Nerve growth factor enhances the excitability of rat sensory neurons through activation of the atypical protein kinase C isoform, PKMζ

Abstract

Our previous work showed that nerve growth factor (NGF) increased the excitability of small-diameter capsaicin-sensitive sensory neurons by activating the p75 neurotrophin receptor and releasing sphingolipid-derived second messengers. Whole cell patch-clamp recordings were used to establish the signaling pathways whereby NGF augments action potential (AP) firing (i.e., sensitization). Inhibition of MEK1/2 (PD-98059), PLC (U-73122, neomycin), or conventional/novel isoforms of PKC (bisindolylmaleimide I) had no effect on the sensitization produced by NGF. Pretreatment with a membrane-permeable, myristoylated pseudosubstrate inhibitor of atypical PKCs (aPKCs: PKMζ, PKCζ, and PKCλ/ι) blocked the NGF-induced increase in AP firing. Inhibitors of phosphatidylinositol 3-kinase (PI3K) also blocked the sensitization produced by NGF. Isolated sensory neurons were also treated with small interfering RNA (siRNA) targeted to PKCζ. Both Western blots and quantitative real-time PCR established that PKMζ, but neither full-length PKCζ nor PKCλ/ι, was significantly reduced after siRNA exposure. Treatment with these labeled siRNA prevented the NGF-induced enhancement of excitability. Furthermore, consistent with the high degree of catalytic homology for aPKCs, internal perfusion with active recombinant PKCζ or PKCι augmented excitability, recapitulating the sensitization produced by NGF. Internal perfusion with recombinant PKCζ suppressed the total potassium current and enhanced the tetrodotoxin-resistant sodium current. Pretreatment with the myristoylated pseudosubstrate inhibitor blocked the increased excitability produced by ceramide or internal perfusion with recombinant PKCζ. These results demonstrate that NGF leads to the activation of PKMζ that ultimately enhances the capacity of small-diameter capsaicin-sensitive sensory neurons to fire APs through a PI3K-dependent signaling cascade.

nerve growth factor (NGF) plays an important role in initiation of the inflammatory response. Local inflammation can result in activation of small-diameter sensory neurons, which contributes to augmented sensitivity, vasodilatation, and plasma extravasation. The role of NGF is supported by observations that injection of NGF into a rat's paw produces elevated sensitivity to thermal and mechanical stimulation (Lewin et al. 1993). Pretreatment with an antibody to NGF prevents thermal hyperalgesia produced by injection of complete Freund's adjuvant into the rat paw (Lewin et al. 1994; Woolf et al. 1994). In an isolated skin-nerve-type preparation, NGF increased the firing frequency of isolated saphenous nerve in response to thermal stimulation (Rueff and Mendell 1996). More recent studies have demonstrated that NGF acts directly on sensory neurons because NGF augments capsaicin-evoked currents (Shu and Mendell 1999, 2001). In addition, our previous work demonstrated that NGF and brain-derived neurotrophic factor (BDNF) increase the number of current-evoked action potentials (APs) in capsaicin-sensitive small-diameter sensory neurons through modulation of voltage-dependent potassium currents (I_K) as well as the tetrodotoxin-resistant sodium current (TTX-R I_Na) (Zhang et al. 2002, 2008).

The intracellular signaling cascades activated by NGF that give rise to neuronal sensitization are poorly understood. Our work has demonstrated that NGF and BDNF augment excitability through activation of the p75 neurotrophin receptor (p75^NTR) and consequent downstream signaling by the sphingolipid pathway wherein the generation of ceramide appears to be a key event (Zhang et al. 2002, 2008; Zhang and Nicol 2004). Important early studies indicated that treatment with NGF led to increased neurite outgrowth in PC12 cells that was mediated by a phorbol ester-insensitive activation of an atypical PKC, PKCζ (Wooten et al. 1994, 1999). This observation raises the question as to how NGF leads to activation of PKCζ. Additional studies in a variety of cell types demonstrated that ceramide (ranging from membrane-permeant C2-ceramide to endogenous C18-ceramide) led to activation of PKCζ through a presently undefined mechanism (Bourbon et al. 2000; Lozano et al. 1994; Müller et al. 1995; Wang et al. 1999, 2005). Thus we speculate that NGF through activation of p75^NTR liberates ceramide, which in turn could lead to the activation of PKCζ. In these studies, we sought to establish the intracellular signaling pathways whereby NGF enhances the excitability of small-diameter sensory neurons with focus on the idea that NGF leads to the activation of this atypical PKC.

MATERIALS AND METHODS

Isolation and maintenance of adult rat sensory neurons.

The procedures for primary culture of rat sensory neurons have been described previously (Lindsay 1988) with slight modification (Jiang et al. 2003). Briefly, male Sprague-Dawley rats (100–150 g) were killed by being placed in a chamber that was then filled with CO₂. Dorsal root ganglia (DRG) were removed and collected in a culture dish filled with sterilized Puck's solution. The ganglia were transferred to a conical tube filled with Puck's solution containing 10 U/ml papain II and incubated for 12 min at 37°C. The tube was centrifuged for 50 s at low speed (∼2,000 g), and the pellet was resuspended in Puck's solution containing collagenase (1 mg/ml, type 1A) and dispase II (2.5 mg/ml). After a 14-min incubation at 37°C, the tube was centrifuged for 50 s before the enzyme-containing supernatant was removed. The pellet was resuspended in F-12 medium supplemented with 30 ng/ml 7S nerve growth factor (Harlan Bioproducts, Indianapolis, IN) and mechanically dissociated with fire-polished pipettes until all obvious chunks of tissues were gone. Isolated cells were plated onto plastic coverslips that had been previously coated with poly-d-lysine and laminin. The cells were maintained in F-12 medium containing NGF at 37°C and 3% CO₂ and were used within 12–24 h for electrophysiological recordings. All procedures were approved by the Animal Use and Care Committee of the Indiana University School of Medicine.

Electrophysiology.

Recordings were made using the whole cell patch-clamp technique as previously described (Hamill et al. 1981; Zhang et al. 2008). Briefly, a coverslip with the sensory neurons was placed in a recording chamber where the neurons were bathed in normal Ringer solution of the following composition (in mM): 140 NaCl, 5 KCl, 2 CaCl₂, 1 MgCl₂, 10 HEPES, and 10 glucose, with pH adjusted to 7.4 with NaOH. Recording pipettes were pulled from borosilicate glass tubing and fire-polished. Whole cell currents or voltages were recorded with an Axopatch 200 patch-clamp amplifier (Molecular Devices, Sunnyvale, CA); the data were acquired and analyzed using pCLAMP 6.04 or pCLAMP 9.0 (Molecular Devices). The whole cell recording configuration was established in normal Ringer solution. All drugs were applied by external superfusion of the recording chamber using a VC-8 bath perfusion system (Warner Instruments, Hamden, CT). In voltage-clamp experiments, both capacitance and series resistance compensation (typically 80%) were used. Linear leakage currents were subtracted by a P/4 protocol in the recordings of TTX-R I_Na but not in those of I_K. To isolate I_K, the Na⁺ in the normal Ringer solution was substituted with equimolar N-methyl glutamine (NMG). The membrane voltage was held at −60 mV; activation of the currents was determined by voltage steps of 350 ms applied at 5-s intervals in +10-mV increments from −80 to +60 mV. Patch pipettes had resistances of 2–5 MΩ when filled with the following solution (in mM): 140 KCl, 5 MgCl₂, 4 ATP, 0.3 GTP, 2.5 CaCl₂, 5 EGTA (calculated free Ca²⁺ concentration = 100 nM, MaxChelator), and 10 HEPES at pH 7.3 adjusted with KOH. This pipette solution was also used in the current-clamp recordings. The TTX-R I_Na was isolated by superfusing cells with the following Ringer solution (in mM): 30 NaCl, 65 NMG-Cl, 30 tetraethylammonium (TEA), 0.1 CaCl₂, 5 MgCl₂, 10 HEPES, 10 glucose, 10 sucrose, and 500 nM TTX, with pH adjusted to 7.4 with TEA-OH. The recording pipette was filled with (in mM) 110 CsFl, 25 CsCl, 10 NaCl, 5 MgCl₂, 4 ATP, 0.3 GTP, 1 CaCl₂, 10 EGTA, 10 glucose, and 10 HEPES at pH 7.3 (maintained with CsOH). Patch pipettes had resistances of 1–1.5 MΩ. Membrane voltage was held at −60 mV; activation of the current was determined by voltage steps 30 ms in duration and applied at 5-s intervals in +5-mV increments from −60 to +60 mV. After the control responses were obtained, the superfusate was changed to the appropriate Ringer solution and cells were superfused continuously for the appropriate times. The currents were filtered at 5 kHz, I_K was sampled at 1 kHz, and TTX-R I_Na was sampled at 25 kHz.

In the current-clamp experiments, the neurons were held at their resting potentials and a depolarizing ramp (1,000 ms in duration) was applied. The amplitude of the ramp was adjusted to produce between two and five APs under control conditions, and then the same ramp was used throughout the recording period for each individual neuron. Voltages were filtered at 5 kHz and sampled at 1 kHz. For experiments involving inhibitors of different signaling molecules, a set of sensory neurons from the same tissue harvest remained untreated as a positive control to assess the effects of NGF on neuronal excitability; parallel recordings were typically obtained from two to three individual untreated neurons. In all these recordings, NGF produced a two- to threefold increase in the number of evoked APs, demonstrating that NGF sensitized these neurons. The data from these parallel neurons (n = 10) have been combined and represent the effects of NGF on neuronal excitability for the untreated condition. These results are summarized in Table 3. At the end of each recording, the neuron was exposed to 400 nM capsaicin. This neurotoxin was used to distinguish capsaicin-sensitive sensory neurons because these neurons are believed to transmit nociceptive information (Holzer 1991). However, the correlation between capsaicin sensitivity and a neuron being a nociceptor is not absolute. Some nociceptive neurons are insensitive to capsaicin, and some capsaicin-sensitive neurons are not nociceptors (see Petruska et al. 2000). Therefore, this agent was used to define a population of small-diameter sensory neurons that could serve a nociceptive function. The results reported below were obtained from capsaicin-sensitive neurons only. All experiments were performed at room temperature (∼22°C).

Table 3.

Effects of NGF on excitability parameters

		Pretreatment
Parameters	Untreated	PD-98059	U-73122	PKCζ inhibitor
Control
RMP, mV	−59.2 ± 1.6	−58.2 ± 2.1	−58.6 ± 2.5	−55.5 ± 2.0
FT, mV	−16.1 ± 3.7	−17.3 ± 1.3	−19.3 ± 2.7	−27.3 ± 6.9
Rheo, pA	440 ± 152	350 ± 81	709 ± 383	382 ± 201
RTh, MΩ	196 ± 48	161 ± 38	198 ± 74	183 ± 60
n	10	7	7	5
NGF (6 min)
RMP, mV	−54.1 ± 1.7*	−50.1 ± 1.6*	−54.3 ± 2.4*	−56.5 ± 2.1
FT, mV	−18.3 ± 3.2	−14.7 ± 5.8	−22.7 ± 2.4	−25.1 ± 7.0
Rheo, pA	212 ± 65*	169 ± 21*	368 ± 175*	385 ± 203
RTh, MΩ	268 ± 48*	214 ± 21	260 ± 101	202 ± 61
Norm Rheo	0.59 ± 0.08*	0.60 ± 0.09*	0.58 ± 0.08*	1.06 ± 0.08†
Norm RTh	1.66 ± 0.29*	1.77 ± 0.43	1.43 ± 0.13*	1.12 ± 0.06†

Values are means ± SE; n = no. of neurons. RMP, resting membrane potential; FT, firing threshold; Rheo, rheobase; R_Th, membrane resistance at threshold; Norm Rheo, rheobase NGF/rheobase control; Norm R_Th: resistance NGF/resistance control.

^*P < 0.05, untreated/pretreatment control vs. NGF (6 min) (within-groups RM ANOVA).

^†P < 0.05, untreated + NGF vs. pretreatment + NGF (across groups, ANOVA).

Small interfering RNA.

