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Research Paper

Voltage-gated calcium currents in human dorsal root ganglion neurons

Hartung, Jane E.a; Moy, Jamie K.a; Loeza-Alcocer, Emanuela; Nagarajan, Vidhyaa; Jostock, Ruthb; Christoph, Thomasb; Schroeder, Wolfgangb; Gold, Michael S.a,*

Author Information
doi: 10.1097/j.pain.0000000000002465

1. Introduction

Voltage-gated calcium channels (VGCCs) in sensory neurons play an essential role in neurotransmitter release at the spinal dorsal horn presynaptic terminal and are therefore necessary for all forms of somatosensation, including pain. Because of this, VGCCs remain a viable therapeutic target for the treatment of pain. However, virtually, everything we know about the biophysical and pharmacological properties of VGCCs in sensory neurons was obtained through the study of rodent sensory neurons or heterologous expression systems. Although heterologous expression systems have played an essential role in drug development, it is now clear that the biophysical30,33 and pharmacological2,12,19 properties of ion channels in heterologous expression systems may differ significantly from system to system as well as from channels present in native tissues. More recent data have documented potentially important species differences as well.31,41,42 To address these issues, we have characterized high voltage–activated (HVA) current subtypes in acutely dissociated human and rat dorsal root ganglion (DRG) neurons.

2. Materials and methods

2.1. Human subjects

Data were collected from a total of 21 organ donors. The L4 and L5 DRGs were recovered by the Center for Organ Recovery and Education staff with consent from next of kin as described in Zhang et al.42 DRGs were placed in ice cold collection media comprised of 129.5 mM NaCl, 5 mM KCl, 1.2 mM MgSO4, 1 mM CaCl2, and 30 mM HEPES that had been filter sterilized after the pH had been adjusted to 7.35 with 5N NaOH. Dorsal root ganglion were transported to the laboratory on ice and processed immediately. All procedures were approved by the University of Pittsburgh Committee for Oversight of Research and Clinical Training Involving Decedents.

2.2. Animals

Data were collected from a total of 10 adult (200-350 g) male Sprague–Dawley rats. At least 3 rats were used for each endpoint. Females were not used simply because this was a convenience sample of rats that were available in the facility. Rats were pair housed in a temperature and humidity controlled, Association for Assessment and Accreditation of Laboratory Animal Care International accredited animal housing facility on a 12:12 light:dark schedule (lights on at 7 am). Food and water were available ad libitum. All procedures were approved by the University of Pittsburgh Institutional Animal Care and Use Committee and performed in accordance with National Institutes of Health guidelines for the use of laboratory animals in research.

2.3. Isolation and plating of human dorsal root ganglion neurons

Dorsal root ganglia were processed as described in Moy et al.21 In brief, after mechanical and enzymatic digestion, cells were spun down for 4 minutes at 450 G. The pellet was resuspended in L-15 basal media containing (in a 500 mL solution) 60 mg imidazole, 15 mg aspartic acid, 15 mg glutamic acid, 15 mg cystine, 5 mg ß-alanine, 10 mg myo-inositol, 10 mg choline-Cl, 5 mg p-aminobenzoic acid, 25 mg fumaric acid, 2 mg vitamin B12, and 5 mg of lipoic acid. Resuspended cells were placed on a Percoll gradient and spun at 199 G for 10 minutes. The pellet was then resuspended in complete media, which contained basal media, as well as 10% fetal bovine serum,, 10 ng/mL nerve growth factor (NGF 2.5S), 0.3 mg/mL glutamine, 0.225 mg/mL glucose, 2.55 mg/mL ascorbic acid, 0.12 mg/mL glutathione, and 0.2% (wt/vol) NaHCO3. Resuspended neurons were plated on 12 mm glass coverslips coated with poly-L-lysine. After plating, cells were placed in an incubator at 37°C with 5% CO2 for 4 to 6 hours before flooding with complete media. Although initial experiments used cells up to 80 hours after plating, we excluded cells in culture >48 hours from data analysis to minimize contamination from the voltage-gated sodium channel–derived outward current as described below.

2.4. Isolation and plating of rat dorsal root ganglion

Sensory neurons from L4 and L5 were obtained from adult rats, enzymatically and mechanically digested, and plated as described in Lu et al., 2006. Cells were plated on 5 mm coverslips coated with laminin and poly-L-ornithine. After plating, cells were stored for 2 hours in a CO2 incubator before flooding with L15 complete media. Rat sensory neurons were used within 12 hours of plating.

2.5. Whole-cell patch clamp

Whole-cell patch clamp recordings were conducted using a HEKA EPC10 using Patchmaster (v2x90.2) software (Harvard Bioscience, Inc, Holliston, MA). Data were acquired at 10 to 20 kHz. Borosilicate glass electrodes (World Precision Instruments, Sarasota, FL) were pulled with a Sutter P-97 (Sutter Instruments, Novato CA) to a resistance of 1 to 2 MΩ when filled with internal solution (see below). Amplifier circuitry was used for capacitance and series resistance compensation. Cells were held at −60 mV. A p/4 leak subtraction protocol was used from a holding potential of −80 mV for all protocols with the exception of the IV protocols, in which a holding potential of −100 or −50 mV was used to include or exclude low voltage–activated (LVA) currents, respectively.

