DRG cultures were obtained as described (Gandini et al., 2014). In brief, DRG were dissected from C57BL/6 mice (P6-P10) in Advanced DMEM Medium (Gibco) supplemented with 20% of Fetal Bovine Serum (Gibco), washed, and digested for 40 min at 37°C with a mixture of trypsin type XI (1.25 mg/ml, Sigma) and collagenase IV (1.25 mg/ml, Sigma), followed by mechanical dissociation. Cells were spun down at 1,000 g for 5 min at 10°C and re-suspended in Advanced DMEM medium supplemented with 10% FBS. Cells were plated onto L-lysine-covered coverslips (12 mm, Carolina Biological Supply, Burlington, NC, USA) and kept in a 5% CO₂ humidified atmosphere at 37°C. The Patch-clamp recordings were made 24 h after dissociation (1 day in vitro, 1 DIV).

Western blots. Crude protein was extracted from at least three WT or KLHL1 KO DRG ganglia pooled together and separated by SDS-PAGE electrophoresis (8%, at 100 V for 90 min) for transfer to a PVDF membrane (BioRad). Membranes were washed in Tris-buffered saline (TBS) supplemented with 0.05% Tween 20 (TBST) and blocked for 1 h in TBST-5% milk at room temperature (Florio et al., 1992). Membranes were incubated at 4°C overnight with primary antibodies against Ca_V3.1 (1:1,000, Millipore, CA, USA), Ca_V3.2 (1:2,000, Santa Cruz Biotechnology, Inc., Santa Cruz, CA, USA), Ca_V3.3 (1:1,000, Alomone), Ca_V2.1 (1:1,000, Alomone) or Ca_V2.2 (1:1,000, Alomone). GAPDH (Santa Cruz Biotechnology, Inc., Santa Cruz, CA, USA; 1:1,000) was used as an internal reference to normalize for protein loading. Horseradish peroxidase (HRP)-conjugated secondary antibodies were used for detection (1:2,000; Pierce) with Supersignal Femto (Pierce, IL, USA) using a ChemiDoc MP System (BioRad).

Immunoprecipitation. Crude membrane preparations were obtained using standard protocols (Aromolaran et al., 2010); a fraction of the sample was reserved prior to immunoprecipitation (input, 30× less concentrated than the IP samples) and the remaining volume was divided up for all experiments. Samples were processed by addition of primary antibodies (Ca_V3.2, 1:40 and KLHL1, 1:40, Santa Cruz Biotechnology, or IgG, 1:40, Alpha Diagnostic Intl. Inc., San Antonio, TX, USA) and incubated for 1–3 h at 4°C followed by overnight incubation with protein A/G agarose beads (Biovision, Mountain View, CA, USA) on a shaking plate at 4°C. Samples were washed and processed for western blot analysis as described.

Whole-cell patch-clamp recordings were obtained at 1DIV using an Axopatch 200B amplifier (Axon Instruments, Union City, CA, USA) at room temperature. Data were acquired at 1 kHz and digitized at 20 kHz. Calcium currents were recorded using an external solution containing (in mM) 5 CaCl₂, 140 TEA-Cl, 10 HEPES and 10 glucose (pH 7.4, 300 mosmol/kgH₂O). The intracellular solution contained (in mM): 108 CsMeSO₃, 4 MgCl₂, 10 Cs-EGTA, 9 HEPES, 5 ATP-Mg, 1 GTP-Li and 15 phosphocreatine-Tris. Pipette resistances were 3.0–4.0 MΩ. Series resistance (Rs) was compensated online (>80%), only cells with Rs <15 MΩ were used. Data were acquired and analyzed using pClamp10 software (Molecular Devices).

Total currents were elicited using depolarizing steps (test potentials, V_T) from −60 to +60 mV (ΔV = 10 mV) from a holding potential (V_H) of −90 mV. HVA currents were obtained from V_H = −50 mV to V_T = −60 to +60 mV (ΔV = 10 mV). HVA currents traces were subtracted from the total current traces at each V_H to obtain the LVA current component.

APs were measured using an external solution containing (in mM): 135 NaCl, 5 KCl, 2 CaCl₂, 1 MgCl₂, 10 HEPES, 10 glucose; the intracellular solution composition was (in mM) 110 K-gluconate, 20 KCl, 2 MgCl₂, 1 EGTA, 10 HEPES, 2 ATP-Mg, 0.25 GTP-Li and 10 phosphocreatine-Tris. The APs were triggered by four consecutive 1.5-s-long current depolarizing ramps at 20, 40, 60, or 80 pA/s. Rheobase was determined as the minimum current necessary to elicit an AP from a membrane potential of −75 mV.

