DD04107 (Palmitoyl-EEQMRR-NH₂), DD04107^RDM (Palmitoyl-EQREMR-NH₂) and non-palmitoylated DD04107 (Ac-EEQMRR-NH₂) were kindly provided by BCN Peptides, Barcelona, and freshly prepared at 20 mM in H₂O. For microelectrode array, cells were incubated with 20 μM DD04107 in HBSS for 1 h (37°C, 5% CO₂). For electrophysiological recordings, non-palmitoylated DD04107 was prepared at 100 μM in standard internal solution from 10 mM stock solution in H₂O. Peptide or vehicle were applied through the patch pipette 10 min before recording. Capsaicin was dissolved in DMSO at 10 mM and further diluted in HBSS at 500 nM for MEAs and 1 μM for patch clamp experiments. BK was dissolved in H₂O at 1 mM and further diluted in HBSS at 1 μM. The BKR1 agonist Sar-[D-Phe⁸]-des-Arg⁹-BK and the BKR2 agonist [Phe⁸ψ(CH-NH)-Arg⁹]-BK (Tocris Bioscience, R&D Systems) were dissolved in H₂O at 1 mM and further diluted in HBSS at 1 μM. The BKR1 antagonist R715 and the BKR2 antagonist HOE140 (Tocris Bioscience, R&D Systems) were dissolved in H₂O at 1 mM and further diluted in HBSS at 1 μM. Unless indicated, all chemicals were obtained from Sigma-Aldrich.

All procedures were approved by the Institutional Animal and Ethical Committee of the University Miguel Hernández de Elche, in accordance with the guidelines of the Economic European Community, the National Institutes of Health, and the Committee for Research and Ethical Issues of the International Association for the Study of Pain. Animals were kept in a controlled environment (21–23°C, 12 h light/dark cycle), and had food and water available ad libitum. Neonatal Wistar rats and wild type C57BL/6J mice were purchased from in house bred stock (originally from Harlan Laboratories). Tac1-deficient mice (B6.Cg-Tac1^tm1Bbm/J) that do not express Substance P were purchased from The Jackson Laboratory. α-CGRP-deficient mice (B6;129-Calca^TM) were produced as described (Salmon et al., 1999).

DRG from neonatal Wistar rats (3–5 days-old) or adult male 12–15 weeks-aged mice (strains: C57BL/6J, αCGRP^-/-, Tac1^-/-), were cultured following previously described protocols with some modification (Baker and Bostock, 1997; Bonnington and McNaughton, 2003; Devesa et al., 2014). Briefly, neonatal rat ganglia were digested with 0.25% (w/v) collagenase (type IA) in DMEM-glutamax (Invitrogen) with 1% penicillin-streptomycin (P/S; 5000 U/mL, Invitrogen) for 1 h (37°C, 5% CO₂). Isolated mouse DRG were incubated with 0.67% (w/v) collagenase type XI and 3% (w/v) dispase (Gibco) in INC mix medium (in mM): 155 NaCl, 1.5 K₂HPO₄, 5.6 HEPES, 4.8 NaHEPES, 5 glucose) for 1 h (37°C, 5% CO₂). After digestion, rat and mouse DRGs were mechanically dissociated using a glass Pasteur pipette. Single cell suspension was passed through a 100 μm cell strainer, and washed with DMEM glutamax plus 10% fetal bovine serum (FBS; Invitrogen) and 1% penicillin/streptomycin. Cells were seeded at the required density for each experiment on 12 mm cover-glass slides, or microelectrode array chambers previously coated with poly-L-lysine (8.3 μg/mL) and laminin (5 μg/mL). After 2 h, medium was replaced with DMEM glutamax, 10% FBS and 1% P/S, supplemented with mouse 2.5S NGF 50 ng/mL (Promega) and 1.25 μg/mL cytosine arabinoside when required (37°C, 5% CO₂). Unless otherwise indicated, all experiments were made 48 h after cell seeding.