A series of voltage-clamp protocols was used to characterize the biophysical properties of voltage-gated Ca2+ currents in DRG neurons. The first of these was an I-V protocol which included a 505 ms prepulse to −80 mV, followed by 50 ms test pulses to potentials ranging between −60 and +55 mV in 5 mV steps, followed by a return step to −80 mV, as used previously.17 This protocol was run with an inter-sweep interval (ISI) of 5 seconds. The second was an I-V protocol used to estimate current decay during a prolonged voltage step, referred to here as open-state inactivation. The protocol was the same as the first I-V protocol, except the test pulse was 500 ms in duration, and the protocol was only run from −60 to +35 mV. The third protocol was used to estimate steady-state inactivation, based on previous studies.22 This protocol consisted of a 500 ms conditioning step ranging from −90 to +10 mV, in 10 mV increments, followed by a 10 ms step back to −90 mV (to enable open channels to close), which was followed by a 50 ms test pulse to the potential corresponding to the peak inward current (generally between −10 and +10 mV). The final voltage step was to −80 mV. This protocol was run with an ISI of 10 seconds. The fourth protocol was a three-pulse protocol used to estimate the extent of G-protein βγ subunit-dependent inhibition of VGCCs. The protocol consisted of an 80 ms conditioning step to either −70 or +80 mV, followed by a 10 ms step to −90 mV, followed by a 30 ms voltage step to the peak inward current. This test step was followed by a return to −70 mV.22 This protocol was run with an ISI of 5 seconds. The fifth protocol was used to evoke tail currents. The protocol consisted of a 100 ms conditioning step to −90 mV, followed by a 50 ms voltage step to the potential associated with peak inward current, followed by a series of voltage steps ranging between −80 and −50 mV in 10 mV steps. This protocol was run with an ISI of 5 seconds. Finally, a subpopulation of neurons was stimulated with a voltage protocol representing an action potential (AP) waveform, similar to protocols previously conducted by others.3,25 To generate this voltage-clamp protocol, a subpopulation of neurons (8 cells from 1 donor) were studied in current clamp with physiological internal and external solutions (see below). Action potentials were evoked with a 5 ms current injection. The waveform used for the voltage-clamp protocol had properties (duration, overshoot, afterhyperpolarization amplitude, etc.) that were most similar to those of the mean of the population of neurons studied. This protocol was used to stimulate neurons 10 times at 5 Hz (to be able to estimate use-dependent changes in the evoked currents).

2.6. Solutions

To isolate Ca2+ currents, we used an extracellular solution containing 100 mM choline-Cl, 3 mM KCl, 10 mM HEPES, 10 mM glucose, 5 mM BaCl2·2H2O, and 0.6 mM MgCl2, adjusted to a pH 7.4 with tris base and to 325 mOsm with sucrose. The electrode solution was composed of 100 mM CsCl, 5 mM NaCl, 40 mM TEA-Cl, 1 mM CaCl2, 2 mM MgCl2, 11 mM EGTA, 10 mM HEPES, 2 mM ATP-Mg, and 1 mM GTP, adjusted to a pH 7.4 with tris base and 310 mOSm with sucrose.

For current-clamp experiments, cells were superfused with bath solution containing in mM: 130 NaCl, 3 KCl, 2.5 CaCl2, 0.6 MgCl2, 10 HEPES, and 10 glucose, pH 7.4 (osmolality 325 mOsm). The electrode solution contained (in mM) K-methanesulfonate 110, KCl 30, NaCl 5, CaCl2 1, MgCl2 2, HEPES 10, EGTA 11, Mg-ATP 2, and Li-GTP 1.

2.7. Drugs and chemicals

The voltage-gated sodium channel blocker tetrodotoxin was diluted in diH2O as a 10 mM stock solution and stored at 4°C. A803467, which blocks NaV1.8, was diluted in dimethyl sulfoxide (DMSO) at 10 mM and stored at -20°C. The L-type HVA current blocker nifedipine (Sigma-Aldrich, St. Louis, MO) was diluted in DMSO as a 100 mM stock solution and stored at −20°C until use. ω-agatoxin IVA and ω-conotoxin GVIA (Alomone Labs, Jerusalem, Israel), which block N-type and P/Q-type HVA currents, respectively, were diluted in diH2O as a 500 µM stock solution and stored at −20°C until use. CdCl2 (Thermo Fisher Scientific, Waltham, MA) was diluted in diH2O to a stock solution of 100 mM and stored at RT until use. [D-Ala2, N-Me-Phe4, Gly5-ol]-enkephalin acetate salt (DAMGO, Sigma-Aldrich) was diluted in diH2O to a stock solution of 1 mM and stored at −20°C until use. On the day of use, stock solutions were diluted in BaCl2 bath to a final concentration, indicated in parentheses: nifedipine (1 µm), ω-agatoxin IVA (200 nM), ω-conotoxin GVIA (200 nM), CdCl2 (100 µM), and DAMGO (1 µM).

2.8. Drug application

For all pharmacological experiments, bath containing VGCC blockers or DAMGO was superfused into the recording chamber through a gravity system at a rate of 1 mL/min. Neurons were considered responders to VGCC blockers or DAMGO if there was a decrease in peak inward current or an increase in the slope ∆I/∆Time of currents evoked at 0.2 Hz before blocker or DAMGO application. The full effect of VGCC blocker or DAMGO was considered to be achieved when Imax values stabilized over 3 pulses. Cells in which there was no detectable change in the slope or peak inward current after blocker or DAMGO application were classified as nonresponders. Because of the slow rates of washout for both ω-agatoxin IVA and ω-conotoxin GVIA, drugs were added sequentially as described in Lu et al., 2010 and the order of drug application for blockers was randomized to minimize the influence of the order in which blockers were added to the bath.

2.9. Data analysis

2.9.1. Inclusion or exclusion criteria

Neurons were excluded (1) if uncompensated voltage error was > 6 mV, (2) if outward current amplitude exceeded inward current amplitude (see below), and (3) if current density was <13 pA/pF. Seven human and 4 rat neurons were rejected for failing the first criterion, 70 human and 1 rat neurons studied < 48 hours in culture were rejected for the second criterion, and 19 human and 0 rat neurons were rejected for the third criterion. Furthermore, if neurons contained LVA currents (ie, currents activated at potentials < −30 mV), the data generated were not included in the analysis of HVA currents.

The biophysical properties of Ca2+ currents were analyzed with a combination of Fitmaster (V2x73.5, Harvard Bioscience, Inc) and SigmaPlot (v14.0, SYSTAT, Chicago, IL). Peak inward current was used for the I-V plots, and tail currents evoked after the first I-V protocol were used for G-V plots. G-V plots were fitted to a Boltzmann equation to extract the V1/2, slope, and Gmax. Pooled data were normalized to Gmax. Steady-state inactivation data were fitted to a Boltzmann equation modified to account for noninactivated current.