Adeno-associated viral constructs containing small hairpin RNAs (shRNA) targeting KLHL1 were designed; shKLHL1-AAV contained two sequences from the mouse gene (NM_053105.2) spanning nucleotides (nt) 1,812–1,830 and 2,121–2,139 (GGCCAGTGATGATGTAAAT and GGGAATGGATAATAACAAA, respectively); these segments were synthesized and cloned into an AAV shuttle vector pZacf-U6-Luc-Zsgreen (U Penn Gene Therapy Core) as described before (Zolotukhin et al., 1999; Sarkey et al., 2011). These shuttle plasmids, along with pHelper and pAAV2/8 were transfected into AAV-293 cells using the Virapack transfection kit (Stratagene) and purified by an iodixanol step gradient as described before (Pradhan et al., 2010). Titer was assessed by serial dilutions of virus and infection of HT1080 cells. AAV particle titer was quantified by SDS-PAGE (Zolotukhin et al., 1999; Kohlbrenner et al., 2012). EGFP-AAV was generated in house by the same method.

All animal protocols used in this study were reviewed and approved by an independent Institutional Animal Care and Use Committee (IACUC). Hind paws of 13- weeks old WT male mice were injected with a control virus (EGFP-AAV, shCtrl,) or shKLHL1-AAV under blind conditions. The summary of the experimental conditions used is depicted in Figure 6A. The initial trial (n = 7) received 4.2 × 10¹⁰ shKLHL1-AAV or 5.5 × 10¹⁰ EGP-AAV vector genomes. The second trial (n = 11) received a high titer, 9.0 × 10¹⁰ shKLHL1-AAV or EGFP-AAV vector genomes over 2 days. Viruses were diluted such that each individual injection volume was 5 μl total. Mice were given pain medication (Buprenorphine, 0.05 mg/kg, s.c.) for the first 2 days following the last injection and were allowed to recover in observation for 4–5 days while checked for any limp or lameness; all mice were confirmed healthy after injections.

Behavioral tests were performed twice a week (and averaged) for a total of 3 weeks after injections. Baseline withdrawal threshold responses were determined for 1 week before injections.

% Paw-withdrawal threshold was reported as the % of mice in the total population displaying withdrawal thresholds at all forces tested.

Hind paw withdrawal experiments were carried out in male mice ~16 weeks old; animals had access to food and water ad libitum, all experiments followed IACUC-approved standard procedures. Only males were used in the first study because our preliminary data show male KLHL1 KO mice display a clear phenotype in contrast with females, where differences are more difficult to establish if present. Mice were placed on a wire mesh-bottom testing apparatus and allowed to acclimate for 15 min before assessing mechanical allodynia. Measurements were recorded by applying von Frey filaments (North Coast, Morgan Hill, CA, USA) ranging from 1.4 to 10 g to the plantar surface of the mouse hind paw; each filament was assessed for a total of five consecutive times. Hind paw withdrawal response times of less than 2 s were considered positive. The withdrawal threshold was calculated as the filament force at which each mouse had a positive response more than three times out of five (Chaplan et al., 1994; Bonin et al., 2014).

Mice were anesthetized with isoflurane before euthanasia; DRG were excised and fixed with 4% paraformaldehyde for 4 h, cryoprotected overnight (30% sucrose in PBS), embedded in OCT (Tissue Tek, Fisher Scientific, Hampton, NH, USA), and frozen with dry ice. 30 μm-thick sections were cut using a cryostat and mounted onto Superfrost Plus slides (Fisher Scientific, Hampton, NH, USA). Slides were washed three times with PBS-glycine, dried and protected with coverslips. Fluorescence images were captured using IX80 Olympus inverted epifluorescence microscope using a 10× objective and analyzed using deconvolution.

Statistical analysis was performed with SigmaPlot 11 Software. Statistical significance was determined as P < 0.05, using student’s t-test or Kolmogorov–Smirnov non-parametric analysis (Kolmogorov–Smirnov test). Results are presented as mean ± SEM.

Co-immunoprecipitation of Ca_V3.2 and KLHL1 is detected in overexpression experiments in HEK-293 cells and in whole brain samples, demonstrating direct interaction between these two proteins (Aromolaran et al., 2009, 2010, 2012). Figure 1 shows an example of pull-down experiments from DRG protein extracts using Ca_V3.2 (top) or KLHL1 antibodies (bottom, IgG was used as negative control). These data confirm the presence of KLHL1 in DRG neurons and its interaction with Ca_V3.2 T-type channels.

