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REPORT
Departments of 1Anesthesiology, 2Neuroscience and 3Neuroscience Graduate Program, University of Virginia Health System, Charlottesville, Virginia; and 4Department of Anesthesiology and Pain Medicine, InJe University, Sanggyepaik Hospital, Seoul, South Korea
Submitted 17 September 2007; accepted in final form 16 April 2008
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ABSTRACT |
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INTRODUCTION |
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). However, the function of these channels in sensory and pain transmission (nociception) has only been discovered recently (Todorovic et al. 2001). New data have established the function of T-channels in supporting acute peripheral nociception. Furthermore, pharmacological and molecular downregulation of the function these channels in DRG neurons also supports the notion that T-channels contribute to the chronic pain associated with peripheral axonal injury (reviewed in Jevtovic-Todorovic and Todorovic 2006). However, in spite of the fact that T-type currents are expressed in several subpopulations of nociceptive DRG cells (Coste et al. 2007; Nelson et al. 2005; Todorovic et al. 2001), the cellular basis for the role of T-channels in chronic pain states is poorly understood. To address this issue, we used patch-clamp recordings to study the properties of T-type currents in acutely isolated and intact small nociceptive DRG neurons in animal model of painful peripheral neuropathy induced by chronic constrictive injury (CCI) of the sciatic nerve. |
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METHODS |
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Abnormalities in pain perception that are similar to those in humans, such as mechanical and thermal hyperalgesia, as well as mechanical allodynia, have been reported to occur in experimental rat models of mechanical injury of peripheral nerves as a consequence of loose ligation of the sciatic nerve, a CCI (Bennett and Xie 1988). We have published experimental procedures for CCI of the right sciatic nerve (Pathirathna et al. 2005; Todorovic et al. 2004) to induce mechanical injury to peripheral sensory nerve. For the present experiments, we used adult female retired-breeder Sprague-Dawley rats (250–350 g, 6–10 mo old). Control age-matched rats received either no operation (naïve animals) or sham operation (sham-operated animals). To determine the neuropathic state in CCI-treated rats, we measured thermal nociception in hind paws using thermal radiant heat testing as previously described (Pathirathna et al. 2005; Todorovic et al. 2001, 2004). For all behavioral data, we used ANOVA to compare the effects of CCI on thermal sensation. Subsequent pairwise comparisons between the pre- and post-CCI paw withdrawal latency (PWL) were done if significant P values resulted from two-way ANOVA. When appropriate, alpha levels were adjusted using the Bonferroni procedure (Pathirathna et al. 2005).
Before tissue harvest, rats were deeply anesthetized with isoflurane and rapidly decapitated. For one experiment, we dissected two dorsal root ganglia (DRGs), L4-5 from ligated (right-side) from one CCI or sham-treated rat. In control (untreated, naive) rats, we used bilateral L4-5 DRGs. We chose L4 and L5 DRGs because they contain the cell bodies of the majority of sensory fibers of the sciatic nerve. We prepared dissociated DRG cells and used them within 6–8 h for whole cell recordings as previously described (Todorovic and Lingle 1998). We focused only on small-size cells with an average soma diameter of 15–27 µm (Scroggs and Fox 1992) because many functional studies have confirmed that the vast majority of them are nociceptors (Caterina and Julius 2001; McCleskey and Gold 1999).
Recordings were made according to the procedures we described previously, using standard whole cell techniques with acutely dissociated DRG neurons (Jagodic et al. 2007; Nelson et al. 2005, 2007; Todorovic and Lingle 1998) and intact DRG neurons (Nelson et al. 2005). Series resistance (Rs) and capacitance (Cm) values were taken directly from readings of the amplifier after electronic subtraction of the capacitive transients. Series resistance was compensated to the maximum extent possible (usually 
60–80%). In most experiments, we used a P/5 protocol for on-line leak subtractions.