Kinetic properties including current activation, inactivation, and deactivation were determined with a single exponential fitted to the rising phase (activation), falling phase (inactivation), and tail currents (deactivation). Activation time constants (tau) were estimated from exponential fits of currents evoked with the first I-V protocol. Inactivation tau were estimated from exponential fits of currents evoked with the second I-V. Deactivation tau were estimated from exponential fits of currents evoked with the tail current protocol. Voltage dependence of these kinetic properties was determined by an exponential fit of the tau values plotted as a function of the voltage to derive the tau of tau. This tau of tau value indicates the voltage range over which the time constant of activation, inactivation, or deactivation has changed by 63%, where a smaller voltage range (smaller tau) reflects a greater voltage-dependent change in the time constant.

VGCC blocker and DAMGO data were plotted as a fraction of the total current. The fraction of blocker or DAMGO sensitive current was determined as the difference between the current evoked before and after the blocker or DAMGO relative to the total current. The total current evoked with the AP waveform was determined by digitally subtracting the current evoked in the presence of CdCl2 from the total current. The integral of the current was used to determine the charge and was measured during the rising and falling phases determined based on the peak of the AP. Blockers or DAMGO sensitive charge was determined relative to total charge. In experiments measuring Ca2+ influx sensitive to VGCC blockers, drugs were added sequentially or, in the case of both nifedipine and conotoxin, were added individually before the application of DAMGO. Data generated using the 2-pulse protocol were analyzed to determine the percent potentiation driven by the prepulse and were calculated as the percent change from P1. Data were analyzed using 2-tailed t tests or one-way, two-way, or three-way analysis of variances (ANOVAs) for parametric data and Mann–Whitney and Kruskal–Wallis tests for nonparametric data. χ2 analysis was used for proportion data. Correlation data were analyzed using a Pearson 2-tailed test. Significance was assigned if P < 0.05. When P < 0.05 for the ANOVA, data were analyzed with a Holm–Sidak post hoc test to determine differences between groups.

3. Results

A total of 195 neurons were studied from 21 human donors (17 male and 4 female), ranging in age from 21 to 77 years (46 average age), and 93 neurons from 12 male rats, ranging in age from 3 to 6 months.

3.1. Characteristics of high voltage–activated currents in rat and human dorsal root ganglion neurons

The same voltage-clamp protocol was used to evoke Ba2+ currents from rat and human DRG neurons (Fig. 1A). Currents in most neurons from both species had a high threshold for activation, demonstrated little inactivation over the 50 ms step duration, had peak inward current between −10 and +5 mV, and inactivated and deactivated relatively slowly (resulting in large tail currents). Although the peak inward currents in human DRG neurons (−6.01 ± 0.40 nA, n = 81) were larger than those in rat DRG neurons (−4.09 ± 0.35 nA, n = 70, P < 0.01), the current density (peak current normalized to membrane capacitance) was larger in rat DRG neurons (Fig. 1B, P < 0.01).

F1
Figure 1.:
Voltage-gated Ca2+ currents in rat and human dorsal root ganglion (DRG) neurons. (A) A series of depolarizing voltage steps were used to evoke high voltage–activated (HVA) currents in rat and human DRG neurons. (B) Pooled HVA I-V data from human and rat DRG neurons. Inset: Variability in peak inward HVA current evoked in human and rat DRG neurons (n = 87 humans and 57 rats), where each point represents a single neuron. (C) Pooled G-V data from the same neurons plotted in B. G-V data for each neuron was used to estimate Gmax, the voltage-dependence of activation (slope), and the voltage (V1/2) at which conductance was half of Gmax. Insets: Variability in slope and V1/2. (D) There was a negative correlation between the mean peak current density (Imax pA/pF) per donor (n = 21) and donor age. Black line is a linear fit of the data with gray area of 95% confidence interval. (E) With time in culture, there was an increase in a voltage-dependent transient outward current (top traces), which was not Ca2+-dependent but was abolished by blockade of TTX-sensitive and TTX-insensitive voltage-gated sodium currents with 300 nM TTX, 300 nM A803467, and changing the holding potential to −40 mV (bottom traces). (F) Low voltage–activated (LVA) currents (activation threshold < −35 mV) were detected in a subpopulation of both human and rat DRG neurons. Low voltage–activated currents detected in rat DRG neurons were slowly activating, slowing inactivating, and slowly deactivating. No open-state inactivation was detected in human LVA currents. Inset: LVA current evoked at −35 mV in rat (top trace) and human (bottom trace) DRG neurons. (G) Pooled I-V data for LVA currents detected in human (n = 18) and rat (n = 18) DRG neurons. Currents were evoked after a 500 ms prepulse to −100 mV. *P < 0.05, **P < 0.01. Data are mean ± SEM.

Of note, although data from several recent studies suggest that properties of putative nociceptors are enriched in human DRG neurons with a small cell body diameter, human DRG neurons selected for study were not chosen based on the cell body size. Indeed, the membrane capacitance of the neurons studied ranged from 55 to 687 pF. Nevertheless, to determine whether there was an association between cell body size and HVA current density, we analyzed HVA currents as a function of membrane capacitance (as a more sensitive measure of cell body size). Results of this analysis indicated that although there was a significant correlation (P < 0.01) between peak inward current and cell body capacitance (ie, there were larger currents in larger neurons), there was no association between current density and cell body size (P > 0.05). This contrasts with the positive relationship between cell body size and HVA current density we have previously reported in rat DRG neurons (where the current density increases as the cell body size increases).17 To further explore this relationship in human DRG neurons, we compared the current density in the 20% of neurons with the smallest cell body size with that observed in the 20% of neurons with the largest cell body size. Again, there was no significant difference between these groups with respect to HVA current density.

Tail currents recorded in response to membrane repolarization at the end of the 50 ms test pulse were used to generate G-V curves. Only neurons that did not contain LVA current (activating < −35 mV) were included in this analysis. Although HVA currents in human DRG neurons were larger than those in rat DRG neurons, the G-V curve for human neurons was right-shifted compared with that of the rat, suggesting that differences in G-V and current density were not because of clamp control. Rather, the difference in the G-V seemed to be largely because of the voltage dependence of activation (slope of the G-V) of HVA currents that was less in human neurons than that of rodent sensory neurons (8.0 ± 0.31 mV per e-fold increase in G, n = 59 vs 6.5 ± 0.30 mV per e-fold increase in G, n = 43, P < 0.01, 2-tailed t test, Fig. 1C, inset). The V1/2 of HVA current activation in human sensory neurons was also significantly more depolarized than that of HVA current in rat sensory neurons (−6.7 ± 0.79 mV, n = 73 vs −12.4 ± 0.51, n = 40, P < 0.01, 2-tailed t test, Fig. 1C, inset).