We next assessed the effect of KLHL1 deletion on Ca_V expression in DRG neurons. HVA channel expression was statistically similar in both KLHL1 KO and WT mice, as seen in Figure 2, which shows the KO/WT protein ratio for Ca_V2.1 (1.3 ± 0.4, n = 4) and Ca_V2.2 (1.0 ± 0.1, n = 3). In contrast, Ca_V3.2 expression was statistically lower among LVA channels (0.3 ± 0.09, n = 4) in the KLHL1 KO tissue (p = 0.04) whereas Ca_V3.1 and Ca_V3.3 expression remained constant (1.0 ± 0.2, n = 3; and 0.9 ± 0.06, n = 4 respectively). Thus, the absence of KLHL1 results in decreased Ca_V3.2 protein expression, which remains uncompensated for in the adult KLHL1 KO mice DRG.

To assess the physiological impact of lower Ca_V3.2 expression in the absence of KLHL1 we analyzed Ca²⁺ current densities in DRG neurons. We found two neuronal populations according to the Ca²⁺ currents they expressed (Figure 3A): 56% of all WT neurons elicited only HVA currents (capacitance = 23.0 ± 3.1 pF, n = 14; gray symbols); the remaining cells (44%) expressed both HVA and LVA currents. The latter group had an average capacitance of 18 pF ± 2.4 pF (black symbols, n = 11); this value was not statistically different from HVA-only neurons (p = 0.1).

Overall the HVA-only population was identical between WT and KLHL1 KO neurons (56% and 53% of the total population, respectively), with current densities of 45.1 ± 5.9 and 50.1 ± 7.9 pA/pF (n = 10 and 13 (WT, KO); p = 0.1). Figure 3B depicts an example of a recording from a neuron displaying HVA currents-only. Figure 3C shows the I-V curves for WT (black circles) and KO HVA-only neurons (white circles; n = 10, 13; p = 0.2).

Neurons expressing LVA+HVA currents represented 44% of the total population in WT vs. 47% in KO neurons (p = 0.1). Figure 3D depicts an example of HVA+LVA currents; the rapidly inactivating LVA current component is noticeable at lower voltages. Figure 3E shows an example of current traces recorded at V_T = −30 mV from V_H = −90 and from −50 mV, respectively, which when subtracted yield the LVA current component (red trace). Figure 3F shows the LVA-only current I–V curves from WT and KO DRGs. The peak LVA current was −37.0 ± 3.6 pA/pF (n = 11) in WT compared to −23.0 ± 4.6 pA/pF (n = 10) in the KO (at −30 mV; p = 0.004). We studied small neurons with capacitances ranging from 18 to 28 pF to ensure only nociceptor neurons were analyzed (Andrade et al., 2010), given that D-Hair cells also display a high density of T-type currents (Dubreuil et al., 2004; Coste et al., 2007; Bernal Sierra et al., 2017), but their capacitance ranges from ~39 to 65 pA (Coste et al., 2007).

The impact of decreased Ca_V3.2 channel expression on DRG neuron excitability was assessed using current clamp experiments. Figure 4A shows representative traces of the APs elicited by a depolarizing current ramp delivered at 60 pA/s. The KLHL1 KO neuron rheobase was significantly larger (39.0 ± 4.1 pA) than WT (22.1 ± 2.9 pA; n = 9, 11, p = 0.03; Figure 4B), in line with a reduction in LVA calcium channel expression. This increase was accompanied by a concomitant reduction in action potential number (AP; 9.2 ± 1.1, n = 9) compared to WT (13.1 ± 1.4, n = 11, p = 0.03; Figure 4C). This rheobase difference was abolished by application of a low dose (100 nM) of NCC 55–0396 (NCC) to partially block T-type channels in WT neurons (27.2 ± 3.7 pA, n = 8, p = 0.003).

Our data shows that Ca_V3.2 expression is down-regulated in the absence of KLHL1; KO DRG neurons display significantly lower neuronal excitability, which may, in turn, alter pain sensation. We assessed the responses to mechanical stimulation by measuring paw withdrawal thresholds in WT and KLHL1 KO mice using von Frey filaments. KLHL1 KO mice displayed significantly higher withdrawal threshold (6.1 ± 0.2 g, n = 20) compared to WT mice (4.7 ± 0.3 g, n = 20; p = 0.009; Figure 5A). Non-parametric analysis of paw withdrawal threshold responses within the mice population demonstrates statistical differences at 4 and 6 g of force (p < 0.05; Figure 5B). Thus, decreased Ca_V3.2 expression in KLHL1 KO DRG neurons results in decreased excitability and altered pain sensitivity, confirming KLHL1 is a physiological modulator of Ca_V3.2 in sensory neurons.