Whole-cell voltage clamp was made in neurons seeded on coverslips, placed in RC-25 chamber and connected to a external perfusion system at ≈22°C. External solution was (in mM): 140 NaCl, 4 KCl, 2 CaCl₂, 2 MgCl₂, 10 HEPES, 5 D-glucose, 20 mannitol, pH 7.4 (adjusted with NaOH). Internal pipette solution was (in mM): 144 KCl, 2 MgCl₂, 10 HEPES, 5 EGTA, pH 7.2 (adjusted with KOH). Membrane currents were acquired using EPC10 HEKA Patch amplifier (HEKA Electronics). Cells were held at a RMP of -60 mV, and at a sampling rate of 2.5 Hz. Patch glass pipettes with OD 1.5 mm × I.D. 1.17 mm (Harvard Instruments) were pulled with a Sutter Pippete puller Equipment (Sutter Instruments) to have a 3–6 MΩ resistance. Data acquisition and offline analysis were performed with PatchMaster software (HEKA Electronics). Rat DRG neurons were labeled with IB4-alexa 568 (10 μg/mL, 10 min, RT) in the external solution (Molecular Probes, Invitrogen), followed by two 5 min-washes. Labeled neurons were visualized through x20 air objective (Axiovert 200 inverted microscope, Carl Zeiss), with an excitation filter ET545 and an emision filter ET605 (CHR-49004, Laser 2000 SAD). Neuronal viabiliy was determined through typical neuronal Na⁺–K⁺ currents. TRPV1 desensitization was evoked by three repetitive 10 s-pulse of 1 μM capsaicin using a gravity-driven perfusion system (2 mL/min; Devesa et al., 2014). BK (1 μM) was applied for 8 min between P2 and P3 capsaicin pulses. Potentiation of TRPV1-mediated currents was calculated as the ratio of P3/P2 current peaks.

Electrical properties of the neuron were determined 2 min after establishing whole cell access using the current-clamp mode. For current-clamp recording, cells were held at 0 pA. Cells were assessed for the presence of spontaneous activity for 1 min and processed to measure electrogenic properties. Both IB4^- and IB4⁺ nociceptors were processed for electrogenic properties before BK application. Firing threshold was measured first by injecting a series of 100 ms depolarising current in 10 pA steps from 0 pA to elicit the first AP. To examine the neurons firing properties, a depolarising current of 40 pA (IB4^-) and 100 pA (IB4⁺) of 100 ms was injected to fire APs. APs were analyzed for the following intrinsic membrane properties: threshold potential (mV), amplitude of AP (mV), duration of AP (ms), overshoot of AP (mV), amplitude of afterhyperpolarization potential (mV). BK (1 μM) was applied to IB4^- and IB4⁺ nociceptors for 4 min to observe BK induced spontaneous neuronal excitabilities. RMP was checked before and after BK application. Mean depolarization was calculated by selecting the maximum depolarized voltage observed upon BK application in IB4^- and IB4⁺ nociceptors. Nociceptors (IB4^- and IB4⁺) were injected with 100 pA for 100 ms and 300 pA for 1 s to measure electrically evoked APs. BK induced changes in electrically evoked APs were measured before and after BK application.

Extracellular recordings were made using multiple electrode planar arrays of 60-electrode thin MEA chips, with 30 μm diameter electrodes and 200 μm inter-electrode spacing with an integrated reference electrode (Multichannel Systems GmbH). The electrical activity of primary sensory neurons was recorded by the MEA1060 System (Multi Channel Systems GmbH¹), and MC_Rack software version 4.3.0 at a sampling rate of 25 kHz. TRPV1-mediated neuronal firing activity was evoked by three repetitive 15 s-applications of 500 nM capsaicin, using continuous perfusion system (2 mL/min). BK (1 μM) in external solution was perfused between P2 and P3 for 8 min. For experiments on BK receptor agonists, selective BK receptor agonists for BKR1 and BKR2 were applied between P2 and P3 for 8 min. Data were analyzed using MC_RACK spike sorter and Neuroexplorer Software (Nex Technologies). An evoked spike was defined when the amplitude of the neuronal electrical activity overcame a threshold set at -20 μV. The recorded signals were then processed to extract mean spike frequency.

All data are expressed as mean ± SEM, with n as number of registered cells and N as the number of independent experiments. The percentage of sensitized neurons was calculated considering those cells that exhibited a fold potentiation (P3/P2) above 1.1. Statistical analysis was made using the Wilcoxon Rank test or Student’s t-test (unpaired or paired), one-way ANOVA or two-way ANOVA followed by Bonferroni’s post hoc test as indicated using the GraphPad Prism 5.0 (Graph-Pad). MEA data were analyzed by Bonferroni’s post hoc test as paired values through comparison of the responses of each electrode in the 30-s time interval upon stimulation. p < 0.05 was considered to be significant for a difference.