Because of evidence from rodent sensory neurons indicating that HVA current decreases with age,14 we assessed if the age-affected HVA current density, as human neurons studied were from donors considerably older than the rats were used in this study. This analysis was performed by averaging the peak inward HVA current density of each neuron studied from a donor and then plotting the average current density per donor as a function of donor age. Consistent with previous rodent data, there was a significant negative correlation between current density and age (P < 0.05, Pearson 2-tailed correlation test Fig. 1D).

To confirm that we were isolating only HVA currents in both rat and human neurons, we added 100 µM Cd2+ to eliminate all HVA currents in humans and rats. In acutely dissociated human DRG neurons, inward currents were completely blocked by Cd2+ (Fig. 1E, inset). This was also true for neurons in culture for > 48 h, but a large outward current emerged with time in culture (Fig. 1E). Initial experiments suggested that this outward current was Ca2+-dependent, which we assumed would other be carried by either Cl or K+ channels. Subsequent analysis ruled out either channel subtype, however, and confirmed that the outward current was Ca2+-independent. This left voltage-gated sodium channels as the only remaining likely possibility based on the composition of the external and internal bath solutions used. This was subsequently confirmed by the observation that the outward current was abolished by a combination of 300 nM tetrodotoxin (TTX, to block TTX-sensitive channels), 1 μM A803467 (to block NaV1.8), and a holding potential of −40 mV (to block NaV1.9). Thus, we suggest that the outward current was carried by a combination of TTX-sensitive and TTX-insensitive voltage-gated sodium channels. Owing to concerns that outward current would contaminate Ca2+ current data, all neurons in this study were in culture for < 48 h. In addition, cells showing a large (peak outward current > peak inward current) outward current, including those within the 48 hours window, were eliminated from analysis (70 of 195 neurons).

Low voltage–activated currents, those activating at potentials < −35 mV, were detected in DRG neurons from both rats and humans (Fig. 1F). Interestingly, in contrast to the transient LVA currents detected in rodent sensory neurons, the LVA currents in human sensory neurons exhibited little open-state inactivation (Fig. 1F inset). Low voltage–activated currents in the rodent also tended to activate at lower potentials than those in human sensory neurons (Fig. 1G) and were present at a higher density. To minimize the potential impact of LVA currents on the properties of HVA currents in human and rodent sensory neurons, neurons with a current > 48 pA at −35 mV were excluded from further analysis. This was performed for 18 human DRG neurons and 18 rat DRG neurons.

3.2. Steady-state inactivation of high voltage–activated Ca2+ currents in rat and human dorsal root ganglion neurons

Typical currents evoked with a steady-state inactivation protocol are shown for rats and humans in Figure 2A. No inactivation (<10% current inactivated) was observed in 6 of 60 rat neurons and 0 of 83 human neurons (P < 0.01, Fisher exact test). In a small fraction of neurons (12 of 60 rat, 0 of 83 human neurons; P < 0.01, Fisher exact test), currents seemed to inactivate in 2 phases with a clear plateau between each inactivation phase. In the remainder of neurons studied (42 of 60 neurons in rat, 83 of 83 neurons in human), steady-state inactivation was present but resulted in the inactivation of only a fraction of the total inward current (Fig. 2B). Of the current that did inactivate, the V1/2 was more depolarized in humans, which appeared to be due to a difference, at least in part, in slope (Fig. 2B, inset). There was also a trend towards a difference in the fraction of inactivated current in rat and human neurons (P = 0.051, Fig. 2B, inset).

F2
Figure 2.:
Steady-state inactivation in high voltage–activated (HVA) currents in rat and human dorsal root ganglion (DRG) neurons. (A) A three-pulse protocol (shown beneath current traces) was used to assess steady-state inactivation of HVA currents. Most of the current evoked during the conditioning prepulse has been eliminated for clarity. Full current sweep is shown in insets. (B) Steady-state inactivation data were fitted with a modified Boltzmann equation enabling an estimate of the fraction of total current subject to steady-state inactivation, in addition to the V1/2 of inactivation and the slope of the inactivation curve. Data for each neuron was normalized to the fitted maximal current value (Imax). Data from rats (n = 49) and humans (n = 34) were pooled and plotted. Insets: Variability in V1/2 slope and fraction of inactivated current. Data are mean ± SEM. **P < 0.01.

3.3. Kinetic properties of high voltage–activated Ca2+ currents in rat and human dorsal root ganglion neurons

Although rat HVA currents were relatively homogeneous with respect to the rate of activation, there was heterogeneity in the activation rate of human HVA currents with a subpopulation in which currents activated relatively quickly (time constant of activation, tau < 3ms at 0 mV) and another in which the currents activated more slowly (tau > 3 ms). These current types were referred to as fast and slow (Fig. 3A). Pooled data supported this impression, such that rat currents generally activated more quickly than those in human neurons (Fig. 3B). The HVA current from rat DRG neurons was also relatively homogeneous with respect to the small but consistent voltage-dependent decrease in the tau of activation. By contrast, there were at least 3 subpopulations of human DRG neurons that could be defined by both the tau of activation (Fast and Slow) and whether the tau of activation was voltage dependent. That is, the tau of activation decreased in a voltage-dependent manner in the majority (19 of 19 and 9 of 17) of neurons with fast (Fast) and slow (Slow) HVA currents. However, in a subpopulation of neurons with slow HVA currents, there was little evidence of a voltage-dependent change in tau (Flat) (Fig. 3B). The voltage-dependent change in the tau of HVA current activation (tau of taus) was significantly different between Fast and Slow currents in human neurons (P < 0.05, one-way ANOVA followed by the Holm–Sidak post hoc test, Fig. 3C), indicating that the voltage-dependent change in the tau of activation was greater in Fast HVA currents than in Slow HVA currents in human DRG neurons.