To reduce expression of PKCζ, we used a previously described small interfering RNA (siRNA) treatment protocol (Chi and Nicol, 2007). Briefly, isolated sensory neurons were maintained for 6 h in normal medium with 30 ng/ml NGF. Normal medium was replaced with Optimem medium overnight. The Metafectene-siRNA complex (200 nM) was added on day 2 in culture wherein the neurons were exposed to the siRNA or Metafectene alone for 48 h at 37°C. After 2 days (day 4 in culture), the Metafectene ± siRNA was washed out and the normal medium containing antibiotics and NGF was then added to the neurons and allowed to incubate for another 2 days before electrophysiological recordings, Western blots, or quantitative real-time PCR (qPCR) were performed. The siRNA targeted to PKMζ/PKCζ (NCBI reference sequence NM_022507.1 for PKCζ) was a mixture of four siRNAs (siGenome SMARTpool, M-09-1560-00) obtained from Thermo Scientific (Lafayette, CO), all of which are targeted to sequences present in both PKMζ and PKCζ mRNAs. The sequences targeted by this siRNA pool (5′ to 3′) are as follows: siRNA1, GAACGAUGGUGUAGACCUU (position 741–759); siRNA2, GGAAACAUGACAAUAUCAA (position 803–821); siRNA3, GCUGAGAUCUGUAUCGCUC (position 1218–1237); and siRNA4, CGAUGCCGAUGGACACAUU (position 1299–1310). Two additional siRNA molecules were obtained from Thermo Scientific: siRNA 638, GCAAGCTGCTTGTCCATAA (5′ to 3′, position 638–656), targeted to sequence present in both species, and siRNA 407, GGGACGAAGTGCTCATCAT (5′ to 3′, position 407–425), targeted to sequence present in PKCζ but not PKMζ. The negative control siRNA was obtained from Ambion (SC-1; Austin, TX) and had the sequence (5′ to 3′) GCGCGCUUUGUAGGAUUCG. Metafectene was purchased from Biontex-USA (San Diego, CA). For the electrophysiological studies, the siRNA and the negative control siRNA were labeled using the Mirus Bio Label IT* siRNA Tracker* intracellular localization kit (fluorescein) available from Fischer Scientific (Pittsburgh, PA).

Western blot.

All procedures were conducted on ice. Isolated sensory neurons maintained in culture were scraped from the dish after addition of 150 μl of ice-cold 1× RIPA lysis buffer (no. 20-188; Millipore, Billerica, MA) containing a 100-fold dilution of Protease inhibitor cocktail set III (539134; EMD Biosciences, San Diego, CA). Cell lysates were transferred to 1.5-ml tubes and sonicated (setting 3 of Fisher Scientific Sonic Dismembrator 550, 2 pulses, 1 s each pulse). Lysates were centrifuged at 2,500 g for 4 min at 4°C. Supernatants were transferred to fresh tubes. Protein concentration was measured using the Bradford method (protein assay dye reagent no. 500–000; Bio-Rad, Hercules, CA). Equivalent amounts of reduced, denatured protein (25 μg) were loaded and separated on a NuPAGE 4–12% Bis-Tris gel (NP0335BOX; Invitrogen, Carlsbad, CA) using NuPAGE MES-SDS running buffer (NP0002; Invitrogen) and then transferred to an Invitrolon polyvinylidene difluoride membrane (LC2005; Invitrogen). To measure expression levels, we used the following antibodies: an atypical PKC polyclonal rabbit antibody that detects PKMζ/PKCζ and PKCλ/ι (sc-216, 1:500; Santa Cruz Biotechnology, Santa Cruz, CA), a PKCζ-specific rabbit polyclonal antiserum (1:5,000; described in Hernandez et al. 2003), a PKCλ/ι-specific monoclonal mouse antibody (610207, 1:500; BD Transduction Laboratories, San Diego, CA), and for a loading control, an actin monoclonal mouse antibody (MS-1295-PABX, 1:2,500; Thermo Scientific, Waltham, MA). These antibodies were added to the blocking solution and incubated overnight at 4°C with agitation. After serial incubation with specific secondary antibodies, immunoreactive bands on the membrane were developed using enhanced chemiluminescence (ECL kit; Thermo Scientific, Rockford, IL) and visualized by exposure to Blue Lite Autorad film (F-9024–8 × 10; ISCBioExpress, Kampenhout, Belgium). The film was photographed with a Kodak DC290 charge-coupled device camera, and the density of each band was measured using Kodak 1D 3.6 software (Kodak Scientific Imaging Systems, New Haven, CT). The summary data are expressed as the density of the experimental bands normalized to the density of their respective actin bands and then normalized to their respective untreated control/actin values.

Quantitative real-time PCR.

DRG, brain, and lung tissue were isolated from male Sprague-Dawley rats (100–150 g). Total RNA was extracted from each tissue and from primary cultures of dissociated DRG neurons (isolated in the absence of papain) using the RNeasy Plus minikit (Qiagen, Valencia, CA) and assessed on a NanoDrop ND-1000 spectrophotometer (Thermo Scientific, Franklin, MA) for concentration (A₂₆₀) and purity by optical density (OD) ratios (A₂₆₀/A₂₈₀, between 2.0 and 2.2). RNA integrity was also assessed using the Experion RNA StdSens analysis kit and Experion automated electrophoresis system (Bio-Rad). RNA samples with a RNA quality indicator value >7.0 were used for this study. After treatment with DNase I (Invitrogen), 1 μg of RNA was reverse transcribed using the iScript cDNA synthesis kit (Bio-Rad) in 20-μl reactions according to the manufacturer's instructions. To assess the specificity of these reactions, no-reverse transcriptase and no-template controls were also generated. Reverse transcription products were diluted with nuclease-free water and amplified in 10-μl reactions using Power SYBR Green PCR Master Mix (Applied Biosystems, Carlsbad, CA) and 500 nM forward and reverse primers (Invitrogen). Two sets of primers were used for each aPKC gene target: one set for PKCζ and PKMζ was designed in our laboratory using PrimerExpress software v3.0 (Applied Biosystems), whereas the primer set for PKCλ/ι was based on Stretton et al. 2010. The second set of sequences was kindly provided by Dr. Sourav Ghosh and his laboratory (Cellular and Molecular Medicine, University of Arizona); these were targeted to aPKC mouse transcripts wherein we modified several base pairs to match rat mRNA sequences. Acidic ribosomal protein P0 (Arbp) was selected as the reference gene for normalization. The accession number, amplicon size and position, and primer sequences for all genes targeted in this study are shown in Table 1. qPCR reactions were run in duplicate with tissue-derived cDNA or in triplicate with primary DRG cell culture cDNA on an Applied Biosystems 7500 Fast Real-Time PCR system using MicroAmp Fast 96-well reaction plates sealed with MicroAmp optical adhesive film. All reactions began with an initial cycle of 95°C for 10 min, followed by 40 cycles of 95°C for 15 s and 60°C for 1 min. Efficiencies for each primer pair were determined from the fitted slope of a seven-point standard curve. The regression fit was considered to be acceptable when r² > 0.99. Calculated efficiencies for all primers are listed in Table 2. The specificity of these amplifications was verified by melt curve analysis and electrophoresis on a 2% agarose gel. The quantification cycle (Cq) was chosen to be number of cycles when the value of normalized fluorescence generated by SYBR green emission attained 0.3. For determination of the expression levels of PKCζ, PKMζ, PKCλ/ι, and Arbp, a calculation based on Pfaffl (2001) was used wherein the number of copies = efficiency^−Cq. Because the calculated efficiencies listed in Table 2 are all very close to 2, we have based the calculation of number of copies for all genes on an efficiency of 2.

Table 1.

Primer sequences

Gene	Ref. Sequence	Sequence (5′ to 3′)	Location	Size, bp
PKCζ	NM_022507.1	F: TGGACCCCACGACAACTTTC	239–258	178
		R: ACTTCGTCCCTGCCCTGAC	416–398
PKMζ	S80195.1	F: CCTTCTATTAGATGCCTGCTCTCC*	50–73	88
		R: CCTCCGGACCGACAATGG	137–120
PKCλ/ι	NM_032059.1	F: CCATGATGCCAATGGACCAGTC†	737–758	136
		R: TTCCCACTCTCCCTGGTGTTC†	872–852
Arbp	NM_022402.2	F: CAGCCAAGGTCGAAGCAAA	957–975	58
		R: CCGAATCCCATGTCCTCATC	1014–995
Ghosh-adapted primers
PKCζ		F: GTCAGGGCAGGGACGAAGT	398–416	136
		R: TGGCTCGGTATAGCTTCCTCC	532–512
PKMζ		F: AGCAGAGAAAGCCGAGTCCA	233–252	96
		R: TTAAAGCGCTTGGCTTGGAA	328–309
PKCλ/ι		F: CCGTGTGTACCAGAGCGTCC	460–479	106
		R: TGTGGCCATTTGCACAATACA	565–545

Adapted primers were provided by Dr. Sourav Ghosh, University of Arizona (see text).

^*From Hernandez et al. 2003 (forward only).

^†From Stretton et al. 2010.

Table 2.

Calculated primer efficiencies

	Our Primers	Ghosh-Adapted Primers
Brain
PKCζ	1.981	1.999
PKMζ	2.058	2.035
PKCλ/ι	1.975	2.013
Arbp	1.998
Lung
PKCζ	2.006	1.974
Arbp	2.019

Internal perfusion.

Recombinant PKCζ and PKCι were aliquoted and stored according to the manufacturer's instructions (PKCζ: Cell Signaling Technology, Beverly, MA; PKCζ and PKCι: ProQinase, Freiburg, Germany). PKCζ was dissolved in the pipette solution at a concentration of 2 ng/μl, and PKCι at an initial concentration of 0.42 ng/μl. At these concentrations, the specific activities were similar. The tip of the patch pipette was filled with pipette solution; the pipette was then backfilled with pipette solution containing recombinant PKCζ or PKCι. A fresh batch of recombinant PKC-pipette solution was made for each recording session and kept on ice during the experiments. Control recordings were obtained as soon as possible after the whole cell configuration was established. A concentration of 2 ng/μl was chosen based on the dose-activity relation for the recombinant PKCζ wherein 100 ng/50 μl reaction yielded maximal kinase activity (see Fig. 3, http://www.cellsignal.com/pdf/7608.pdf). Recombinant PKCζ (2 ng/μl in pipette solution) was heat inactivated by being placed in boiling water for 10 min and then immediately placed on ice for the remainder of the recording session.

Fig. 3.

A pseudosubstrate peptide inhibitor of the atypical PKCζ blocks the sensitization produced by NGF. A: a 30-min pretreatment with 1 μM bisindolylmaleimide I (BIM) did not affect the increased number of evoked APs produced by bath application of 100 ng/ml NGF (n = 5). B: a 30-min pretreatment with 10 μM myristoylated pseudosubstrate inhibitor (PS-Inhib PKCζ via the bath, n = 5) completely blocked the sensitization produced by 100 ng/ml NGF. C: treatment with 1 μM C2-ceramide (C2-Cer) significantly increased the number of evoked APs by 2-fold (n = 3) over 20 min. D: a 30-min pretreatment with 10 μM PS-Inhib PKCζ blocked the increased excitability produced by C2-Cer (n = 5). E: a 30-min pretreatment with 10 μM of the myristoylated pseudosubstrate inhibitor for PKCη (PS-Inhib PKCη) had no effect on the baseline levels of excitability (no NGF, n = 3) and did not alter the increased excitability produced by NGF (n = 6). F: PS-Inhib PKCζ (n = 6) had no effect on the increased excitability produced by bath application of 1 μM PGE₂. *P < 0.05 compared with the respective control values (RM ANOVA).

Data analysis.

Data are means ± SE. The parameters of excitability described in Tables 3 and 4 were determined, in part, by differentiating the voltage trace (dV/dt) in the current-clamp recordings (sampling frequency of 500 Hz). The firing threshold and time at which the first AP was fired were taken as the point that exceeded the baseline value of dV/dt by >20-fold. The baseline value of dV/dt was determined by averaging the points between the onset of the ramp and the next 100 ms (135–235 ms). The rheobase was measured as the amount of ramp current at the firing threshold. The resistance at threshold (R_Th) was calculated as the difference between the firing threshold and the resting membrane potential divided by the rheobase current. The voltage dependence for activation of I_K and TTX-R I_Na was fitted with the Boltzmann equation, where G/G_max = 1/[1 + exp(V_0.5 − V_m)/k], in which G is the conductance [G = I/(V_m − E_rev)], G_max is the maximal conductance obtained from the Boltzmann fit under control conditions, E_rev is the reversal potential for the targeted current, V_0.5 is the voltage for half-maximal activation, V_m is the membrane potential, and k is a slope factor. The Boltzmann parameters were determined for each individual neuron and were used to calculate the mean ± SE. Statistical differences between the control recordings and those obtained under various treatment conditions were determined by using either an ANOVA or repeated-measures (RM) ANOVA. When a significant difference was obtained with ANOVA, post hoc analyses were performed using a Holm-Sidak all pairwise test. If the data set failed the normality test, a Kruskal-Wallis analysis of variance on ranks was performed using Dunn's all pairwise test. Values of P < 0.05 were judged to be statistically significant.