As a preliminary step, we characterized the effect of BK on the electrical properties of peptidergic (IB4^-) and non-peptidergic (IB4⁺) nociceptors. For this purpose, BK-induced changes in spontaneous neuronal activity, RMP and the properties of electrically evoked APs were investigated in primary cultures of neonatal rat nociceptors. For these experiments, IB4⁺ nociceptors were fluorescently labeled with Alexa-IB4, and nociceptor excitability was evaluated by whole cell patch-clamp. First, neurons were recorded for their RMP for 1 min after seal formation. Thereafter, these nociceptors were perfused with 1 μM BK for 4 min and an electrical response typically began within a minute of the pro-inflammatory agent application (Figure 1A). As seen, exposure of IB4^- nociceptors to BK produced a depolarization of the RMP that resulted in the firing of APs. In marked contrast, exposure of IB4⁺ neurons to BK did not result in a significant alteration of the RMP nor firing of APs. Quantification of the mean RMP reveals a more depolarized value for IB4^- nociceptors (-45 ± 2 mV) than for IB4⁺ neurons (-56 ± 1 mV; Figure 1B). Furthermore, despite of the variability inherent in primary nociceptor cultures, BK produced a significant depolarization in the mean RMP value (≈7 mV) of IB4^- nociceptors, indicating a direct effect of the pro-algesic agent in the excitability of peptidergic nociceptors.

Increase in nociceptor excitability or sensitization by inflammatory mediators has been associated with lower threshold requirement to fire APs and to an increased firing frequency in response to depolarizing stimuli (Gold and Traub, 2004). Next, we investigated the impact of BK exposure on electrically evoked APs in nociceptors. First, nociceptors (IB4^- and IB4⁺) were examined for evoked excitability in basal conditions upon injecting a depolarizing current pulse of 100 pA for 100 ms or of 300 pA for 1 s (Figure 2, vehicle). In both conditions, IB4^- nociceptors displayed significantly higher electrically evoked activity than IB4⁺ neurons. Second, 1 μM BK was applied for 4 min and nociceptors were retested for changes in evoked APs. Exposure of IB4^- nociceptors to BK depolarized the RMP, and increased the frequency of electrically activated APs by ≥1.5-fold, as evidenced for the two current injections (Figures 2A,C). Similarly, exposure of IB4⁺ nociceptors to BK also resulted in an increment in the electrically evoked AP firing, especially when 300 pA were applied (Figures 2B,D). Collectively, these findings indicate that peptidergic nociceptors are more excitatable and more reactive to acute BK sensitization than non-peptidergic nociceptors.

We next investigated the potentiating effect of BK on TRPV1 channels. For these experiments, we used the MEA technology to monitor the electrical activity of primary cultures of rat DRG neurons evoked by capsaicin. To register neuronal activity, a continuous protocol was used where two 15 s pulses of 500 nM capsaicin interspersed by a washing period of 3 min were applied, followed by an 8-min incubation with vehicle or 1 μM BK, and finalized by a third (P3) capsaicin pulse (Figures 3A,B). As seen, successive capsaicin pulses led to TRPV1 tachyphylaxia as evidenced by the lower AP firing evoked by the second (P2) and third (P3) pulses (Figure 3A). Application of BK after the second capsaicin pulse resulted in an increment of neuronal activity directly evoked by BK (Figure 3B). Notably, a subsequent capsaicin pulse (P3) produced a significant augment of the vanilloid response as compared with the second pulse (Figure 3B), consistent with a BK-induced potentiation of TRPV1 responses. Analysis of the mean spike frequency of each capsaicin pulse and the fold potentiation (estimated as the ratio P3/P2) substantiates this finding, revealing that BK increased the TRPV1 activity by 2.6 ± 0.4 fold (Figure 3E).