F3
Figure 3.:
Kinetic properties of high voltage–activated (HVA) currents in rat and human dorsal root ganglion (DRG) neurons. (A) The initial increase in current associated with a depolarizing voltage step was fitted with a single exponential to estimate the time constant (tau) of current activation. There was little variability between rat neurons with respect to the rates of HVA current activation. However, there were at least 2 types of HVA currents in humans that could be distinguished based on the rate of current activation across a range of voltage steps. (B) The tau of HVA current activation was plotted against the voltage at which the current was activated. The voltage-dependent decrease in the tau of activation of HVA current from rat DRG neurons was well fitted with a single exponential. The voltage-dependent decrease in the tau of activation of Fast and Slow HVA currents observed in human DRG neurons, illustrated in A could also be fitted with a single exponential. However, there was a third subpopulation of human DRG neurons in which the rate of HVA current activation demonstrated little voltage-dependence (Flat). (C) Variability in the voltage-dependence of current HVA current activation in rats and the 2 subpopulations of human neurons in which it was possible to estimate the voltage dependence of the time constant of current activation (tau of the tau). (D) The presence of open-state inactivation of HVA currents was estimated with 500 ms depolarizing voltage steps. There was a subpopulation of rat DRG neurons in which HVA currents demonstrated little inactivation (left traces) and a second subpopulation in which an initial phase of fast inactivation was detected (middle traces). High voltage–activated currents in human DRG neurons demonstrated a consistent rate of inactivation across the entire voltage step (right traces). (E) The time constant (tau) of HVA current inactivation in all rat neurons in which fast inactivation was detected decreased with increasing voltage. By contrast, there were 2 subpopulations of HVA currents in human neurons, one in which there was a small but consistent voltage-dependent decrease (V-Dependent) in the tau of inactivation and a second in which there was no voltage-dependent (V-Independent) change in the tau of inactivation. (F) Variability in the voltage-dependent change in the tau of HVA current inactivation. (G) A tail current protocol was used to determine the voltage-dependence of HVA current deactivation. High voltage–activated current deactivation was relatively consistent in both rat and human DRG neurons. (H) Although the time constant of HVA current deactivation in human DRG neurons was consistently larger than that in rat DRG neurons, currents in both species demonstrated a voltage-dependent increase in the tau of deactivation. (I) Variability in the voltage-dependent change in the tau of HVA current deactivation. *P < 0.05, **P < 0.01.

In contrast to the activation data, there was greater heterogeneity in the open-state inactivation data in rat neurons, resulting in 2 neuronal subpopulations, one with little inactivation and another showing a rapidly inactivating component (26 of 58 neurons, Fig. 3D). Interestingly, while slow inactivation was detected in all human neurons, there seemed to be 2 subpopulations of neurons based on whether the time constant of slow inactivation was voltage dependent. That is, in one subpopulation of human DRG neurons (17 of 59), there was a small but consistent decrease in the tau of inactivation with increasing voltage (V Dependent, Fig. 3E). No voltage-dependent change in the tau of inactivation was detected in the other subpopulations of neurons (42 of 59, V Independent, Fig. 3E). By contrast, HVA currents in all rat neurons, in which open-state inactivation was detected, demonstrated a voltage-dependent decrease in the tau of inactivation (Fig. 3E). There was a significant difference between HVA currents in rat and human DRG neurons with respect to the voltage dependence of the tau of inactivation (P < 0.01, Student t test, Fig. 3F), which was greater HVA current evoked in DRG neurons from rats than humans.

Consistent with activation and inactivation, HVA current deactivation rates seemed to be faster in DRG neurons from rats compared with humans (Fig. 3G). Pooled data supported this impression, confirming that the tau of deactivation was faster at all potentials assessed (Fig. 3H). Furthermore, although the tau of deactivation was voltage-dependent in all HVA currents in both rat and human DRG neurons, there was a significant difference between rat and human with respect to the voltage dependence of the tau of deactivation (P < 0.01, Student t test, Fig. 3I), which was greater in HVA current evoked in DRG neurons from rats than humans.

3.4. Constitutive inhibition of high voltage–activated currents in rat and human dorsal root ganglion neurons

Prepulse potentiation was observed in rat and human DRG neurons before the application of the mu-opioid receptor agonist DAMGO (Fig. 4A). Variability in the degree of potentiation was relatively small in most rat neurons compared with human neurons. Furthermore, there seemed to be a subpopulation of human neurons in which the percent increase in current associated with the prepulse (% Potentiation) was relatively large (>2 SDs from the mean). Because of the presence of this subpopulation, potentiation of HVA currents was, on average, significantly greater in neurons from humans than rats (P < 0.01, Student t test, Fig. 4B).

F4
Figure 4.:
Prepulse potentiation in rat and human neurons. (A) Prepulse potentiation was assessed by comparing the current evoked at 0 mV after a 500 ms conditioning voltage step to either −70 mV (black trace) or +80 mV (green trace). Inset: Magnified view of the current evoked during the test pulse. (B) Variability in the prepulse potentiation (% Potentiation) of high voltage–activated (HVA) currents in rat and human dorsal root ganglion (DRG) neurons. Data are mean ± SEM. *P < 0.05.

3.5. High voltage–activated current subtypes in rat and human dorsal root ganglion neurons

There was heterogeneity in both rat and human neurons with respect to the amount of current blocked by nifedipine, ω-conotoxin GVIA, and ω-agatoxin IVA, with the greatest source of heterogeneity being the proportion of nifedipine or conotoxin-sensitive current (Fig. 5A). Nevertheless, there were significant species-dependent differences in the proportion of Ca2+ currents sensitive to the HVA blockers, with significantly more nifedipine-sensitive currents in human neurons and significantly more conotoxin-sensitive currents in rat neurons (two-way ANOVA, Holm–Sidak post hoc, Fig. 5B). There were no species differences in the proportion of current sensitive to ω-agatoxin IVA or the current resistant to the 3 HVA blockers used (Fig. 5B).