Table 4.

Effects of recombinant PKCζ on excitability parameters

	Control (0 min)	PKCζ (6 min)
No pretreatment
RMP, mV	−64.0 ± 2.5	−57.4 ± 1.2*
FT, mV	−10.5 ± 2.4	−19.9 ± 1.6*
Rheo, pA	727 ± 138	226 ± 84*
RTh, MΩ	100 ± 22	189 ± 24*
Norm Rheo	1.0	0.37 ± 0.06*‡
Norm RTh	1.0	2.18 ± 0.28*
Pretreatment with Myr-PKCζ inhibitor
RMP, mV	−57.0 ± 2.4	−57.5 ± 2.1
FT, mV	−29.1 ± 3.7	−28.3 ± 3.6
Rheo, pA	317 ± 270	306 ± 252
RTh, MΩ	651 ± 211	620 ± 215
Norm Rheo	1.0	1.13 ± 0.07†
Norm RTh	1.0	0.96 ± 0.05†

Values are means ± SE for 6 neurons. Myr, myristoylated.

^*P < 0.05, control vs. PCKζ (6 min) (RM ANOVA).

^†P < 0.05, PKCζ (6 min) vs. PKCζ + inhibitor (ANOVA).

^‡P < 0.05, PKCζ (6 min) vs. untreated + NGF (6 min) (ANOVA; cf. Table 3).

Chemicals.

Bisindolylmaleimide I (BIM), PD-98059, U-73122, LY-294002, LY-303511, wortmannin, and pseudosubstrate inhibitor peptides for PKCζ and PKCη were obtained from EMD Chemicals. Purified recombinant human PKCι was purchased from ProQinase, and PKCζ was obtained from Cell Signaling Technology and ProQinase. C2-ceramide was obtained from Avanti Polar Lipids (Alabaster, AL). Tissue culture supplies were purchased from Invitrogen. All other chemicals were obtained from Sigma Chemical (St. Louis, MO). Capsaicin, BIM, PD-98059, U-73122, LY-294002, LY-303511, and wortmannin were dissolved in 1-methyl-2-pyrrolidinone (HPLC grade). The stock solutions were then diluted with Ringer to yield the appropriate concentration. We have previously demonstrated that the vehicle 1-methyl-2-pyrrolidinone had no effect on AP firing or the activation of either TTX-R I_Na or I_K (Zhang et al. 2002, 2008).

RESULTS

NGF enhances the excitability of small-diameter capsaicin-sensitive sensory neurons.

As we previously demonstrated, exposure to NGF augments the excitability of small-diameter capsaicin-sensitive neurons (Zhang et al. 2002). Bath application of NGF to different sets of untreated sensory neurons used in parallel experiments involving the inhibitors of different signaling cascades (described below) produced a significant increase in the number of evoked APs (control, 3.8 ± 0.3 APs vs. NGF, 9.5 ± 1.2 APs after 6 min, n = 10, RM ANOVA, data not shown). From these 10 neurons, the effects of NGF on different parameters of excitability were assessed. Although NGF did not alter the firing threshold, there were significant changes in resting membrane potential, rheobase, and R_Th. The results from these parallel neurons represent the effects of NGF on neuronal excitability for the untreated condition and are summarized in Table 3.

Neither ERK nor PLC pathways are involved in NGF sensitization.

In this report we sought to establish the downstream signaling pathways that contribute to the increased excitability. Therefore, to determine whether ERK/MAPK played a role in increased AP firing produced by NGF, neurons were exposed to PD-98059, a selective inhibitor of MEK1/2 (IC₅₀ 4 μM; Alessi et al. 1995; English and Cobb 2002). As shown for a representative recording (Fig. 1, A and B), bath application of NGF (100 ng/ml) to an untreated neuron greatly increased the number of APs evoked by a depolarizing ramp of current. In three untreated sensory neurons, NGF significantly increased the number of evoked APs by greater than twofold (see Fig. 1C, RM ANOVA). A parallel set of isolated neurons was pretreated with 10 μM PD-98059 for 30 min and then exposed to NGF. PD-98059 did not affect the capacity of NGF to increase the number of evoked APs in small-diameter sensory neurons (n = 7) compared with NGF alone. These results are summarized in Fig. 1C. Pretreatment with PD-98059 had no significant effects on the NGF-induced changes in the parameters of excitability, which are summarized in Table 3. Zhu and Oxford (2007) demonstrated that the NGF-induced phosphorylation of ERK (p42/p44) was greatly reduced by treatment with 10 μM PD-98059 (see their Fig. 4), indicating that this concentration was sufficient to block activation of MEK1/2. In addition, pretreatment with 10 μM PD-98059 alone for 30 min did not alter the number of evoked APs (3.3 ± 0.5 APs at 0 min vs. 3.3 ± 0.5 APs after 20 min, n = 4, RM ANOVA, data not shown) or membrane potential (−54.8 ± 2.4 at 0 min vs. −55.4 ± 2.5 mV after 20 min) over time.

Fig. 1.

PD-98059 (PD), a selective inhibitor of MEK1/2, does not block the enhanced excitability produced by nerve growth factor (NGF). A and B: representative recording of action potential (AP) firing in response to the depolarizing ramp of current wherein under control conditions (A, untreated), 5 APs were evoked; after a 6-min bath application of NGF (B, 100 ng/ml), 9 APs were generated. C: summary of the effects of NGF on AP firing obtained from a parallel group of untreated control neurons (n = 3) and another group that were pretreated with PD (n = 7). Cont, control. *P < 0.05 compared with the respective control groups (untreated and PD) [repeated-measures (RM) ANOVA].

Fig. 4.

Inhibition of phosphatidylinositol 3-kinase (PI3K) blocks increased AP firing produced by NGF. Neurons were untreated or pretreated with 10 μM LY-294002 or LY-303511 for 30 min before bath superfusion with 100 ng/ml NGF (n = 6 for NGF alone, n = 12 for LY-294002, and n = 7 for LY-303511). Although not shown, 100 nM wortmannin inhibited the NGF-induced increase in AP firing (n = 7). *P < 0.05 compared with the respective control values (RM ANOVA). In an all pairwise multiple comparisons (ANOVA, Holm-Sidak test), the 4 control values were not significantly different (untreated, LY-294002, wortmannin, and LY-303511). For untreated sensory neurons, the NGF-induced increase in AP firing at all time points (2, 6, and 10 min) was significantly different from that for the untreated control, the LY-303511 control, and all time points for LY-294002 and wortmannin. For neurons pretreated with LY-303511, the NGF-induced increase in AP firing at the 6- and 10-min time points was significantly different from that for the untreated and LY-303511 controls and all time points for LY-294002 and wortmannin. There were no differences at 2, 6, and 10 min for the NGF-induced increase in AP firing for untreated and LY-303511-pretreated neurons.

To examine the role of PLC in the NGF-induced increase in AP firing, sensory neurons were exposed to U-73122, a selective inhibitor of PLC (IC₅₀ 1–2 μM; Bleasdale et al. 1990; Smith et al. 1990). U-73122 also blocks activation of PLCγ-1, including that produced by NGF (Jones et al. 2005; Sato et al. 2000; Zapf-Colby et al. 1999). As shown in Fig. 2, pretreatment with 5 μM U-73122 for 30 min did not affect the capacity of NGF to augment the number of evoked APs (n = 8) and was similar to the NGF-induced sensitization observed in a parallel set of neurons in the absence of U-73122 (n = 5, RM ANOVA). Pretreatment with U-73122 had no significant effects on the changes in the parameters of excitability produced by NGF alone, which are summarized in Table 3. Pretreatment with 5 μM U-73122 (30 min) alone did not alter the number of evoked APs (3.3 ± 0.8 APs at 0 min vs. 3.5 ± 0.9 APs after 20 min, n = 4, RM ANOVA, data not shown) or membrane potential (−53.8 ± 1.1 at 0 min vs. −51.8 ± 2.0 mV after 20 min) over time. Although U-73122 had no effect on the capacity of NGF to increase AP firing, recent work has raised questions regarding the ability of U-73122 to inhibit PLC specifically (see Horowitz et al. 2005; Klein et al. 2011). To corroborate whether PLC plays an important role in the NGF-mediated increase in excitability, sensory neurons were pretreated with 30 μM neomycin, which can be an effective inhibitor of PLC (Lipsky and Lietman 1982; Schwertz et al. 1984), for 30 min before recording. Pretreatment with neomycin also failed to block the increased AP firing produced by NGF. Under control conditions the ramp evoked 3.8 ± 0.4 APs; exposure to 100 ng/ml NGF significantly increased the number of APs to 7.6 ± 0.9, 8.2 ± 0.6, 7.6 ± 0.6, and 8.3 ± 0.3 APs after exposures of 2, 6, 10, and 20 min, respectively (n = 5 with 1 neuron lost between 10 and 20 min, ANOVA, Holm-Sidak posttest, data not shown). The NGF-induced increase in AP firing in the presence of neomycin for all time points (2 through 20 min) was not significantly different from the increased firing obtained for the untreated control sensory neurons, pretreatment with U-73122, and pretreatment with PD-98059 (P = 0.98, ANOVA, all pairwise). Together, these results indicate that neither activation of ERK nor PLC contributed to the elevated excitability produced by NGF.

Fig. 2.

U-73122, a selective inhibitor of PLC, does not suppress the increased excitability produced by NGF. Pretreatment with 5 μM U-73122 for 30 min did not block the increased AP firing produced by bath application of 100 ng/ml NGF (U-73122 + NGF, n = 8). The increased firing by NGF was similar to that observed in a parallel set of neurons in the absence of U-73122 (n = 5, untreated condition). *P < 0.05 compared with the respective control values (RM ANOVA).

NGF sensitization involves activation of an atypical PKC but not conventional or novel PKC subtypes.

To examine whether PKC played a role in increased excitability produced by NGF, neurons were treated with BIM. After 30-min pretreatment with 1 μM BIM, neurons exposed to NGF (100 ng/ml) still produced a significant increase in the number of APs (Fig. 3A, n = 5, RM ANOVA). The NGF-induced increase in AP firing after pretreatment with BIM was not different at any of the time points compared with the increases produced by NGF in the untreated neurons and the PD-98059-, U-73122-, and neomycin-pretreated neurons (P = 0.79, ANOVA). However, at a concentration of 1 μM, BIM inhibits only conventional (IC₅₀ 10–30 nM) and novel (IC₅₀ 100–200 nM), but not atypical (IC₅₀ ∼6 μM), isoforms of PKC (Martiny-Baron et al. 1993; Toullec et al. 1991). Because pretreatment with 1 μM BIM did not block the sensitizing actions of NGF, it is possible that NGF modulated excitability through activation of an atypical PKC.