Drugs were prepared as 100 mM stock solutions of NiCl2 in H2O. The external solution used to isolate Ca2+ currents contained (in mM) 10 BaCl2, 152 TEA-Cl, and 10 HEPES adjusted to pH 7.4 with TEA-OH. To minimize contamination of T-type currents with even minimal high-voltage-activated (HVA) components, we used only fluoride (F–)-based internal solution to facilitate HVA Ca2+ current rundown; this solution contained (in mM) 135 tetramethyl-ammonium- hydroxide (TMA-OH), 10 EGTA, 40 HEPES, and 2 MgCl2, adjusted to pH 7.2 with hydrofluoric acid (HF). This allowed studies of well-isolated and well-clamped T-type currents in acutely isolated DRG cells (Todorovic and Lingle 1998). For voltage-clamp recordings with intact ganglia, the external solution contained the following (in mM): 140 NaCl, 4 KCl, 2 MgCl2, 2 CaCl2, 10 glucose, and 10 HEPES, adjusted to pH 7.4 with NaOH.; the pipette solution contained the following (in mM): 130 KCl, 5 MgCl2, 1 EGTA, 40 HEPES, 2 Mg-ATP, and 0.1 Na-GTP, adjusted to pH 7.2 with KOH.
All chemicals were obtained from Sigma (St. Louis, MO) unless otherwise noted. Unless otherwise indicated, statistical comparisons were made, where appropriate, using an unpaired Student's t-test, Mann-Whitney sum test, signed-rank test and 
2 test. All quantitative data are expressed as means of multiple experiments ± SE. The percent reductions in peak current at various Ni2+ concentrations were used to generate a concentration-response curve. Mean values were fit to the following Hill function
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 | (2) |
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In these forms, Imax is the maximal amplitude of current and Gmax is the maximal conductance, V50 is the voltage where half of the current is activated or inactivated, and k represents the voltage dependence (slope) of the distribution. The amplitude of T-type current was measured from the peak, which was subtracted from the current at the end of the depolarizing test potential to avoid contamination with residual HVA currents that was present at more positive membrane potentials (typically –20 mV and higher).
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RESULTS |
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). We confirmed the presence of CCI-induced thermal hyperalgesia before sacrifice (typically 7–14 days after CCI) for all animals used in voltage-clamp recordings. This was indicated by the reduction in thermal PWLs to 51.20 ± 0.01% of baseline values in right-side (ipsilateral, ligated) paws (n = 34, P < 0.001). In contrast, thermal PWLs in these rats in left-side (contralateral, nonligated) paws were not significantly affected: 99.80 ± 0.01% of baseline recorded before surgery (data not shown). Baseline values for PWLs were 10.47 ± 0.08 s in the right paws and 10.40 ± 0.08 s in the left paws. A sham treatment did not significantly affect thermal PWLs 
15 days after the procedure (data not shown). We recorded from a total of 167 small-size acutely dissociated DRG cells. The average diameter of cell somas was (in µm): 24.2 ± 0.3 in the control group, 24.3 ± 0.2 in the CCI group, and 24.6 ± 0.9 in the sham group (P > 0.05). These included 77 cells from control naïve rats (n = 22 animals), 14 cells from sham-operated rats (n = 2 animals) and 76 cells from CCI-subjected rats (n = 34 animals).