F5
Figure 5.:
Subtypes of high voltage–activated (HVA) currents differ in rats and humans. (A) HVA currents were evoked in rat and human neurons before (baseline) and after serial application of HVA channel blockers nifedipine (1 μM), ω-conotoxin GVIA (200 nM), and ω-agatoxin IVA (200 nM). Blockers were applied in a random order. Each family of traces is from a different neuron. (B) In DRG neurons from both species, there was heterogeneity with respect to the percent of total HVA current (% inhibition) blocked by each compound. Pooled data revealed a species difference with respect to the relative contribution of nifedipine-conotoxin and ω-conotoxin GVIA-sensitive current. There was a significant interaction between species and blocker (two-way ANOVA, P < 0.01). The results of post hoc comparisons (Holm–Sidak test) are indicated. (C) HVA current evoked with a typical action potential (AP) used as the voltage-clamp protocol was associated with Ca2+ influx during both the rising (Rise) and falling (Fall) phase of the waveform. The application of HVA channel blockers enabled determination of the channel subtypes contributing to the AP-induced Ca2+ influx. (D) There was a species difference in Ca2+ influx (charge) evoked with the AP waveform. (E) There was a species difference in the relative contribution of HVA current subtypes to the Ca2+ influx evoked with the AP waveform, although this was only manifest in the influx associated with the falling phase of the AP. There was again a significant interaction between species and blocker (P < 0.01 two-way ANOVA), with post hoc comparisons (Holm–Sidak test) indicated. Data are mean ± SEM. **P < 0.01. ANOVA, analysis of variance.

Although there were clear species differences in HVA currents evoked by a square voltage step, as a more physiologically relevant voltage protocol, we assessed the relative contributions of these HVA subtypes evoked by an AP waveform, as has been used previously.3 Data were collected by sequentially adding each HVA blocker as in Figure 5A. Because a p/4 protocol was not used with the AP waveform, Cd2+ was added at the end of recording to enable subtraction of capacitive and leak currents. Consistent with larger current amplitudes in human neurons, larger currents were evoked in human neurons with the AP waveform (Fig. 5C, black traces), as total Ba2+ influx was significantly larger in human (11.48 ± 1.49 pC) than rat (7.36 ± 0.90 pC, P < 0.05) neurons, a difference largely due to greater Ba2+ influx during the falling phase of the AP (Fig. 5D). Similar to results generated with a square pulse, the relative contribution of HVA current subtypes to total charge influx evoked with the AP waveform was primarily blocked by nifedipine and conotoxin (Fig. 5E). Interestingly, nifedipine-sensitive current seemed to contribute to Ba2+ influx during both the rising and falling phase of the AP waveform in humans, but not rat neurons. Finally, to assess the presence of a difference in the relative contribution of nifedipine-sensitive and conotoxin-sensitive current to the Ba2+ influx evoked with a square pulse or an AP waveform, proportion of block data was analyzed with a three-way ANOVA (species × stimulus type × channel blocker). Consistent with results of the analysis of data generated with each stimulus independently, there was a significant interaction between species and drug. However, the absence of a significant interaction between all 3 factors suggests that the apparent differences between the proportion of Ba2+ influx sensitive to an HVA blocker is stimulus independent.

As noted above, there was no evidence of an association between cell body size and HVA current density in human DRG neurons. However, previous evidence suggests that N-type currents are enriched in putative nociceptive rat DRG neurons,18,29 consistent with the results of this study. We therefore determined if there was a relationship between cell size and the proportion of current blocked by L-type/N-type channel blockers. We ran a two-way ANOVA on % inhibition by either L-type (nifedipine) or N-type (conotoxin) channel blockers in the smallest 20% and largest 20% of neurons and found that although there was a significant effect of drug (P < 0.05), there was no significant difference by size or interaction.

3.6. µ-opioid receptor-induced suppression of high voltage–activated currents in human and rat dorsal root ganglion neurons

Given evidence of species differences in the relative contribution of HVA current subtypes and of HVA subtype specific modulation by G-protein–coupled receptors (GPCRs),5 we assessed the impact of the µ-opioid receptor, a canonical inhibitory GPCR, on HVA currents in rat and human DRG neurons. In line with previously published data from our laboratory21 and others,4 the selective µ-opioid receptor agonist, DAMGO (1 μM) blocked ∼50% of the Ba2+ current elicited by a square pulse in 58.8% of rat and 72% of human neurons (Figs. 6A and B). High voltage–activated currents evoked during both the rising and falling phase of the AP waveform were blocked by DAMGO in neurons from both rats and humans (Fig. 6C). When only neurons considered responders based on data collected in the square pulse were included, there were neurons in which there was no detectable suppression of the current evoked with the AP waveform. Nevertheless, on average, the percentage of current blocked during the rising and falling phases was similar between rat and human neurons (Fig. 6D).

F6
Figure 6.:
Inhibition of high voltage–activated (HVA) currents with μ-opioid receptor agonist DAMGO. (A) HVA current was evoked in rat and human dorsal root ganglion (DRG) neurons before (baseline, black trace) and after (DAMGO, orange trace) application of DAMGO (1 μM). (B) The percentage of neurons responsive to DAMGO (% responders) and the magnitude of the HVA current inhibition (% inhibition) was comparable in rat and human DRG neurons. (C) HVA current evoked during the rising and falling phase of an action potential (AP) waveform was inhibited by DAMGO in DAMGO responsive rat and human DRG neurons. (D) Pooled data indicate that the percent of HVA current blocked by DAMGO during the rising and falling phase of the AP was comparable (P > 0.05, two-way ANOVA). Data are mean ± SEM. ANOVA, analysis of variance.

Based on previous data suggesting that the Gβγ subunit displaces the VGCC β subunit from the α subunit, resulting in a depolarizing shift in the voltage dependence of activation,24 we used the prepulse potentiation protocol to quantify the extent of the DAMGO-induced inhibition of HVA current that could be reversed with a depolarizing prepulse. Only neurons considered responsive to DAMGO were included in this analysis. DAMGO-induced suppression of HVA current was reversed with a depolarizing prepulse in both rat and human DRG neurons (Fig. 7A). There was heterogeneity in rats and humans with respect to the number of neurons that responded to DAMGO in which current suppression was reversed with prepulse potentiation (Fig. 7A), which was detected in 85.2% of human neurons and 66.7% of rat neurons (P > 0.05). Of those neurons that prepulse potentiation was detected, there was no species-dependent difference in the magnitude of the potentiation (Fig. 7B). These results are consistent with a Gβγ-dependent suppression of HVA current in most rat and human DRG neurons.