Our previous work showed that NGF activation of p75^NTR enhanced neuronal excitability through the sphingomyelin/ceramide pathway (Zhang et al. 2002; Zhang and Nicol, 2004). Studies using other cellular model systems have demonstrated that ceramide can activate the ζ-subtype of atypical PKC (PKCζ) (Bourbon et al. 2000; Lozano et al. 1994; Müller et al. 1995; Wang et al. 1999, 2005). To examine the potential role of NGF-mediated activation of PKCζ, neurons were exposed via the bath to a selective peptide inhibitor targeted to the pseudosubstrate domain of aPKCs (EC₅₀ 10–20 μM; Standaert et al. 1997). As shown in Fig. 3B, a 30-min pretreatment with 10 μM myristoylated pseudosubstrate inhibitor completely blocked the sensitization produced by NGF. Furthermore, in the presence of the PKCζ inhibitor, NGF failed to alter the resting membrane potential, the rheobase, or R_Th (see Table 3). In an all pairwise comparison, the NGF-induced reduction in the normalized values for the rheobase were significantly different for untreated neurons and for pretreatment with PD-98059 and U-73122 compared with pretreatment with the pseudosubstrate inhibitor (P = 0.004, ANOVA, Holm-Sidak all pairwise test), although the increase for the normalized values of R_Th was not different between these groups (P = 0.27, ANOVA on ranks). In addition, internal perfusion of the nonmyristoylated PKCζ pseudosubstrate inhibitor (although at a higher concentration, 60 μM) via the recording pipette blocked the NGF-induced increase in excitability wherein 4.0 ± 0.3 APs were evoked under control conditions compared with 4.0 ± 0.3 APs after a 20-min exposure to 100 ng/ml NGF (n = 6, P = 0.93, ANOVA, data not shown). In further studies, exposure of sensory neurons to 1 μM C2-ceramide produced a significant twofold increase in the number of evoked APs (Fig. 3C) and is similar to our previously reported results wherein there was a threefold increase in the number of APs generated after a 20-min exposure to 1 μM C2-ceramide (Zhang et al. 2002). In a parallel set of neurons, pretreatment with the PKCζ pseudosubstrate inhibitor blocked the increased excitability produced by C2-ceramide (Fig. 3D). Consistent with the results obtained for NGF, treatment with the inhibitor prevented the ceramide-induced changes in the resting membrane potential, the decrease in normalized rheobase, and the increase in normalized R_Th (data not shown). Previous studies have indicated that myristoylation of peptides can lead to the activation of eNOS in endothelial cells through the Akt pathway (Krotova et al. 2006). Therefore, as a control experiment for the myristoylated peptide, neurons were pretreated for 30 min with 10 μM of the pseudosubstrate inhibitor for the novel η-isoform of PKC, PKCη. Treatment with the PKCη inhibitor did not alter the increased excitability produced by NGF or the stability of AP firing over a 20-min period in the absence of NGF (Fig. 3E). The NGF-induced increase in AP firing after pretreatment with the PKCη inhibitor was not different from the NGF-mediated increase in untreated sensory neurons (P = 0.40, ANOVA). As a positive control, treatment with the myristoylated pseudosubstrate inhibitor of aPKCs (60 μM for 30 min, 5 times the EC₅₀ value) had no effect on the increased excitability produced by bath application of 1 μM PGE₂ (Fig. 3F). These results indicate that the PKCζ pseudosubstrate inhibitor lacked any nonspecific actions on the capacity of these neurons to be sensitized, since it is well established that PGE₂ acts through the G_s-cyclic AMP-PKA pathway (Cui and Nicol 1995; Hingtgen et al. 1995; Taiwo et al. 1989). Thus these observations indicate that ceramide is a key upstream modulator of PKCζ activity that can alter neuronal excitability.

In addition to ceramide, previous studies have demonstrated that phosphatidylinositol 3-kinase (PI3K) is a key upstream regulator of PKCζ activity (Chou et al. 1998; Herrera-Velit et al. 1997; Mendez et al. 1997; Standaert et al. 1997). To determine whether PI3K had any role in the NGF-induced increase in excitability, sensory neurons were pretreated with 10 μM LY-294002, a selective PI3K inhibitor (IC₅₀ 1.4 μM; Vlahos et al. 1994), for 30 min before recording. As shown in Fig. 4, LY-294002 suppressed the ability of NGF to augment the number of evoked APs. Similar results were obtained with exposure to 100 nM wortmannin, a structurally different PI3K inhibitor (IC₅₀ 5 nM; Powis et al. 1994) (control, 3.4 ± 0.5 APs vs. 10 min of NGF, 4.0 ± 0.5 APs, n = 7, P = 0.48, RM ANOVA, data not shown). In contrast, pretreatment with 10 μM LY-303511, the inactive analog of LY-294002, did not alter the increased AP firing produced by NGF. The increase in AP firing produced by NGF in untreated neurons and in the presence of LY-303511 were not different at any of these time points (ANOVA, see Fig. 4 legend for details). For both untreated and LY-303511-pretreated neurons, exposure to NGF produced a significant depolarization of the resting membrane potential neurons (untreated control, −57.0 ± 1.5 mV vs. 10 of min NGF, −52.2. ± 2.3 mV; and LY-303511 control, −60.3 ± 2.0 mV vs. 10 of min NGF, −53.7 ± 1.6 mV, RM ANOVA), whereas for those neurons pretreated with LY-294002, NGF had no effect (control, −56.3 ± 1.8 mV vs. 10 min of NGF, −56.2 ± 1.8 mV, P = 0.57, RM ANOVA). Together, these results demonstrate that NGF via ceramide and PI3K led to the activation of PKCζ, which through presently undefined mechanisms increases the excitability of small-diameter sensory neurons.

siRNA targeted to PKMζ/PKCζ blocks NGF-induced sensitization.

Because the sequences of the pseudosubstrate binding domains are the same for the aPKCs (PKCζ, PKMζ, and PKCλ/ι; Moscat et al. 2006), pseudosubstrate peptide inhibition will not distinguish these isoforms. Therefore, to further establish whether one of the PKCζ isoforms was the key mediator of the enhancement of excitability produced by NGF, initial studies examined the expression of these different isoforms of aPKC in untreated naive sensory neurons and the potential effects of siRNA molecules targeted to PKMζ/PKCζ. Isolated sensory neurons were treated with a pool of four different siRNA molecules targeted to PKCζ as well as two additional siRNA molecules targeted to PKCζ (siRNA 407 and siRNA 638). The effects of these siRNAs on the expression of each subtype of aPKC (PKCζ, PKMζ, and PKCλ/ι) was determined by Western blotting and quantified by densitometry. Previous studies established that the full-length isoforms of PKCζ and PKCλ/ι were detected at a molecular mass of ∼75 kDa (Akimoto et al. 1994; Hernandez et al. 2003; Sacktor et al. 1993; Selbie et al. 1993), whereas the PKMζ isoform was detected at 51–55 kDa (Hernandez et al. 2003; Sacktor et al. 1993). Representative Western blots are shown in Fig. 5, A–C, left. Figure 5A, left, depicts the Western blot obtained with the antibody sc-216 (Santa Cruz), which is known to have overlapping specificity for PKMζ/PKCζ and PKCλ/ι. This blot indicates that for the naive untreated control conditions, there was moderate expression for full-length PKCζ or PKCλ/ι (75-kDa band) and more prominent expression of PKMζ that was detected as the 49-kDa band. Treatment of sensory neurons with the siRNA pool or siRNA 638, either of which targets sequences in PKMζ and PKCζ mRNA, greatly reduced the expression of PKMζ but had little effect on the full-length isoforms. Neither siRNA 407, which specifically targets PKCζ sequences, the negative control siRNA, nor Metafectene altered expression of the 75-kDa band; however, the negative control siRNA and Metafectene appeared to have a small effect on PKMζ. Figure 5A, right, summarizes the ability of siRNAs 407, 638, and the pool to alter the expression of PKCζ/PKCλ/ι or PKMζ as measured with the sc-216 antibody. The results for siRNA 407 and 638 were obtained from five different tissue harvests, whereas the results for the siRNA pool were obtained from four of those five tissue harvests. The summary results indicate that siRNA 638 and the pool significantly reduced the expression of PKMζ by an average value of 64 ± 12% (n = 5) and 78 ± 9% (n = 4), respectively, compared with naive controls (P < 0.001, ANOVA, Holm-Sidak all pairwise test, see details in Fig. 5 legend). These treatments had no significant effect on expression of the 75-kDa band representing PKCζ/PKCλ/ι (P = 0.23, ANOVA, average reductions of 22 ± 11% and 12 ± 10% for siRNA 638 and the pool, respectively). In these five experiments, treatments with Metafectene or negative control siRNA did not affect expression of either the 49- or the 75-kDa bands. To further establish the effects of these siRNAs, a specific PKMζ/PKCζ antibody was used (Hernandez et al. 2003). Figure 5B, left, shows a representative Western blot obtained with this antibody. Consistent with the results shown in Fig. 5A, siRNA 638 and the siRNA pool reduced the expression of the 49-kDa band but had no effect on the 75-kDa band. As summarized in Fig. 5B, right, the 49-kDa band was significantly reduced by 64 ± 8% (n = 5) and 65 ± 13% (n = 4) by treatment with siRNA 638 and the siRNA pool, respectively, compared with the naive controls (P < 0.001, ANOVA, Holm-Sidak all pairwise test). Treatment with the PKCζ-specific siRNA 407, negative control siRNA, or Metafectene alone had no effect on the levels of the 49-kDa band. None of the treatments had a significant effect on the expression of the 75-kDa band representing full-length PKCζ (P = 0.15, ANOVA). To assess whether these siRNA molecules targeted to PKMζ/PKCζ affected the expression of PKCλ/ι, a selective antibody to PKCλ/ι (BD Transduction) was used. Figure 5C, left, shows a representative Western blot wherein the expression of the 49- or the 75-kDa band was not altered after siRNA exposure. The summary results shown in the Fig. 5C, right, demonstrate that these different treatments had no significant effect on the expression of the 49- or the 75-kDa band (P = 0.36 and P = 0.57, respectively, ANOVA). Thus these results indicate that after exposure to the siRNA pool and siRNA 638, the expression of PKMζ was significantly and specifically reduced, whereas neither full-length PKCζ nor PKCλ/ι was altered.

Fig. 5.

Treatment with the small interfering RNA (siRNA) pool and siRNA 638 targeted to PKMζ/PKCζ reduces the expression of PKMζ but not full-length PKCζ or PKCλ/ι in sensory neurons. A, left: a representative Western blot for the detection of immunoreactive PKMζ and PKCζ/ PKCλ/ι using the antibody sc-216 in untreated control (naive), after treatment with Metafectene (Meta), in negative control siRNA (NC), and with siRNAs targeted to PKMζ/PKCζ (638 and pool) or PKCζ (407). The primary antibody was used at 1:500 and actin at 1:2,000. The film for the atypical PKC (aPKC) blot was exposed for 30 s; actin was exposed for 2 s. A, right: levels of expression for the 49-kDa (PKMζ) and the 75-kDa bands (full-length PKCζ/PKC λ/ι) after the various treatments represented by the different bars and normalized to their respective untreated naive/actin levels. Results after treatments with siRNA 407 and 638 were obtained from 5 different tissues harvests; for the siRNA pool, results were obtained from 4 of those 5. B, left: a representative Western blot for the detection of immunoreactive PKMζ and PKCζ using the PKCζ-specific antibody for the same treatment conditions described in A. The primary antibody was used at 1:5,000 and actin at 1:2,000. The film for the PKC blot was exposed for 60 s; actin was exposed for 1 s. B, right: levels of expression for the 49-kDa (PKMζ) and the 75-kDa bands (full-length PKCζ) after the various treatments. Results after treatments with siRNA 407 and 638 were obtained from 5 different tissues harvests; for the siRNA pool, results were obtained from 4 of those 5. C, left: a representative Western blot for the detection of immunoreactive PKCλ/ι using the PKCλ/ι-specific antibody for the same treatment conditions described in A. The primary antibody was used at 1:500 and actin at 1:2,000. The film for the PKCλ/ι blot was exposed for 5 s; actin was exposed for 1 s. C, right: levels of expression for the 49-kDa (PKMζ) and the 75-kDa bands (full-length PKCλ/ι) after the various treatments. Results after treatments with siRNA 407 and 638 were obtained from 5 different tissues harvests; for the siRNA pool, results were obtained from 4 of those 5. Asterisks indicate a significant reduction (P < 0.001) in the densities obtained for that treatment condition (ANOVA with a Holm-Sidak all pairwise test; see text for details).

qPCR assessment of PKCζ, PKMζ, and PKCλ/ι expression and the effects of siRNA treatment.