Induction of excitability changes by the manipulation of KLHL1 levels could represent a novel method in the regulation of Cav3.2 expression. Therefore, we tested whether knockdown of KLHL1 expression alters mechanical sensitivity in WT mice by injecting adeno-associated viral particles (AAV) containing shRNA designed against KLHL1 into the mice hind paws (US Patent 10,047,377).

Preliminary data from neuronal cultures indicated that titers ~5.0 × 10¹⁰ shRNA-containing viral particles appeared to be less efficient in vitro. Therefore, we carried out two blind trials assessing the effect of two titer viral loads (4.2 × 10¹⁰ vs. 9.0 × 10¹⁰ viral particles; Figure 6A). All trials were performed following the timeline showed in Figure 6B and described in “Materials and Methods” section.

Figure 6C shows representative images of DRG slices obtained from WT mice injected with EGFP-AAV or shKLHL1-AAV, confirming successful delivery and uptake of the AAV. Ca_V3.2 and KLHL1 levels from protein samples pooled from three L4 DRGs ipsilateral to the shKLHL1-AAV-injected paw were analyzed by western blot (sh). L4 DRG ipsilateral to the EGFP-AAV injection were also collected as a negative control (Ctrl; Liu et al., 2019).

Baseline behavioral tests were performed a week prior to injection in all mice (untreated). Blinded experimental measurements started 4–5 days after AAV injection and were performed 2–3 times weekly for 3 weeks thereafter (Figure 7). As seen in all figures, baseline withdrawal threshold values were indistinguishable in mice injected with either EGFP-AAV or shKLHL1-AAV at both titers, trial 1: EGFP-AAV, 5.2 g vs. 5.0 g for shKLHL1-AAV2; n = 7, p = 0.2; trial 2: EGFP-AAV, 5.2 g vs. 5.5 g in shKLHL1-AAV; n = 11, p = 0.2).

AAV injections caused some pain and inflammation, as expected (Ishihara et al., 2012), resulting in lower threshold values after injection compared with untreated values, as seen after injection in trial 1 at weeks 1–3 compared to untreated. Overall, the doses of shKLHL1 delivered in trial 1 exerted no effect on mechanical sensitivity, as seen in Figure 7A; the individual weekly averages were: week 1: EGFP-AAV, 4.4 ± 0.4 g vs. 4.1 ± 0.3 g for shKLHL1-AAV-2 (n = 7, p = 0.2); week 2: EGFP-AAV, 4.0 ± 0.2 g vs. 4.2 ± 0.2 g for shKLHL1-AAV-2 (n = 7, p = 0.3); and week 3: EGFP-AAV = 4.2 ± 0.2 g vs. 4.7 ± 0.1 g in shKLHL1-AAV-2 (n = 7, p = 0.056).

The dose delivered in trial 2 (~9.0 × 10¹⁰ viral particles, Figure 7B) induced significant differences in mechanical threshold values in shKLHL1-injected mice after 1 week. Unlike trial 1 and trial 2 control conditions, mechanical thresholds in shKLHL1-injected mice did not decrease compared to untreated conditions and they were significantly higher than their corresponding controls at all times tested. Week 1: EGFP-AAV, 3.6 ± 0.3 g vs. 4.6 ± 0.5 g for shKLHL1-AAV-1 (n = 11, p = 0.006); week 2: EGFP-AAV, 3.8 ± 0.5 g vs. 5.8 ± 0.3 g for shKLHL1-AAV-1 (n = 11, p = 0.006); and week 3: EGFP-AAV, 3.5 ± 0.4 g vs. 5.6 ± 0.5 g for shKLHL1-AAV-1 (n = 11, p = 0.001).

Further analysis is shown in Figure 8 where trial 2 data is shown as the percentage of the mice population displaying paw withdrawal threshold (PWT %) at a given Von Frey filament force value. There are no significant differences in baseline values (A) or after the first week after injection (B) between the experimental and control-treated mice populations (Kolmogorov–Smirnov test). However, 69% of EGFP-AAV injected mice responded to the 4 g von Frey filament stimulus 2 weeks after injection (C) in comparison with only 34% in the shKLHL1-AAV injected population (p < 0.05). This difference was more pronounced after 3 weeks of injection (D), with changes between the two populations at 4 g (35% shKLHL1-AAV vs. 64% EGFP-AAV) and 2 g (0% shKLHL1-AAV vs. 24% EGFP-AAV). Note that all mice were more sensitive after injections as a result of the AAV injections (compare to untreated).