To compare the expression of T-type voltage-gated Ca2+ currents in small DRG cells after the induction of CCI, we held cells at –90 mV, then imposed voltage commands of depolarizing pulses from –60 to 0 mV in 10-mV increments. A representative family of inactivating inward currents in small DRG cell from control and CCI-treated animals is depicted in Fig. 1, A and B, respectively. Note that in both cells, T-type Ca2+ currents activate with small membrane depolarization, have a characteristic criss-crossing pattern, and display fast and almost complete inactivation during 250-ms-long test potentials. The average current-voltage curves were constructed from similar experiments, which indicated significant enhancement of T-type Ca2+ current amplitudes (measured from peak to the end of the depolarizing pulse) in CCI-treated animals; these currents were most prominent at negative membrane potentials and peaked at about –20 mV (Fig. 1C). To further determine the magnitude of the T-type current increase in CCI rats and to express it as current density, we normalized peak inward currents evoked at –30 mV to the cell capacitance in neurons from sham-operated, CCI-treated, and control rats. The histograms in Fig. 1D indicate that T-type current density was enhanced 1.5-fold in DRG cells from CCI rats as compared with cells from control (P < 0.05) and sham-operated rats (P < 0.01, Mann-Whitney test). Similarly, we found that the average T-current density in small DRG cells per rats had higher values in the CCI group (31.5 ± 4.2 pA/pF, n = 22 rats) than in the control group (20.6 ± 2.1 pA/pF, n = 16 rats, P < 0.05, Mann-Whitney test; data not shown). The average capacitance in these cells was (in pF): 19.4 ± 0.8 for the control group, 15.8 ± 0.7 for the CCI group (P < 0.01, t-test), and 19.3 ± 1.5 for the sham-operated group (data not shown). We also recorded T-currents from acutely isolated medium-size cells (n = 18) from CCI rats (n = 5) and found the following: average T-current density, 64 ± 12 pA/pF; average cell soma diameter, 33.8 ± 4.4 µm, average cell capacitance, 31.4 ± 2.7 pF; data not shown). None of these parameters was statistically different from those for medium DRG cells in control rats recorded under identical experimental conditions (Jagodic et al. 2007). Next we used data presented in Fig. 1D to generate linear correlation curves from scatter plots of T-type current density against cell capacitance in control and CCI groups. We found significant correlation only in the CCI group (P < 0.001), not in the control one (P = 0.2, data not shown). These data suggest that CCI-induced increase in T-type current density occurred predominantly in the subpopulation of smallest DRG cells.
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) (single-exponential fit of decaying portion of the current waveforms, Fig. 1F) from current-voltage curves in these cells over the range of test potentials from –50 to 0 mV. We found a small but significant difference between the controls and CCI-treated groups only for inactivation at 0 mV (Fig. 1F). The proportion of cells expressing T-type current was not significantly different in these two groups: control, 69% and CCI, 70% (2 test, data not shown).
We also tested voltage-dependent (steady-state) inactivation, finding that CCI caused a very small depolarizing shift in the midpoint (V50) of inactivation in these cells. For example, Fig. 2 B shows that the inactivation, V50, was about –69 mV in control cells (n = 13) and –66 mV in DRG cells from CCI-induced neuropathic rats (n = 17; P > 0.05). Likewise, the average V50 for T-type channel activation calculated from current-voltage curves was not significantly different in cells from the control (–42 mV, n = 15) and CCI (-41 mV, n = 24) groups (Fig. 2C). In contrast, we recently reported upregulation and a depolarizing shift in voltage-dependent inactivation of T-type channels in a subpopulation of medium-size DRG cells in rats with streptozotocin-induced diabetic neuropathy (Jagodic et al. 2007). Thus it appears that diabetic and mechanical neuropathy may affect T-type channels differently in different subpopulations of DRG cells.
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), knock-out (Chen et al. 2003), and molecular knock-down (Bourinet et al. 2005) have established that CaV3.2 is a predominant isoform of T-type channels in small DRG nociceptors from normal rats. Thus we examined the pharmacological properties of these cells in a CCI model of peripheral neuropathy, testing their sensitivity to nickel, a CaV3.2-specific pharmacological tool that is a traditional T-type channel blocker (Lee et al. 1999). Figure 3 A shows representative traces and B shows representative time courses from experiments in which nickel reversibly blocked T-type current in a concentration-dependent manner at 10 and 30 µM. However, when we compared IC50 for nickel in control cells (29 µM, n = 4 cells) versus DRG cells from CCI-treated rats (34 µM, n = 7 cells), there was very little difference between the two groups (Fig. 3C). There was no significant difference between cells in the control and CCI group with respect to the amplitude of T-type current blocked by 10, 30, and 100 µM nickel. For example, 30 µM nickel blocked 42.5 ± 4.5% T-type current in DRG cells from CCI rats (n = 4), and 50 ± 5% in DRG cells from control rats (P > 0.05).The fact that pharmacological sensitivity to nickel is very similar in both cells from CCI rats and control cells, together with the minimal changes in channel kinetics, strongly suggest that the molecular composition of T-type currents is little affected in small DRG cells from CCI-subjected rats.