F7
Figure 7.:
DAMGO-induced inhibition of high voltage activated (HVA) current in rat and human dorsal root ganglion (DRG) neurons was reversed with a prepulse potentiation protocol. (A) HVA currents evoked with the protocol described in Figure 4 before and after application of DAMGO (1 μM). (B) Pooled data indicate that there was no difference between rat and human DRG neurons with respect to the increase in current associated with prepulse potentiation (% potentiation). That is, there was a main effect of DAMGO (P < 0.01), but not species difference (P > 0.05) or a significant interaction between DAMGO and species (P > 0.05, two-way ANOVA). ANOVA, analysis of variance

3.7. The impact of DAMGO on N-type and L-type currents

Rapid Gi-protein–dependent suppression of HVA currents is generally believed to be due to a change in N-type channel gating.35 However, our data suggest that other HVA subtypes may also be involved because DAMGO blocked 54% of the current in human neurons, whereas N-type currents comprised only ∼20% of the total HVA current in human neurons. Therefore, we assessed the impact of nifedipine and ω-conotoxin GVIA on the current suppressed by DAMGO (Fig. 8A). In the rat, ω-conotoxin-GVIA reduced the percent responders and there was a trend toward a reduction in the magnitude of inhibition. In the human neurons, nifedipine had a dominant effect on the actions of DAMGO, at least with respect to the percent of neurons responsive to DAMGO (Fig. 8B).

F8
Figure 8.:
High voltage-activated (HVA) current subtypes inhibited by DAMGO in rat and human dorsal root ganglion (DRG) neurons. (A) HVA currents were evoked in rat and human DRG neurons before and after application of ω-conotoxin GVIA (200 nM, left traces) or nifedipine (10 μM, right traces) and then again after the application of DAMGO (1 μM). (B) There was a species difference in the percentage of neurons responsive to DAMGO when applied alone or in the presence of an HVA channel blocker (P < 0.05, χ2 left panel), but the impact of HVA channel blocker on the magnitude of the current inhibited by DAMGO (% DAMGO inhibition) was not significant (right panel).

4. Discussion

The aim of this study was to compare properties of HVA Ca2+ currents in rat and human DRG neurons. High voltage–activated currents isolated in both rat and human DRG neurons were similar in their biophysical and pharmacological properties. However, we identified 3 major differences between rat and human HVA currents: (1) human HVA current density was significantly smaller than that of rats, (2) a subpopulation of human neurons displayed significantly more constitutive inhibition relieved by a depolarizing prepulse, and (3) the proportion of HVA current subtypes was significantly different with a greater proportion of nifedipine-sensitive current and smaller proportion of ω-conotoxin IVA-sensitive current in humans compared with rats. A smaller species-dependent difference was the heterogeneity in kinetic properties of HVA currents in human DRG neurons not observed in rat. Finally, although the focus of this study was HVA currents, there were potentially interesting species-dependent differences in the magnitude and biophysical properties of LVA currents.

Although we suggest that the differences observed were species dependent, other factors may also have contributed, including age, time in culture, length of time between death and DRG recovery, and cause of death. Age is important to consider because of evidence in rodent neurons that HVA current density decreases with age14 and the fact that the median age for human donors was older than the rat by any comparative aging scale. Indeed, consistent with the age effect previously observed in rats, current density in humans DRG neurons decreased with increased age (Fig. 1D). Nevertheless, even when data from only the youngest human donors was included in the HVA current density analysis, the species difference persisted (data not shown). Similarly, evidence against a time-in-culture effect is that statistically significant differences in current density and kinetics persisted when only acutely dissociated human DRG neurons were analyzed (data not shown). The impact of time after death is difficult to account for, as there seems to be a species difference in the ability of sensory neurons to survive under hypoxic conditions: human neurons survive longer after cross clamp than rodent sensory neurons.42 Cause of death is also difficult to address experimentally. Nevertheless, we detected no influence of variables that could be used to differentiate the donor populations including cause of death, on HVA properties. This provides at least indirect evidence that cause of death is unlikely to have played a large role in the species differences in HVA currents. Thus, we suggest differences observed between rat and human DRG neurons with respect to HVA currents are largely species dependent.

There are at least 2 implications of lower HVA current density in human DRG neurons. First, Ca2+ influx at any level of activity will be smaller. Previous data from rat suggest that although the activity-dependent increase in intracellular Ca2+ is entirely dependent on the activation of VGCC, the Ca2+ influx through these channels is only responsible for the initiation of the evoked transient. Other Ca2+-regulatory machinery (eg, that underlying Ca2+-induced Ca2+ release from intracellular stores and the plasma membrane Ca2+-ATPase) is largely responsible for determining the magnitude and duration of the evoked transient at even moderate activity levels for relatively short burst durations.8,9,17 As a result, the magnitude and duration of activity-dependent increases in intracellular Ca2+ are largely independent of VGCCs at even moderate activity levels for relatively short burst durations.10,17,39 This dissociation is even more apparent with inflammation, where the magnitude and duration of activity-dependent increases of intracellular Ca2+ are increased despite decreased HVA current density.18,27,28 The relatively small current evoked by AP discharge in human DRG neurons21 suggests other Ca2+ regulatory processes do not compensate for low HVA current density. Thus, the first implication of lower HVA current density in human DRG neurons is that activity-dependent increases in intracellular Ca2+ will be smaller and more spatially restricted than in rat neurons, suggesting that cellular processes dependent on Ca2+ (eg, kinases and phosphatases) would have to respond to smaller increases in intracellular Ca2+ or be closer to the point of influx. That said, as discussed below, the slower kinetics of the HVA currents observed in human DRG neurons (Fig. 3) may also help compensate for the lower HVA current density when Ca2+ influx is driven by more physiologically relevant stimuli such as the AP waveform. Second, lower HVA current density and more localized, activity-dependent increases in Ca2+ may impact activity-dependent changes in gene expression.36 In light of compelling data linking neural activity, Ca2+, and changes in gene expression in rodent DRG neurons,7 initial injury-induced changes in gene expression in sensory neurons15,38 are believed to be influenced by activity-dependent increases in intracellular Ca2+.6 However, if activity fails to drive global increases in intracellular Ca2+ in human DRG neurons, then this form of plasticity would not be as robust in human DRG neurons, or other mechanisms may instead link neuronal activity and gene expression. From an experimental perspective, the tight association between VGCC activity and the magnitude and duration of Ca2+ transients in human DRG neurons suggests that Ca2+ imaging may provide a more accurate measure of activity in human DRG neurons than rats, even if transients are smaller and have a shorter duration.10