It has been documented that BK induces nociceptor potentiation mainly through BKR2 receptors, which are expressed constitutively in peptidergic nociceptors (Seabrook et al., 1997; Lee et al., 2002). In contrast, BKR1 receptors expression appears to be induced during inflammation in IB4⁺ non-peptidergic nociceptors (Davis and Perkins, 1994; Vellani et al., 2004; Petho and Reeh, 2012), with limited studies reporting a constitutive expression of BK1R receptors in nociceptors (Wotherspoon and Winter, 2000; Ma, 2001). Thus, the question that raises is which receptor isoform is mediating BK-induced TRPV1 sensitization in nociceptors. To explore the contribution of BKR1 and BKR2 receptors on BK induced TRPV1 sensitization, the BKR1 agonist Sar-[D-Phe⁸]-des-Arg⁹-BK) and the BKR2 agonist ([Phe⁸ψ(CH-NH)-Arg⁹]-BK) were used in an identical experimental paradigm as above. Figure 3C illustrates that BKR1 agonist modestly evoked neuronal firing and marginally promoted TRPV1-mediated neuronal spikes. In marked contrast and akin to BK, the BKR2 agonist notably increased nociceptor firing and strongly potentiated TRPV1 induced neuronal spiking (Figure 3D). Quantification of the mean spike frequency further substantiates the prominent effect of the BKR2 ligand as compared with that of BKR1. Notably, the fold potentiation of TRPV1-evoked neuronal spikes induced by BKR2 ligands (2.9 ± 0.3) is virtually identical to that evolved by BK (2.6 ± 0.4; Figure 3E) implying that BK exerts mainly its action through BKR2. In support of this tenet, no significant increase in the fold potentiation of TRPV1 evoked neuronal spikes was observed in neurons treated with the BKR1 agonist (Figure 3C). These results were further corroborated by using specific blockers of both receptor subtypes (Figure 3F). Collectively, these findings suggest that BK induces sensitization of TRPV1 mainly through activation of BKR2 in nociceptors.

Recent studies revealed that acute incubation of IB4^- nociceptors with BK induced the release of αCGRP in a dose-dependent manner through activation of the Gα_q/11 signaling pathway (Supowit et al., 2011). Our earlier findings showed that the algogen ATP, that also activates the G_q/11 signaling pathway, sensitizes TRPV1 in peptidergic nociceptors promoting the exocytosis of LDCVs carrying the receptor (Devesa et al., 2014). Henceforth, we hypothesized that BK may also induce TRPV1 sensitization through the exocytotic recruitment of new TRPV1 channels present in LDCVs.

To investigate if BK promotes the membrane mobilization of TRPV1 we evaluated the impact of inhibiting neuronal exocytosis of LDCVs using compound DD04107, a small lipidated peptide that interferes with the formation of the SNARE complex (Camprubí-Robles et al., 2009). For these experiments, we also used the MEA technology to monitor the electrical activity of primary cultures of rat DRG neurons (Devesa et al., 2014) utilizing an identical experimental protocol as above. In control cultures, two successive capsaicin pulses led to TRPV1 tachyphylaxia as evidenced by the lower AP firing evoked by the second (P2) pulse (Figure 4A). Application of BK after the second capsaicin pulse to cultures exposed to vehicle resulted in an increment of neuronal activity directly evoked by the kinin (Figure 4A, vehicle). A subsequent capsaicin pulse (P3) produced a significant augment of the vanilloid response as compared with the second pulse, consistent with a BK-induced potentiation of TRPV1 responses. Analysis of the mean spike frequency of each capsaicin pulse and the fold potentiation of these experiments (estimated as the ratio P3/P2) substantiates this finding, revealing that BK increased the TRPV1 activity in nociceptors by ≥2.0 fold (Figures 4B–D).

We next evaluated the effect of peptide DD04107 on the BK potentiation of capsaicin responses. For these experiments, primary cultures of rat nociceptors were pre-incubated with 20 μM DD04107 for 1 h. As a control, we used 20 μM compound DD04107^RDM, a peptide with the same amino acid composition but random sequence. Neither peptide affected the capsaicin nor BK responses as illustrated in Figure 4A (DD04107 and DD04107^RDM). As for control conditions, two consecutive pulses of the vanilloid-induced TRPV1 tachyphylaxia. Notably, exposure of nociceptors to 1 μM BK did resulted in a significantly lower potentiation of the response to the third capsaicin pulse in nociceptors incubated with peptide DD04107 as compared with those incubated with vehicle or DD04107^RDM. This is clearly discerned in the mean spike frequency and in the fold potentiation (Figures 4B–D). As illustrated in Figure 4, DD04107 notably inhibited the fold potentiation of vanilloid responses evoked by BK (Figure 4C). In contrast, DD04107^RDM did not affect the sensitizing effect of the kinin (Figure 4D). Though we observe a lower potentiation of TRPV1 activity by BK in DD04107-exposed neurons compared to vehicle and DD04107^RDM-treated nociceptors, we still notice potentiation of TRPV1 evoked excitability upon inhibiting neuronal exocytosis with DD04107 (Figures 4A,B). This result is consistent with the tenet that other mechanisms, apart from channel recruitment to the cell surface, are involved in BK-induced TRPV1 sensitization. Taken together, our findings imply that TRPV1 sensitization by BK in nociceptors is partially mediated by the exocytotic recruitment of new TRPV1 channels to the neuronal surface.