In addition to Western blotting, qPCR with SYBR green chemistry was used to establish the expression levels of mRNA for PKCζ, PKMζ, and PKCλ/ι in the DRG as well as the reduction in expression produced by the siRNA in isolated sensory neurons. Using our primer set described in Table 1, the expression of PKCζ, PKMζ, and PKCλ/ι relative to the reference gene Arbp was determined in the DRG. The expression of PKCζ, PKMζ, and PKCλ/ι also was determined in tissue isolated from the lung and brain to provide perspective to the levels in the DRG. These results are summarized in Table 5. In the lung, the expression of PKCλ/ι relative to Arbp was ∼2 times higher than that of PKCζ, whereas expression of PKMζ was ∼250–500 times lower than that of PKCζ or PKCλ/ι, respectively (P < 0.001, ANOVA, Holm-Sidak all pairwise test). In contrast, the levels of expression for the three aPKCs in the brain were significantly different wherein PKMζ > PKCλ/ι >> PKCζ (P < 0.001, ANOVA, Holm-Sidak all pairwise test). In the DRG, expression levels of PKMζ and PKCλ/ι were similar, with PKCζ expression being significantly lower (P < 0.001, ANOVA, Holm-Sidak all pairwise test). Using another primer set adapted to rat from mouse sequences (kindly provided by Dr. Sourav Ghosh, University of Arizona), we obtained very similar results for the expression of PKCζ, PKMζ, and PKCλ/ι in all three tissues (see Table 5). Thus these results are consistent with observations using Western blots, wherein the expression of PKMζ greatly exceeds that of full-length PKCζ in the DRG, and with previous results reported by Hernandez et al. (2003).

Table 5.

Cq values and expression levels for PKCζ, PKMζ, PKCλ/ι, and Arbp

	PKCζ	PKMζ	PKCλ/ι	Arbp
Our primers
Lung Cq	23.91 ± 0.25	31.83 ± 0.14	22.74 ± 0.20	18.50 ± 0.09
Lung expression	0.0242 ± 0.0033*	0.0001 ± 0.000004*	0.0537 ± 0.0052*
Brain Cq	29.65 ± 0.39	23.14 ± 0.45	24.02 ± 0.14	20.30 ± 0.16
Brain expression	0.0015 ± 0.0005*	0.1920 ± 0.0171*	0.0836 ± 0.0082*
DRG Cq	33.96 ± 0.62	25.19 ± 0.44	24.88 ± 0.30	20.20 ± 0.31
DRG expression	0.00011 ± 0.00005†	0.0369 ± 0.0080	0.0403 ± 0.0049
Ghosh-adapted primers
Lung Cq	23.77 ± 0.22	33.51 ± 0.70	22.60 ± 0.16
Lung expression	0.0265 ± 0.0032*	0.00004 ± 0.00001*	0.0588 ± 0.0050*
Brain Cq	29.37 ± 0.29	23.13 ± 0.42	23.64 ± 0.08
Brain expression	0.0020 ± 0.0006*	0.1900 ± 0.0147*	0.1030 ± 0.0081*
DRG Cq	33.66 ± 0.83	25.50 ± 0.39	24.94 ± 0.35
DRG expression	0.00013 ± 0.00007†	0.0306 ± 0.0075	0.0397 ± 0.0036

Quantification cycle (Cq) and expression values are means ± SE. Gene expression was determined as the no. of copies of PKCx per the no. of copies of Arbp. Results were obtained from n = 4 lung, n = 5 brain, and n = 5 dorsal root ganglion (DRG) different tissue harvests.

^*P < 0.001 (ANOVA).

^†P < 0.001, PKCζ vs. PKMζ or PKCλ/ι (ANOVA). PKMζ and PKCλ/ι were not different.

The capacity of the siRNA molecules (siRNA pool and siRNAs 407 and 638) to knock down the expression of mRNA for PKCζ, PKMζ, and PKCλ/ι was determined in isolated sensory neurons using our primer set described in Table 1. These findings are summarized in Fig. 6, where the results for siRNA 407 and 638 were obtained from four different tissue harvests; the results for the siRNA pool were obtained with three of those four tissue harvests. Figure 6A1 demonstrates that the expression of PKCλ/ι relative to Arbp (0.013 ± 0.003, Cq 23.27 ± 0.50, n = 4) was the highest of the three isoforms of aPKC in the naive untreated neurons. The expression of PKCλ/ι/Arbp for the different treatment groups was not significantly different than that for the untreated naive neurons (P = 0.07, ANOVA on ranks). In addition, the levels of the reference gene, Arbp, were not affected by any treatment compared with the naive neurons (P = 0.66, ANOVA). For example, the Cq for Arbp in the untreated neurons was 16.86 ± 0.22 (n = 4) compared with 17.04 ± 0.07 (n = 4) after treatment with siRNA 638 (data not shown). To reduce the variance in the extent of knockdown produced by the siRNA, the levels of PKCλ/ι/Arbp after treatment were normalized to their respective untreated naive values; these results are summarized in Fig. 6A2. These treatment groups were not significantly different from the untreated naive neurons (ANOVA on ranks, P = 0.10). Thus these results demonstrate that siRNA molecules targeted to PKCζ had no effect on the expression of PKCλ/ι. Figure 6B1 illustrates the expression level of PKMζ relative to Arbp (0.0030 ± 0.0010, Cq 25.70 ± 0.52, n = 4) in the naive untreated neurons, which is approximately four times less than that of PKCλ/ι/Arbp. In contrast to PKCλ/ι, treatment with the siRNA pool, but not with siRNA 407 or 638, significantly reduced the levels of PKMζ/Arbp compared with the naive condition (ANOVA on ranks, P = 0.009, Dunn's all pairwise test). After normalization to their respective untreated naive values (see Fig. 6B2), treatments with siRNA 638 and the pool, but not siRNA 407 or the negative control siRNA, significantly reduced the expression of PKMζ/Arbp by 86.3 ± 6.1% (n = 4) and 88.0 ± 6.7% (n = 3), respectively, (P = 0.011, ANOVA on Ranks, Dunn's test). The levels of Arbp were not affected by any treatment compared with the naive neurons (P = 0.19, ANOVA). The Cq for Arbp in the untreated neurons was 17.07 ± 0.04 (n = 4) compared with 17.15 ± 0.22 (n = 4) after treatment with siRNA 638 (data not shown). These results indicate that siRNA 638 and the pool effectively reduced the expression of PKMζ mRNA by about 90%. Figure 6C1 shows that the expression level of PKCζ relative to Arbp was quite low (6.82 ± 0.91 × 10⁻⁶/Arbp, n = 4), ∼2,000 times less than PKCλ/ι/Arbp. For PKCζ, the treatments had no significant effect on expression before (P = 0.13, ANOVA) or after normalization (Fig. 6C2, P = 0.14, ANOVA). The lack of effect of siRNA on full-length PKCζ is similar to that observed for the detection of full-length PKCζ with the sc-216 and PKCζ-specific antibodies. It is possible that the PKCζ-specific siRNA 407 is ineffective because the levels of mRNA for PKCζ (average Cq of 33.96) are quite low compared with those for PKMζ (average Cq of 25.19) under normal conditions. Thus, taken together, these results demonstrate that siRNA 638 and the siRNA pool, but not siRNA 407 or the negative control siRNA, significantly reduced the expression of PKMζ and did not alter the expression of either PKCζ or PKCλ/ι.

Fig. 6.

Treatment with the siRNA pool and siRNA 638 reduces the expression of mRNA PKMζ but not full-length PKCζ or PKCλ/ι in sensory neurons. A1: the expression level of mRNA for PKCλ/ι relative to the reference gene Arbp for the different treatment conditions. A2: the results obtained in A1 after normalization to the levels of PKCλ/ι/Arbp obtained for the untreated naive condition. Results after treatments with siRNA 407 and 638 were obtained from 4 different tissues harvests; for the siRNA pool, results were obtained from 3 of those 4. B1 and B2: the expression of PKMζ/Arbp and the results after normalization to the untreated naive condition, respectively, for the different treatment conditions. C1 and C2: the expression of PKCζ/Arbp and the results after normalization to the untreated naive condition, respectively, for the different treatment conditions. Asterisks represent a significant reduction in the mRNA levels obtained for that treatment condition (ANOVA with a Holm-Sidak all pairwise test; see text for details).

On average, immunoreactive PKMζ was reduced by 65–75% and mRNA for PKMζ was reduced by nearly 90% after treatment with siRNA; however, the critical question is whether the NGF-induced enhancement of excitability was affected by siRNA treatment. As summarized in Fig. 7A, in recordings from sensory neurons that exhibited uptake of the labeled siRNA pool, the capacity of NGF to increase the number of APs was blocked (n = 7, P = 0.55, RM ANOVA), whereas after exposure to labeled negative control siRNA, NGF significantly augmented the excitability by about twofold compared with the control (n = 5, P < 0.001, RM ANOVA). The increases in AP firing produced by NGF at 2, 6, and 10 min were not different. After treatment with the siRNA pool, NGF failed to depolarize the resting membrane potential (control, −57.3 ± 1.7 mV vs. NGF, −55.9 ± 1.6 mV after 6 min, data not shown). NGF had no effect on either rheobase or R_Th after siRNA pool treatment. In contrast, after treatment with negative control siRNA, NGF significantly depolarized the resting membrane potential, reduced the rheobase, and increased R_Th (data not shown); these values were similar to those obtained for NGF alone. As described for untreated sensory neurons, NGF did not alter the firing threshold for either siRNA or negative control siRNA treatments. Similar to the siRNA pool, treatment with siRNA 638 blocked the ability of NGF to increase the excitability of sensory neurons (Fig. 7B, n = 5, P = 1.0, RM ANOVA). However, after treatment with negative control siRNA, NGF significantly increased the number of APs compared with the control (n = 4, P < 0.001, RM ANOVA). As above, increases in AP firing produced by NGF at 2, 6, and 10 min were not different. In addition, the increase in AP firing produced by NGF was similar after either corresponding treatment with the negative control siRNAs (P = 0.35, ANOVA on ranks). As a control experiment for siRNA labeling, treatment with the unlabeled siRNA pool also blocked the ability of NGF to augment excitability (control, 3.0 ± 0.4 APs vs. 10 min of NGF, 4.0 ± 0.4 APs, n = 7, data not shown). Together, these results show that either inhibition or reduced expression of PKMζ prevents the increase in the number of evoked APs produced by NGF.

Fig. 7.

Treatment with the siRNA pool and siRNA 638 targeted to PKMζ blocks the increased excitability produced by NGF. Recordings were obtained from neurons that exhibited uptake of labeled siRNA. A: treatment with the siRNA pool blocked the capacity of NGF to increase the number of APs (n = 7), whereas after exposure to labeled negative control siRNA, NGF significantly augmented the excitability (n = 5). B: treatment with labeled siRNA 638 also blocked the ability of NGF to augment AP firing (n = 5), whereas the negative control siRNA did not affect the sensitization produced by NGF (n = 4). *P < 0.05 compared with the respective control values (RM ANOVA).

Internal perfusion with recombinant PKCζ enhances excitability.