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) and measured peak inward currents evoked from Vh –90 mV in 5-mV increments. We reasoned that voltage for half-maximal activation of total voltage-gated inward currents in these cells may be shifted to more depolarized potentials in the presence of 100 µM nickel. Indeed, Fig. 3D shows that nickel significantly increased the half-maximal activation threshold from –40 ± 1 to –31 ± 3 mV (P < 0.01; n = 11 cells from 4 rats) in CCI but not in DRG cells from sham-operated rats (baseline: –37 ± 3 mV, nickel –36 ± 3 mV; P > 0.05; n = 8 cells from 3 rats). Interestingly there was no significant difference in baseline half-threshold activation in the two groups. This could be explained by the concomitant decrease in TTX-resistant Na+ current density reported previously in small DRG cells in rats with CCI (Dib-Hajj et al. 1999). TTX-resistant Na+ channels activate over a range of potentials similar to that which activates T-type channels and are often co-expressed in the same DRG cells (Coste et al. 2007). |
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DISCUSSION |
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) as well as high-threshold mechano-sensory currents (Coste et al. 2007). Thus it is very likely that these T-channel-containing small DRG cells are polymodal nociceptors capable of responding in vivo to noxious heat, chemical, and mechanical stimuli. Recent data indicate that T-type channels have an important function in enhancing the cellular excitability of at least some small DRG cells by reducing the threshold for action potential firing (Nelson et al. 2005) and contributing to Ca2+ entry during action potentials (Blaire and Bean 2002; Nelson et al. 2005). In spite of their modest expression in most small DRG cells, increased T-type current density similar to that described here in our CCI model could be sufficient to increase the probability of the firing of action potentials because these cells have high-input resistance, as we have directly demonstrated in our current-clamp recordings using T-type channel modulators and CaV3.2 knock-out mice (Nelson et al. 2007).
It is important to note that remodeling of other voltage- and ligand-gated ion channels that can alter the excitability of the sensory neurons has been proposed to have a critical function in the development and maintenance of neuropathic pain symptoms such as hyperalgesia and allodynia (Campbell and Meyer 2006; Woolf 2004). Thus it is unlikely that changes in T-type current expression in the CCI model are the only culprit but may contribute to complex plasticity and to overall alteration in the cellular excitability of injured sensory neurons. In a simple model of measure of cellular excitability, we found that nickel at concentrations thought to be selective for T-type current significantly increased the threshold for half-maximal activation of total inward current largely carried by voltage-gated Na+ channels in these cells from CCI-subjected but not sham-operated rats. These data suggest that upregulated T-type currents may have a more prominent part in lowering threshold for spike firing in small DRG cells from CCI rats than in healthy rats.
Unlike pain that is caused by acute tissue injury (nociceptive pain), neuropathic pain resulting from constrictive nerve damage is a debilitating disorder that is inconsistently responsive to currently available conventional treatments. Of particular interest is our finding that several pharmacological blockers and modulators of T-type channels in vivo alleviate neuropathic pain in CCI. We determined that a series of 5
-reduced neuroactive steroids [e.g., (+)-ECN] are potent and selective blockers of DRG T-type channels in vitro (Todorovic et al. 1998). Furthermore, consistent with our present findings, (+)-ECN had a more potent analgesic effect when injected locally in peripheral receptive fields of sensory neurons in rats with CCI than it did when injected in control rats (Pathirathna et al. 2005). Dogrul and colleagues (2003) found that the preferential T-type channel blockers mibefradil and ethosuximide effectively reverse hyperalgesia and allodynia from CCI. We also found that oxidizing agents that block DRG T-type channel in vitro are capable of reversing CCI-induced thermal hyperalgesia in vivo (Todorovic et al. 2004). Moreover application of the T-type channel blocker Ni2+ blocks ectopic discharges from peripheral nerves in a model of segmental spinal mechanical injury (Liu et al. 2001). Toward this end, specific molecular silencing of CaV3.2 T-type channels in DRG cells with antisense reverses both hyperalgesia and allodynia in rats with CCI (Bourinet et al. 2005).
