system, the role of LVACC may be entirely different.

Not all neuropathic cells were without T-type Ca²⁺current in our study, nor was it fully expressed in all sham cells. Indeed, several features of currents in our cells differ from published reports of LVA Ca²⁺current. For example, the midpoints of activation were −25 mV in our sham cells compared to −36 ± 0.5 mV in systems expressing only α¹subunits without auxiliary β or δ subunits. 48However, despite the difference in LVA Ca²⁺current profile in these cells, a significant loss of LVA Ca²⁺current accompanied neuropathic injury.

The current that remained after injury was not due to Ca²⁺passing through TTX-insensitive Na channels because the remainder current was sensitive to the Ca channel blockers Cd²⁺(data not shown) and Ni²⁺. Furthermore, 160 mm TEA and 1 μm TTX were present in the external solution to substitute for Na⁺and block the TTX-sensitive Na channels, respectively. A more likely explanation for the remainder current in neuropathic cells involves the newly recognized heterogeneity of LVA Ca channels. Of the three subtypes, α1^G, α1^H, and α1^I, only α1^Hand α1^Iare differentially expressed in the DRG, according to in situ  hybridization examination. 49The α1^Isubtype inactivates more slowly than α1^H, 50which corresponds to earlier observations of fast- and slowly inactivating T-current subtypes. 51,52It would therefore appear that the slowly activating α1^Isubtype is preferentially inhibited by nerve injury, leaving intact a faster-inactivating α1^H, although the exact nature of the remainder current has yet to be determined.

Other studies of Ca²⁺currents in the DRG of neuropathic rats found decreases in whole cell Ca²⁺current, but they did not observe differences in the number of cells with detectable LVA Ca²⁺currents in sham versus  neuropathic groups. 20,53However, these studies did not specifically isolate T-currents with pharmacological agents. Furthermore, these studies either selectively studied identified cutaneous neurons 20or nonselectively examined injured and noninjured neurons. 53Deep, persistent pain characterized by hyperalgesia and allodynia preferentially arises from muscle afferents that become spontaneously active after nerve injury. We labeled more proximally at the sciatic nerve injury site, which means that both muscle and cutaneous afferents were included.

In sum, neuropathy sharply reduces inward current through LVA calcium channels of sensory neurons by shifting the voltage dependence of activation in a more depolarized direction, reducing calcium conductance and increasing the extent and rate of inactivation. These changes are reflected in a decreased total calcium current induced by a virtual AP waveform in injured neurons. Mimicking this loss by addition of low micromolar nickel or mibefradil produces a reversible increase in neuronal excitability. Future studies in nondissociated DRG tissue and intact neuropathic rats will be necessary to determine whether this loss of Ca²⁺current is responsible for increased excitability.

The authors thank Rich Rys (Senior Research Engineer, Department of Anesthesiology, Medical College of Wisconsin, Milwaukee, Wisconsin) for his valuable engineering help.