Our findings demonstrate that pseudosubstrate inhibition of aPKCs and siRNA knockdown of PKMζ prevented the increased AP firing produced by NGF, suggesting that NGF leads to the activation of PKMζ. It is well established that there is a great deal of homology in the catalytic domains for the aPKCs, i.e., there is ∼85% homology between PKCζ and PKCλ/ι, which is the most homology for any two PKC isoforms (Akimoto et al. 1994; Selbie et al. 1993). With such a large extent of overlap, this raises the question as to whether internal perfusion with active recombinant PKCζ could recapitulate the sensitization produced by NGF. As shown in Fig. 8, this is the case. A representative recording of AP firing evoked by the depolarizing ramp is shown in Fig. 8A. The control recording was obtained shortly (<2 min) after the whole cell configuration was attained. After internal perfusion of recombinant PKCζ (2 ng/μl in the pipette) for 6 min, there was a large increase in the number of evoked APs that remained elevated up to 20 min. A scattergram (Fig. 8B) of the effects of internal perfusion with PKCζ on AP firing shows that some neurons responded with a modest increase, whereas others exhibited a robust increase in firing. One neuron (cell 7) after 10 min became spontaneously active. The time-dependent effects of internal perfusion of PKCζ for the remaining six neurons are summarized in Fig. 8C. After 6 min of perfusion, there was a significant threefold increase in the number of evoked APs (RM ANOVA). The increase in AP firing produced by PKCζ at all time points (2, 6, 10, and 20 min) was not significantly different from the NGF-induced increase in evoked APs observed in untreated neurons (P = 0.54, ANOVA). As shown in Table 4, 6 min of internal perfusion with PKCζ had effects on the parameters of excitability very similar to those of exposure to NGF. PKCζ significantly depolarized the resting membrane potential, lowered the rheobase, and increased R_Th (P < 0.05, ANOVA, Holm-Sidak all pairwise test for each respective parameter group). As observed above, the value of the firing threshold was not altered compared with that of NGF after 6 min of exposure for the untreated neurons or after pretreatment with either PD-98059 or U-73122 (P = 0.51, ANOVA on ranks). To assess the specificity of the sensitizing effect, another group of sensory neurons was pretreated with the myristoylated pseudosubstrate peptide inhibitor (10 μM for 30 min) before recording. After treatment with the pseudosubstrate inhibitor, internal perfusion with recombinant PKCζ failed to increase the number of evoked APs (Fig. 8D, n = 6, P = 0.15, RM ANOVA). In addition, after treatment with the inhibitor, internally perfused PKCζ had no effect on the parameters of excitability, as observed with PKCζ alone (summarized in Table 4). The blocking effects of the pseudosubstrate inhibitor on elevated excitability produced by PKCζ are similar to the inhibitory actions on the NGF-induced modifications of excitability. In further support of the high homology in the catalytic domain of aPKCs, internal perfusion with active recombinant human PKCι also sensitized AP firing in sensory neurons (Fig. 8E). A concentration of 0.42 ng/μl PKCι was added to the pipette solution; this value was chosen so that the specific activity of PKCι was equal to that of 2 ng/μl PKCζ. Internal perfusion with PKCι produced a significant increase in the number of APs compared with control values after only 2 min (control, 3.7 ± 0.3 vs. 6.7 ± 0.9 APs, n = 3, P = 0.017, RM ANOVA, Holm-Sidak test). A higher concentration of PKCι (2 ng/μl) produced a similar effect wherein after only 2 min, the number of APs was significantly higher compared with controls (control, 4.0 ± 0.6 APs vs. PKCι after 2 min, 7.0 ± 1.3 APs, n = 3, P = 0.018, RM ANOVA, Holm-Sidak test). Neither the values for the control APs (P = 0.64, t-test) nor those for the increase in APs produced by PKCι at all time points (P = 0.49, ANOVA) were different for concentrations of 0.42 vs. 2 ng/μl; therefore, these values have been combined and are summarized in Fig. 8E. PKCι produced a significant increase in AP firing at all time points compared with the control value, whereas the increases at 2, 6, 10, and 20 min were not different (P < 0.001, ANOVA, Holm-Sidak all pairwise test). In contrast, internal perfusion with 0.08 ng/μl PKCι had no effect on AP firing over time (control, 4.5 ± 0.2 APs vs. PKCι after 20 min, 4.2 ± 0.3 APs, n = 6, P = 0.85, ANOVA on ranks). As an additional control experiment, heat-inactivated recombinant PKCζ was internally perfused into sensory neurons. Under control conditions, the ramp evoked 3.2 ± 0.6 APs (n = 5); after 10 min of internal perfusion, the number of APs remained unchanged at 3.6 ± 0.7 APs (n = 5). After 10 min of internal perfusion with heat-inactivated PKCζ, these neurons then were exposed to 100 ng/ml NGF wherein after 5 min of NGF, the number of APs in four of the five neurons was significantly increased to 7.8 ± 1.1 APs (n = 4, P = 0.003, ANOVA, Holm-Sidak all pairwise test); one of these five neurons became spontaneously active. Together, these results indicate that activation of the highly homologous catalytic domains of aPKCs can result in enhanced excitability and that these effects recapitulated the sensitization produced by NGF.

Fig. 8.

Internal perfusion with recombinant PKCζ increases the excitability of sensory neurons. A: representative recordings of AP firing evoked by the depolarizing ramp over a 20-min period. The recording pipette contained recombinant PKCζ (2 ng/μl). After 6 min, there was a large increase in the number of evoked APs that remained elevated for the 20-min duration. B: a scattergram of the effects of internal perfusion with PKCζ on AP firing for 7 individual neurons. Cell 7 became spontaneously active after 10 min. C: the time-dependent effects of internal perfusion of PKCζ for the remaining 6 neurons. D: pretreatment with PS-Inhib (10 μM for 30 min) blocked the ability of recombinant PKCζ to increase the number of evoked APs (n = 6). (Descriptions for each bar apply to C and D.) E: internal perfusion with active recombinant PKCι enhanced the excitability of sensory neurons. The sensitizing effects for 0.42 and 2 ng/μl PKCι were not significantly different and have been combined (see results for details), whereas 0.08 ng/μl PKCι was without effect. *P < 0.05 compared with the control value (RM ANOVA).

Recombinant PKCζ suppresses I_K.

To explore a possible mechanism giving rise to the increased excitability, the effects of internal perfusion with recombinant PKCζ on the amplitude and activation properties of the total outward I_K were examined. Internal perfusion with recombinant PKCζ (2 ng/μl) produced a significant time-dependent suppression of I_K (Fig. 9). Figure 9A shows current traces for a representative neuron wherein the peak amplitude of I_K obtained for the step to +60 mV was reduced from a control value (left) of 6.74 to 3.40 nA after a 20-min exposure to PKCζ (middle). The PKCζ-sensitive I_K (Fig. 9A, right) was obtained by subtraction of the traces shown in the middle panel from those shown at left. The time-dependent effects of PKCζ on the current-voltage relation for I_K are summarized for seven sensory neurons in Fig. 9B. Internal perfusion with recombinant PKCζ significantly reduced the peak amplitude of I_K (at +60 mV) from a control value of 6.42 ± 0.88 to 3.84 ± 0.78 and 3.43 ± 0.76 nA (n = 7, RM ANOVA) after 10- and 20-min exposures, respectively. The current values were transformed to conductances wherein the conductance-voltage relations obtained for the different times were fitted with the Boltzmann relation and normalized to their respective values of G_max; these results are summarized in Fig. 9C. There were small but significant increases in the values of G/G_max after 10 and 20 min of internal perfusion for voltages between −10 and +10 mV and at 0 mV, respectively (RM ANOVA, see Fig. 9 legend for details) and are consistent with the small but significant leftward shift in the value of V_0.5 (control, −4.4 ± 1.7 mV vs. 10 min, −10.6 ± 3.4 mV and 20 min, −10.1 ± 2.2 mV, n = 7, RM ANOVA); the values at 2 and 6 min were not different from control. The value of k remained unaltered over the 20-min period (control, 12.8 ± 0.5 mV vs. 10 min, 12.2 ± 0.7 mV and 20 min, 13.1 ± 0.9 mV, n = 7, RM ANOVA). For these seven neurons, the PKCζ-sensitive I_K was obtained by subtraction of the current traces obtained at 20 min from their respective controls; the current-voltage relation is summarized in Fig. 9D. The PKCζ-sensitive current begins to activate around −20 mV and thus may contribute to the increased excitability (e.g., decreased rheobase and increased R_Th). Recordings obtained under control conditions over a 20-min period demonstrated that I_K was stable wherein the average peak I_K at +60 mV at time 0 was 8.74 ± 2.55 nA compared with 8.75 ± 2.57 nA (n = 4, P = 0.96 RM ANOVA), and when normalized to their respective peak values at +60 mV, the average value after 20 min was 1.09 ± 0.04 compared with their time 0 controls (n = 4). Previously, we demonstrated that treatment with C2-ceramide produced a time-dependent suppression of I_K (Zhang et al. 2002). In addition, recent evidence suggests that ceramide directly activates PKCζ (Bourbon et al. 2000; Wang et al. 2005); together, this would suggest that currents modulated by ceramide and PKCζ should be similar. To determine whether the PKCζ-sensitive I_K was similar to the C2-ceramide-sensitive I_K, we reexamined these previous results to determine the properties of the I_K sensitive to 10 μM C2-ceramide (see Fig. 5, B and C, Zhang et al. 2002). As shown in Fig. 9, E and F, both the current-voltage and the G/G_max-voltage relations for the C2-ceramide-sensitive I_K (V_0.5 = 4.5 ± 5.6 mV, k = 11.5 ± 0.9 mV, n = 4) were quite similar to those for the PKCζ-sensitive I_K (V_0.5 = 6.1 ± 6.3 mV, k = 16.3 ± 2.9 mV, n = 7). Thus these results indicate that NGF, through downstream activation of ceramide and PKCζ, leads to increased excitability, in part, via the inhibition of voltage-dependent potassium currents.

Fig. 9.

Internal perfusion with recombinant PKCζ suppresses the total outward potassium current (I_K) in sensory neurons. A: a representative recording of the total I_K obtained under control conditions (left), after internal perfusion with recombinant PKCζ for 20 min (middle), and the PKCζ-sensitive I_K obtained by subtraction of the traces in the middle from those at left. The recording pipette contained recombinant PKCζ (2 ng/μl). Traces shown are for voltage steps from −80 to +60 mV in 20-mV increments. “0” lines indicate the zero-current level. B: the average current-voltage relation for the peak value of I_K obtained at different times during internal perfusion with PKCζ (n = 7). The values of I_K after PKCζ were significantly different from control values for voltages between −10 and +60 mV and −50 and +60 mV for the 10- and 20-min perfusion times, respectively, whereas there were no differences between the control and after 2 and 6 min (RM ANOVA). C: the conductance (G/G_max)-voltage relations obtained from the 7 neurons for the control condition and at different times after internal perfusion with PKCζ. The values for each neuron were normalized to their respective fitted values of G_max for each time point. The values of G/G_max were significantly different between the control and after 10 min for voltages between −10 and +10 mV and between the control and 20 min for 0 mV (RM ANOVA). There were no significant differences between the control and after 2 and 6 min. The lines represent the fit to the data points using the Boltzmann relation. D: the current-voltage relation for the PKCζ-sensitive I_K obtained by the subtraction of the current remaining after 20 min of internal perfusion from the respective controls; the peak value at +60 mV was 2.30 ± 0.41 nA (n = 7). E and F: the I/I_max- and G/G_max-voltage relations, respectively, for the PKCζ-sensitive I_K (PKCζ-sens I_K) compared with the C2-ceramide-sensitive I_K (C2-sens I_K) obtained from our previous report (Zhang et al. 2002). The lines through the data points in F were fitted using the Boltzmann relation.

Recombinant PKCζ enhances TTX-R I_Na.

The increased excitability of small-diameter sensory neurons produced by the proinflammatory prostaglandin PGE₂ results, in part, from an augmentation of the TTX-R I_Na (England et al. 1996; Gold et al. 1996). Previously, we showed that NGF increased the amplitude of TTX-R I_Na in a time-dependent manner and that under control conditions, the values of TTX-R I_Na were stable over time (Zhang et al. 2002, 2006). Therefore, we examined whether PKCζ modulated TTX-R I_Na after 2, 6, and 10 min of internal perfusion and thus could account for the enhanced AP firing produced by recombinant PKCζ. As shown in a representative recording (Fig. 10A), internal perfusion with PKCζ increased the amplitude of TTX-R I_Na wherein after 10 min, the peak amplitude was increased from a control value of −2.25 to −3.34 nA. Figure 10B, left, summarizes the current-voltage relations for TTX-R I_Na obtained for the control condition and after 10 min of internal perfusion with PKCζ (for clarity, the values obtained at 2 and 6 min are not shown). After the 10-min exposure, PKCζ significantly increased the average peak amplitude from a control value of −2.71 ± 0.54 to −3.60 ± 0.79 nA (n = 9 neurons, RM ANOVA, see Fig. 10 legend for details). Normalization of the PKCζ-modulated currents to their respective control peak currents (Fig. 10B, right) indicated that PKCζ significantly augmented TTX-R I_Na by 1.35 ± 0.13-fold compared with the control values (0.95 ± 0.02 for the step to −10 mV, see Fig. 10 legend for details). Surprisingly, there was a small but significant time-dependent shift in E_rev for TTX-R I_Na. For the control condition, E_rev was 32.1 ± 1.9 mV (n = 9, the calculated E_rev was 27.7 mV); however, E_rev significantly shifted over time (RM ANOVA, Holm-Sidak all pairwise test) wherein after both 6- and 10-min exposures to PKCζ, E_rev values were 37.3 ± 2.2 and 39.9 ± 1.9 mV, respectively (n = 9). E_rev after 2 min was unchanged from the control value (35.9 ± 2.0 mV, n = 9, RM ANOVA). The rightward shift was only about 8 mV. A similar small shift in E_rev was observed with the enhancement of TTX-R I_Na by BDNF (Zhang et al. 2008). The origins of this shift are unclear. Recombinant PKCζ could possibly modify posttranslationally the screening of surface charge on the membrane, which alters channel permeability. Elevations in extracellular Ca²⁺ or a lowering of intracellular Ca²⁺ concentration can produce a small rightward shift in E_Na; this has been attributed to alterations in screening of surface charge. However, such a shift was accompanied by a rightward shift in the activation curve with a large decrease in the peak amplitude of I_Na (Kostyuk and Krishtal 1977; Ohmori and Yoshii 1977). This is opposite to the leftward shift in the current-voltage relation observed after internal perfusion with PKCζ. In addition, it has been reported that angiotensin II may increase the activity of the Na⁺-K⁺-ATPase via a PKCζ-dependent pathway such that this enhanced activity reduces intracellular levels of Na⁺ (Marsigliante et al. 2003). Consistent with this idea are calculations using the Nernst equation wherein lowering intracellular Na⁺ concentration from an initial value of 8.4 mM (E_rev = 32.1 mV) to 6.2 mM (E_rev = 39.9 mV) would account for this rightward shift in E_Na. Current values of TTX-R I_Na were converted to conductance and then normalized to the value of G_max obtained for each respective neuron; the G/G_max-voltage relations are summarized in Fig. 10C. This relation for each neuron was then fitted with the Boltzmann equation. After 10 min of internal perfusion with PKCζ, there was a small but significant increase in the value of G/G_max compared with the controls for only −20 and −15 mV (RM ANOVA, Holm-Sidak all pairwise test). Consistent with this observation is a small but significant leftward shift in V_0.5 (control, −18.1 ± 2.7 mV vs. PKCζ for 10 min, −23.7 ± 3.2 mV, n = 9, RM ANOVA). The values of V_0.5 obtained at 2 and 6 min were not different from the control (2 min, −21.0 ± 3.7 mV; and 6 min, −22.2 ± 3.7 mV; n = 9 RM ANOVA). The values obtained for k were not changed over time (control, 6.4 ± 1.0; 2 min, 5.5 ± 1.5; 6 min, 5.2 ± 1.6; and 10 min, 4.8 ± 1.4; n = 9, P = 0.13, RM ANOVA). Therefore, these results show that PKCζ enhanced TTX-R I_Na with a small hyperpolarizing shift in the voltage dependence for activation.