Although this study was not designed to identify subpopulations of sensory neurons, there were no unique characteristics of the human DRG neurons with large prepulse potentiation of HVA current. This population of neurons (12 neurons with potentiation > 1 standard deviation above the mean) had similar cell body diameter, HVA current density, and proportions of L-type and N-type currents (although only 7 neurons received a subtype selective channel blocker). Nevertheless, we ruled out 3 potential functional consequences of the large constitutive influence on channel gating. First, constitutive suppression of HVA currents did not influence current density in human DRG neurons as current density was still lower when potentiated currents were used to estimate peak current density. Similarly, if a peak inward HVA current was potentiated by a depolarizing prepulse, this might counter other mechanisms associated with decreased HVA current, thereby maintaining Ca2+ influx during a burst of APs. However, we observed a negative correlation between prepulse potentiation and the degree of change in current influx in response to 10 stimuli at 10 Hz (Pearson, P < 0.05). Furthermore, and potentially most importantly, there was no association between the magnitude of the prepulse potentiation and either the magnitude of the DAMGO-induced suppression of HVA current or the percentage of neurons responsive to DAMGO. Therefore, if prepulse potentiation reflects long-term GPCR signaling, the underlying mechanisms are distinct from those underlying short-term GPCR-dependent suppression of HVA currents. Additional experiments will be needed to elucidate the mechanisms and functional implications of constitutive suppression of HVA currents in human DRG neurons.

Given the therapeutic efficacy of ziconotide, an N-type VGCC blocker presumed to block transmitter release from central terminals of nociceptive afferents in the spinal dorsal horn,34 we were surprised that L-type currents comprised most human HVA currents. There are at least 2 likely explanations for differences in the relative density of HVA current subtypes. One is species differences in HVA channel subtype trafficking within the neuron (eg, in the cell body vs axon terminals). The second possibility is species differences in HVA channel subtype transcription, translation, or protein turnover. If the difference in HVA subtype density is due to trafficking, then results demonstrating that N-type channels underlie Ca2+ influx in presynaptic terminals in rodents are likely also true in humans.40 However, 2 observations favor the second possibility: (1) Despite the presence of N-type channels in human DRG neurons, DAMGO seemed to act preferentially through L-type channels, suggesting these, and not N-type channels,11 may be a target for presynaptic suppression of transmitter release; (2) genes encoding L-type channels (CACNA1(C/D/F) are expressed at relatively higher levels in human DRG than those encoding N-type channels (CACNA1B), whereas the opposite is true in mouse.26 If the differences in HVA current subtypes are not just restricted to the cell soma, then the relatively higher density of L-type channels in human DRG neurons would explain the limited therapeutic efficacy of drugs such as gabapentin that seem to preferentially reduce N-type currents.37

Although not as dramatic as differences in current density prepulse potentiation, and HVA current subtypes, there were interesting implications associated with slower kinetics of activation, inactivation, and deactivation observed in human HVA currents compared with rats. For example, slower rates of activation and deactivation delay Ca2+ influx through HVA channels during an AP, with influx corresponding to the falling phase and afterhyperpolarization of the AP. This shift in timing maximizes the driving force on Ca2+ ions and, as noted above, may compensate for lower HVA current density observed when current is evoked with a square pulse (Fig. 1). Ca2+ influx data generated with an AP waveform (Fig. 5D) are consistent with this suggestion. Similarly, the heterogeneity observed in the voltage-dependence of activation and deactivation of HVA current in human neurons may suggest species differences in expressed splice variants of HVA genes. For example, splice variants for genes encoding L-type and N-type channels can influence voltage-dependence of activation1,16,25 and the kinetics of channel activation.23

Finally, although the focus of this study was on HVA currents, it was difficult to ignore what seemed to be marked differences between rodent and human DRG neurons with respect to the amplitude and kinetics of LVA currents. That is, the LVA current density in human DRG neurons was smaller than in rat neurons, and the current detected demonstrated little open-state inactivation. Because rodent data suggest LVA currents play important roles in neuronal excitability and pain associated with tissue injury,13,20,32 further investigation of LVA currents in human sensory neurons is warranted. Taken together, these data add to a growing list of differences between rodent and human neurons and once again underscore the need to conduct research using human neurons to successfully develop effective therapeutics.

Conflict of interest statement

The authors have no conflicts of interest to declare.

Supplemental audio content

An audio abstract associated with this article can be found at https://links.lww.com/PAIN/B479.

Acknowledgments

The authors thank members of the Gold lab and the Pittsburgh Center for Pain Research for their feedback in writing this article. The authors especially thank the Center for Organ Recovery for tissue recovery. Most importantly, the authors thank the next of kin of tissue donors for their generous consent to the use of the donor's tissue for research purposes. This work was supported by the National Institutes of Health (R01NS083347 and R01DK107966 (M.S.G.), F32NS103231 and T32NS073548 (J.E.H.) and a grant from the Bloch Foundation (M.S.G.) and from Grünenthal GmbH. R. Jostock, T. Christoph, and W. Schroeder are or were employees of Grünenthal GmbH. While M.S. Gold has received research grants from Grünenthal, none of the authors have a conflict of interest with the material in this article.

Author Contributions: J.E. Hartung contributed to experimental design, collection and analysis of data, and manuscript preparation. J.K. Moy, E. Loeza-Alcocer, and V. Nagarajan cultured human DRG tissue. R. Jostock, T. Christoph, and W. Schroeder contributed to experimental design. M.S. Gold contributed to experimental design, data collection and analysis, and manuscript preparation.

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Keywords:

Conotoxin; Agatoxin; DAMGO; GPCR; MOR; Nociceptor; Voltage-clamp

Supplemental Digital Content

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