From our previous results, a question that emerges is whether BK-induced exocytotic mobilization of TRPV1 channels to the plasma membrane is a mechanism used by both types of nociceptors or by a specific subtype. Thus, we next examined the effect of peptide DD04107 on BK-induced TRPV1 sensitization in IB4^- and IB4⁺ nociceptors. For these experiments, IB4⁺ nociceptors were fluorescently labeled with Alexa-IB4, and TRPV1 channel activity was evaluated by whole cell patch-clamp. Figure 5A shows that capsaicin-induced TRPV1 desensitization in IB4^- nociceptors was strongly potentiated by 1 μM BK. Notably, BK-promoted TRPV1 sensitization in IB4^- sensory neurons was virtually abolished by 100 μM of the non-palmitoylated DD04107 peptide derivative delivered to the neuronal cytosol through the patch pipette. These results are corroborated upon calculating the fold potentiation of TRPV1 currents by taking the ratio of P3 and P2 (P3/P2; Figure 5E). No significant change in the fold potentiation was observed under control conditions for vehicle and peptide groups. However, the TRPV1 activity was ≈1.7-fold higher after BK incubation compared to control (Figure 5E). Noteworthy, the BK-induced increase in TRPV1 potentiation in IB4^- was fully blocked by the non-palmitoylated peptide DD04107 (Figures 5C,E), consistent with the tenet that BK sensitizes TRPV1 by promoting its mobilization to the neuronal surface.

At variance with IB4^- nociceptors, BK did not induce significant TRPV1 sensitization in IB4⁺ sensory neurons (Figure 5B). Consistent with the current recordings, there was no fold potentiation of TRPV1 evoked by BK in IB4⁺ nociceptors (Figure 5E) and, consequently, there was no effect of the non-palmitoylated DD04107 peptide (Figures 5D,E). Collectively, these results suggest that most of the TRPV1 sensitizing action produced by BK occurs in peptidergic nociceptors with a marginal contribution of the IB4⁺, non-peptidergic population. This is consistent with the finding that BK signals mainly through BKR2.

In a previous study, we reported that ATP-evoked TRPV1 potentiation in IB4^- peptidergic nociceptors was strictly dependent on the expression of αCGRP in these nociceptors (Devesa et al., 2014). Genetic ablation of αCGRP expression in mice fully abolished inflammatory recruitment of TRPV1 to the neuronal surface. Thus, we next evaluated whether BK-evoked TRPV1 sensitization also required the expression of pro-inflammatory peptides, αCGRP and/or SP, in peptidergic nociceptors. To address this question, we used DRG nociceptors from αCGRP and SP null mice, and monitored the electrical activity evoked with capsaicin using the MEA technology. We first examined the effect of BK sensitizing TRPV1 activity in mice nociceptors in culture. As illustrated in Figure 6A, BK potentiated capsaicin responses in mice nociceptors in a slightly lower extend than their counterpart from rat (Figures 3B and 6A). This is clearly discerned in the mean spike frequency (Figures 3E and 6C). Notice that BK also produced less spikes in mice nociceptors than in rat nociceptors. Furthermore, BK-evoked TRPV1-sensitization was highly sensitive to the inhibitory action of DD04107 (Figures 6B,C), indicating a similar sensitizing mechanism to that reported for rat nociceptors, namely recruitment of new channels to the neuronal surface.

Next, we performed the same measurements in nociceptors from αCGRP (αCGRP^-/-, Figure 7) and SP (Tac^-/-, Figure 8) null mice. In sensory neurons from αCGRP^-/- mice, BK was unable to potentiate TRPV1-evoked neuronal excitability (Figures 7A,B), similar to the blockade exerted with DD04107. In contrast, BK evoked strong TRPV1 sensitization in nociceptors from Tac^-/- mice (Figure 8). These results clearly imply that, akin to ATP, BK-promoted TRPV1 sensitization in IB4^- peptidergic nociceptors was dependent on the expression of αCGRP, and further imply that a population of TRPV1 is trafficked to the neuronal terminals in LDCVs loaded with αCGRP. Thus, pro-inflammatory mediators sensitize nociceptors by the concomitant release of αCGRP and the membrane recruitment of TRPV1 channels. This finding further corroborates that inflammatory sensitization of TRPV1 in peptidergic nociceptors involves the exocytotic delivery of new channels trafficked by LDCVs to the neuronal surface.