Fig. 10.

Internal perfusion with recombinant PKCζ enhances the tetrodotoxin-resistant sodium current (TTX-R I_Na) in sensory neurons. A: a representative recording of TTX-R I_Na obtained under control conditions (left) and after 10 min of internal perfusion with recombinant PKCζ (right). The recording pipette contained recombinant PKCζ (2 ng/μl). Traces shown are for voltage steps from −60 to +40 mV in 5-mV increments. “0” lines indicate the zero-current level. B, left: the average current-voltage relation for the peak value of TTX-R I_Na obtained for the control condition (n = 9) and after internal perfusion with PKCζ for 10 min (n = 9). The current values obtained at 2 and 6 min are not shown for clarity. The values of TTX-R I_Na after PKCζ were significantly different from control values for voltages between −25 and +45 mV for the 10-min perfusion time and between +35 and +45 mV for the 6-min perfusion time, whereas there was no difference between the control and the 2-min perfusion time (RM ANOVA, Holm-Sidak all pairwise test). B, right: the I/I_max-voltage relation for the control condition and after 10 min of internal perfusion with PKCζ. Peak currents were normalized to their respective values obtained under control conditions. The values of I/I_max after PKCζ were significantly different from control values for voltages between −30 and +45 mV for the 10-min time point, between −25 and −15 mV and +25 and +45 mV for the 6-min perfusion time, and at −20 and −15 mV for the 2-min perfusion time (RM ANOVA, Holm-Sidak all pairwise test). The peak values of I/I_max were 1.07 ± 0.05 and 1.20 ± 0.10 (n = 9 neurons) for the 2- and 6-min perfusion times, respectively (data not shown). C: the G/G_max-voltage relation for TTX-R I_Na for the control condition and after 10 min of internal perfusion with PKCζ from these 9 neurons. The values of G/G_max after PKCζ were significantly different from control values for voltages at −20 and −15 mV for the 10-min time point only; there were no significant differences between the values of G/G_max for the control and at the 2- and 6-min perfusion time points (RM ANOVA, Holm-Sidak all pairwise test). The lines through the data points represent the fit using the Boltzmann relation. The values of G/G_max between 30 and 40 mV have been excluded because of their proximity to the reversal potential, which gives rise to a large variance in the data and affects the quality of the fit.

DISCUSSION

PKC is a critical effector molecule in many signaling cascades and structurally consists of two important molecular domains, the NH₂-terminal regulatory domain and the COOH-terminal catalytic domain. The PKC family is made up of three subtypes: the conventional isoforms require phosphatidylserine (PS), diacylglycerol (DAG), and Ca²⁺ for activation; the novel isoforms require PS and DAG for activation and are insensitive to Ca²⁺; and the atypical isoforms require only PS for activation (reviewed by Gould and Newton 2008; Hirai and Chida 2003; Moscat and Diaz-Meco 2000). The regulatory domain contains C1/C2 second messenger/lipid binding sites and an autoinhibitory pseudosubstrate sequence that binds to and inactivates the catalytic domain. In conventional and novel PKCs, second messenger binding to C1/C2 produces a conformational change that relieves autoinhibition by the pseudosubstrate domain, unhinging it from the catalytic domain and activating PKC. Specific synthetic pseudosubstrate peptides can bind to the active site on the catalytic domain and thus act as exogenous inhibitors of kinase activity (pseudosubstrate inhibitors). In contrast, aPKCs lack C1B/C2 so that they are neither activated by second messengers nor downregulated by phorbol esters. The aPKCs rely on PS (perhaps other lipids as well) and upstream effectors such as PI3K for activation. aPKCs have a unique PB1 domain involved in protein-protein interactions that also are critical for downstream signaling.

Our understanding of the physiological actions of aPKCs are beginning to emerge. The atypical PKCζ plays an important role in cell growth and survival through its activation of the NF-κB signaling pathway (see reviews: Hirai and Chida 2003; Moscat and Diaz-Meco 2000; Moscat et al. 2006). In addition, PKCζ may have a regulatory role in the adaptive immune response. PKCλ/ι appears to have a significant functional role in metabolic regulation (Farese and Sajan 2010) as well as the proliferation of cancer cells (Win and Acevedo-Duncan 2009). Generally speaking, the role of aPKCs in neurophysiological function is poorly understood. However, PKMζ has received much attention for its critical role in the maintenance of both late-long-term potentiation (LTP) and memory retention (reviewed by Sacktor 2008, 2011). Sacktor's laboratory has shown that tetanizing stimulation of the CA1 region of the hippocampus produced a dramatic increase in the level of PKMζ that proved to be necessary and sufficient for the maintenance of LTP (Ling et al. 2002; Sacktor 2011; Sacktor et al. 1993). Additional studies showed that PKMζ was a 51- to 55-kDa protein expressed only in the brain and that an internal promoter within the PKCζ gene generates PKMζ mRNA that codes for only the catalytic domain (Hernandez et al. 2003; Sacktor 2008, 2011). Because there is no regulatory domain and no autoinhibition, PKMζ is constitutively active. However, translation of PKMζ mRNA is repressed; in this state, inactive mRNA is transported to neuronal dendrites. Upon synaptic activation, repression is released by local kinases such as PI3K (Kelly et al. 2007) and PKMζ is synthesized (essentially translation on demand). At this point, PKMζ has low activity; however, phosphoinositide-dependent kinase-1 (PDK1) phosphorylation of the activation loop (necessary for activation of all aPKCs) shifts PKMζ to high persistent activity wherein phosphorylation of specific targets is believed to maintain LTP. The key role of PKMζ is supported by in vivo observations where pseudosubstrate inhibition of PKMζ prevented the retention of cortical long-term memory and overexpression of PKMζ enhanced this retention (Shema et al. 2007, 2011). Specific targets whereby PKMζ promotes LTP maintenance are unknown, although recent work suggests that PKMζ may regulate the trafficking of AMPA receptors (Migues et al. 2010). A physiological role for PKMζ in the regulation of neuronal function in the peripheral nervous system remains to be defined.

PKMζ and persistent nociception.

Two recent studies suggested that PKMζ may play a significant role in nociceptive sensitization. After neuropathic injury (ligation of the peroneal nerve), PKMζ activation (measured as phosphorylation of Thr⁴¹⁰ in the activation loop) was increased in the anterior cingulate cortex, a key cortical area involved in chronic pain (Li et al. 2010). Also, injection of the aPKC pseudosubstrate peptide inhibitor into the anterior cingulate cortex prevented the mechanical allodynia that resulted from the injury. Interestingly, these authors reported no change in PKMζ expression in the spinal cord after injury. In another study, PKMζ appeared to maintain the persistent sensitization of mechanical responses after a priming injection of IL-6, because intrathecal injection of the pseudosubstrate inhibitor reversed this sensitization (Asiedu et al. 2011). Expression of constitutively active PKCζ in the dorsal horn of the spinal cord also enhanced mechanical sensitivity, which persisted for at least 12 days. Additional studies are necessary to establish the exact functional role of PKMζ in regulating the nociceptive sensitivity in the spinal cord. These reports suggest that PKMζ mediates enhanced nociceptive sensitivity; however, these interpretations must be tempered by the fact that the phosphospecific antibodies and the pseudosubstrate inhibitor are not selective for PKMζ but target all aPKCs. This suggests that PKCζ and/or PKCλ/ι could be equally important. However, our results for the expression of the different aPKCs as well as the siRNA-induced specific reduction obtained in isolated sensory neurons clearly define PKMζ as a key effector of sensitization. Nevertheless, these results indicate that aPKCs play a significant role in regulating the sensitivity of nociceptive neurons after injury or inflammatory conditions.

NGF, PKMζ, and sensory neuron excitability.

In this report, we show that NGF enhanced the excitability of small-diameter capsaicin-sensitive sensory neurons through a depolarization of the resting membrane potential, a decrease in the rheobase, and an increase in the resistance at threshold; however, NGF did not affect the firing threshold. In experiments to establish the signaling pathways whereby NGF augments excitability, we found that inhibition of MEK1/2 (PD-98059), PLC (U-73122 and neomycin), or conventional/novel isoforms of PKC (BIM) had no effect on the sensitization produced by NGF. In contrast, studies with a pseudosubstrate inhibitor of aPKCs and specific knockdown by siRNA indicated that NGF somehow leads to the activation/synthesis of the atypical PKC, PKMζ, which plays a critical role in enhancing neuronal excitability. Both Western blots using selective antibodies and qPCR using gene-specific primers demonstrated that neither full-length PKCζ nor PKCλ/ι were altered by siRNA treatment. Importantly, inhibitors of PI3K suppressed the capacity of NGF to increase AP firing. This PI3K-dependent inhibition is consistent with a role of PKMζ in neuronal sensitization wherein the enhanced expression of PKMζ in the hippocampus after tetanizing stimulation was blocked by inhibition of PI3K (400 nM wortmannin; Kelly et al. 2007). In addition, PI3K and ceramide are thought to be critical upstream regulators of aPKC activity, although to our knowledge, the actions of ceramide on PKMζ have not been examined. It is well established that the catalytic domains of aPKCs are highly homologous. Based on this similarity, internal perfusion with active recombinant PKCζ or PKCι augmented neuronal excitability and recapitulated the sensitization produced by NGF. Together, these findings demonstrate that the activation/synthesis of PKMζ by NGF is a critical factor in the regulation of neuronal excitability.