Surprisingly, in vivo study using CaV3.2 knock- out mice did not find a difference in pain perception in a CCI model (Choi et al. 2006). The exact reason for this discrepancy is not known, but it is possible that developmental elimination of CaV3.2 channels allows compensatory changes that are not feasible during acute downregulation of channel function using pharmacological agents or antisense applications. Interestingly, previous patch-clamp recordings of DRG Ca2+ channels in CCI also gave contrasting results. Hogan et al. (2000) reported no change in total inward Ca2+ currents in small DRG cells and loss of T-type current in medium-size DRG cells in rats with CCI (McCallum et al. 2003). These authors also reported that in control animals they did not observe T-type currents in any small DRG cells (<29 µm soma diameter). This is in sharp contrast to the results of this and previous studies (Blaire and Bean 2002; Cardenas et al. 1995; Coste et al. 2007; Scroggs and Fox 1992; Todorovic and Lingle 1998; Todorovic et al. 2001;), which found that T-type currents are expressed in the majority of small DRG nociceptors and that T-type current in medium-size DRG cells is not affected with peripheral nerve injury (Baccei and Kocsis 2000). Thus it is possible that different experimental conditions or the selection of different cells can account for the different findings with regard to the effect of CCI on expression of Ca2+ channels in small and medium DRG cells.
Our present study further implicates T-type channels as possible targets for the treatment of neuropathic pain resulting from mechanical injury to peripheral axons of sensory neurons. Thus blocking T-type channels may offer new therapeutic options for alleviating patients' suffering from chronic intractable pain resulting from partial mechanical injury of peripheral nerves. Future studies must be focused on the mechanisms of T-type channel alterations in these cells by CCI and in other animal models of painful neuropathy.
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GRANTS |
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FOOTNOTES |
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Address for reprint requests and other correspondence: S. M. Todorovic, Dept. of Anesthesiology, University of Virginia Health System, Mail Box 800710, Charlottesville, VA 22908-0710 (E-mail: st9d{at}virginia.edu)
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ABSTRACT
INTRODUCTION
) and CCI-subjected rats (B: Cm 25.3 pF, Rs 1.0 M
) and CCI (
) small-size DRG cells. CCI more prominently increased the absolute average amplitude of inward Ca2+ current at negative test potentials. For example, at Vt –30 mV, average control peak current was –274 ± 34 pA, while average peak in CCI group was –378 ± 29 pA (P < 0.05); Vertical bars indicate ±SE of multiple determinations. The amplitude of the total inward current at any given potential was measured from the end of the pulse to its peak. *, significance (P < 0.05 or higher) by t-test or Mann-Whitney U test. The numbers in parentheses indicate the number of cells in samples. D: histograms indicate average T-type current amplitudes in small DRG cells from control, sham-operated and CCI-subjected rats expressed as current density in pA/pF to correct for differences in cell size. Amplitude of T-type current was measured as maximal peak current subtracted from the small sustained current at the end of the depolarizing pulse at Vt –30 mV. Size of each sample is in the parenthesis; vertical bars are SE of multiple determinations. Peak T-type current density averaged 20 ± 2 pA/pF in the control, 30 ± 3 pA/pF in the CCI group and in sham group averaged 16 ± 4 pA/pF. **, significant value by Mann-Whitney test (P < 0.01); *, significant value by Mann-Whitney test (P < 0.05). E and F: we measured time-dependent activation (10–90% rise time, E) and inactivation 