Our current observations demonstrate that internal perfusion with recombinant PKCζ plays a key role in regulating the activity of potassium channels wherein PKCζ led to a large suppression of I_K (∼40%) with a small but significant leftward shift in the V_0.5 of activation. Our previous work showed that NGF could suppress I_K in a similar manner. Also, 10 μM C2-ceramide inhibited I_K by ∼60% and produced a leftward shift in V_0.5 from a control value of −0.3 ± 5.2 to −11.1 ± 1.3 mV after a 20-min exposure to ceramide (Zhang et al. 2002). The idea that PKMζ modulation of I_K gives rise to enhanced excitability is consistent with the observation that overexpression of constitutively active PKCζ in the dorsal horn produced a persistent sensitization to mechanical stimulation (Asiedu et al. 2011). In a variety of cell types, agonist-mediated activation of PKCζ led to a depolarization that resulted from a decrease in potassium current; these depolarizations were blocked by the pseudosubstrate inhibitor of PKCζ (Cogolludo et al. 2003; Ramström et al. 2004). These studies also raise questions as to the possible actions of aPKCs on other types of ion channels that regulate excitability. Our earlier study showed that NGF and ceramide enhanced TTX-R I_Na in small-diameter sensory neurons (Zhang et al. 2002). In this report, we now demonstrate that recombinant PKCζ also augments TTX-R I_Na. To our knowledge, nothing is known about the capacity of PKMζ to modulate voltage-dependent ion channel activity. However, previous studies have established that both conventional and novel isoforms of PKC can modulate I_Na. For example, phorbol ester activation of PKC augmented the amplitude of TTX-R I_Na in both sensory neurons (Gold et al. 1998) and astrocytes isolated from spinal cord (Thio and Sontheimer 1993). In addition, the epinephrine-induced increase in TTX-R I_Na was partially suppressed by the conventional/novel PKC inhibitor BIM (Khasar et al. 1999b) and the more selective peptide inhibitor of PKCε, eV1-2 (Khasar et al. 1999a). Understanding the cellular mechanisms whereby PKMζ and potentially other aPKCs modulate ion channel activity and ultimately neuronal excitability are important studies for the future.

To our knowledge, there has been only one study (Martiny-Baron et al. 1993) that has examined the concentration dependence for the inhibition of the three classes of PKC by BIM. However, Toullec et al. (1991) examined the effects of BIM on the conventional and novel isoforms and reported IC₅₀ values similar to those of Martiny-Baron et al. Thus this is partly a problem in not having a greater understanding of the specific actions of BIM on the three classes of PKC. Nevertheless, there is an order of magnitude between the IC₅₀ values for BIM between the three classes: conventional isoforms at 10–20 nM, novel isoforms at 100–200 nM, and the atypical PKCζ at 5.8 μM. The exact IC₅₀ depends on the specific isoform of PKC. In support of the effectiveness of BIM, Bonnington and McNaughton (2003) demonstrated that under normal conditions, 500 nM BIM blocked the NGF-induced enhancement of capsaicin-evoked increases in intracellular Ca²⁺. McNaughton's group provided clear evidence that the novel PKC isoform PKCε is involved in the enhancement of the transient receptor potential (TRPV1) current (Cesare et al. 1999; Vellani et al. 2001), showing that the functionally effective concentration of BIM is only 2.5–5 times the IC₅₀ value for the novel isoforms. Furthermore, Cogolludo et al. (2003) found that BIM (1 μM) failed to suppress the U46619-induced (a thromboxane A₂ agonist) contraction of isolated pulmonary arteries, whereas the contraction was blocked by Gö-6983 (10 nM), which is an inhibitor of conventional PKCs, some novel isoforms, and PKCζ (see Way et al. 2000). Because contractions were insensitive to BIM but suppressed by Gö-6983, these authors suggest that U46619 may act through PKCζ. Indeed, they found that treatment with the pseudosubstrate inhibitor of PKCζ (10 μM) blocked U46619-mediated contractions (Cogolludo et al. 2003). In this report, we show that with 1 μM BIM, the sensitizing effects of NGF are not affected. This concentration is 50–100 times the IC₅₀ for the conventional isoforms and 5–10 times the IC₅₀ for the novel isoforms. If either the conventional or novel isoforms contribute to the increased excitability produced by NGF, then at 1 μM BIM, we would expect to measure some amount of diminution of the increase in AP firing. However, this is clearly not the case given that the NGF-induced increase in firing was not significantly different between the untreated controls and in the presence of BIM. In contrast, inhibition produced by the atypical pseudosubstrate inhibitor is complete, demonstrating that the atypical PKCs play a critical role in the NGF-mediated enhancement of neuronal excitability.

Our previous work demonstrated that NGF and membrane-permeant C2-ceramide (N-acetyl sphingosine) enhanced the excitability of small-diameter sensory neurons (Zhang et al. 2002). The sensitization produced by NGF was blocked by an inhibitor of neutral sphingomyelinase (SMase), suggesting that NGF led to the liberation of an endogenous ceramide(s). Other studies have indicated that short-chain ceramides, such as C2-ceramide, do a poor job of mimicking the actions of the natural longer chain ceramides, e.g., C16-ceramide (see Hashizume et al. 1998; Simon and Gear 1998; Takeda et al. 2006; Upham et al. 2003). For example, C2-ceramide inhibits aggregation of platelets, whereas C6-ceramide, C8-ceramide, and SMase augment aggregation, suggesting that these ceramides can have different actions (Hashizume et al. 1998). In sensory neurons, our results indicate that C2-ceramide does effectively recapitulate the actions of native ceramides. This is based on the following observations. NGF, the activating ligand of p75^NTR, and C2-ceramide produce very similar increases in AP firing over time, and both are associated with quite similar changes in different parameters of excitability. NGF and C2-ceramide have nearly identical effects in augmenting TTX-R I_Na and suppressing I_K in sensory neurons. Treatment with a bacterial SMase, which should liberate endogenous ceramides, increased AP firing in a manner that was very similar to that produced by NGF and exogenous C2-ceramide (Zhang et al. 2002). The increased AP firing produced by NGF and C2-ceramide was blocked by pretreatment with 20 μM dimethyl-sphingosine, an inhibitor of sphingosine kinase, suggesting that C2-ceramide was effective in stimulating downstream signaling (Zhang et al. 2006). In the present report, the sensitization produced by both NGF and C2-ceramide was blocked by the pseudosubstrate inhibitor of PKCζ, again supporting the idea that C2-ceramide is capable of activating downstream cascades that modulate AP firing. In additional support of our contention that C2-ceramide effectively activates downstream signaling, intradermal injection of C2-ceramide (10 μg) into the hind paw of a rat produced mechanical and thermal hyperalgesia (Doyle et al. 2011; Joseph and Levine, 2004), whereas dihydro-C2-ceramide, an inactive analog, had no effect (Doyle et al. 2011). In addition, injection of TNF-α produced a mechanical hyperalgesia that was suppressed by GW4869, a selective inhibitor of neutral SMase (Joseph and Levine 2004). Early studies demonstrated that C2-ceramide produced a dose-dependent activation of a ceramide-activated phosphatase that was very similar to the dose dependency produced by either C6-ceramide or native ceramides isolated from bovine brain (Dobrowsky and Hannun 1992). C2- and C6-ceramide were capable of inducing DNA synthesis in Swiss 3T3 cells in a manner that was analogous to that produced by bacterial SMase (Olivera et al. 1992). Additional studies have indicated that these short-chain ceramides can be converted into longer chain ceramides and mimic the biosynthetic actions of the longer chain ceramides (Ardail et al. 2002; Ridgway and Merriam 1995; Venkataraman and Futerman 2001). Particularly relevant to our findings are two observations. First, treatment with C2-ceramide was capable of activating PKCζ purified from rat brain (Wang et al. 1999), and C2-ceramide and SMase caused translocation of PKCζ to the particulate fraction isolated from rat astrocytes (Galve-Roperh et al. 1997). Second, application of C2-ceramide led to significant activation of PI3K in both alveolar macrophages (500 nM or 10 μM C2-ceramide, Monick et al. 2001) and rat2 fibroblasts (40 μM C2-ceramide, Hanna et al. 1999). These observations support the notion that NGF via ceramide and PI3K could lead to the activation of PKMζ and thereby enhance neuronal excitability. Together, these results indicate that C2-ceramide effectively parallels the actions of NGF in sensory neurons; however, caution must be taken, because the efficacy of these short-chain ceramides in signaling cascades appears to be dependent on cell type.

NGF signaling and sensitization.

Activation of multiple signaling cascades by either TrkA or p75^NTR could function as the intracellular pathways mediating NGF-induced peripheral sensitization. On the basis of the existing literature that has focused on small-diameter sensory neurons, it appears that NGF augments the current conducted by TRPV1 via activation of TrkA receptors, whereas NGF enhances the capacity of these neurons to fire APs by activation of p75^NTR. This idea is based on observations wherein an inhibitor of tyrosine kinase receptors, K-252a (Knüsel and Hefti 1992, but see Kase et al. 1987), blocked NGF-induced sensitization of TRPV1 (100 nM, Shu and Mendell 1999; 100 nM, Zhu and Oxford 2007), whereas 400 nM K-252a (or the structurally different inhibitor AG-879, 30 μM) had no effect on increased AP firing produced by either NGF or BDNF (Zhang et al. 2008). Furthermore, the enhanced excitability produced by NGF or BDNF was prevented by pretreatment with the p75^NTR blocking antibody (Zhang and Nicol 2004; Zhang et al. 2008).

Despite similar neuronal preparations and experimental designs, the results regarding the nature of the signaling pathways that are causal in NGF-mediated sensitization have been surprisingly inconsistent, and this remains an unanswered question (see review by Nicol and Vasko 2007). The role of PLC in NGF-induced sensitization is controversial. In human embryonic kidney (HEK) cells, a signaling complex comprising TrkA, TRPV1, and PLCγ is thought to form wherein NGF leads to the enhancement of TRPV1 activity (Chuang et al. 2001). These authors demonstrated that NGF activation of a mutant TrkA incapable of coupling to PLCγ did not augment capsaicin-evoked current. In another study, treatment with 5 μM U-73122, a PLC inhibitor, suppressed the NGF-induced sensitization of the heat response in sensory neurons (Galoyan et al. 2003). Similarly, inhibition of PLC by 10 μM neomycin reduced enhancement of the capsaicin-evoked increase in intracellular Ca²⁺ levels produced by NGF (Bonnington and McNaughton 2003). In contrast, other studies indicated that PLC inhibition by U-73122 had no affect on NGF enhancement of the capsaicin-evoked current (20 μM, Zhu and Oxford 2007; 10 μM, Zhuang et al. 2004). In many different cell systems, NGF activates Ras with consequent activation of Erk. The MEK inhibitor PD-98059 reduced the ability of NGF to augment capsaicin-evoked currents (10 μM, Zhu and Oxford 2007; 20 μM, Zhuang et al. 2004). Another MEK inhibitor, U0126 (10 μM), similarly diminished the NGF-induced enhancement of the capsaicin-evoked increase in intracellular Ca²⁺ levels (Bonnington and McNaughton 2003). As in the PLC studies, other investigations demonstrated that MEK inhibition had little effect on the NGF-induced augmentation of capsaicin-evoked currents (Chuang et al. 2001; Shu and Mendell 2001). In studies examining the role of PKA, the PKA inhibitors H89 (500 nM) or PKI_14–22 (20 nM) reduced the NGF-induced enhancement of the capsaicin-gated current recorded from adult sensory neurons (Shu and Mendell 2001). In contrast, inhibition of PKA by 200 nM KT-5720 had no effect on the capacity of NGF to augment the capsaicin-evoked increase in intracellular Ca²⁺ in neonatal sensory neurons (Bonnington and McNaughton 2003). Finally, novel findings in a heterologous expression system indicated that NGF produced a rapid insertion of additional TRPV1 channels into the plasma membrane, which could account for the increased capsaicin-evoked influx of Ca²⁺ (Zhang et al. 2005). This sensitization was blocked by PP2, an inhibitor of Src kinase. Additional experiments showed that Src kinase phosphorylation of Tyr-200 on TRPV1 was critical for membrane insertion of TRPV1. One must also consider the fact that among these signaling pathways activated by NGF, there is likely to be significant cross talk between them. Establishing the exact sequence of events has been limited by the paucity of specific inhibitors to block targeted effector molecules in these pathways.

There does, however, appear to be a point of commonality between TrkA and p75^NTR signaling in that both the NGF-mediated sensitization of TRPV1 and excitability are blocked by inhibition of PI3K (Bonnington and McNaughton 2003; Zhu and Oxford 2007; Zhuang et al. 2004; present study). NGF via TrkA activates PI3K (reviews by Huang and Reichardt 2003; Reichardt 2006). NGF via p75^NTR also activates PI3K through interaction with specific adaptor proteins (Roux et al. 2001). It is downstream of PI3K that the signaling cascades activated by NGF appear to diverge. As an example, consider the role of the PKC pathway. BIM, a PKC inhibitor that is more selective for conventional and novel isoforms, blocks sensitization of TRPV1 currents by NGF (500 nM, Bonnington and McNaughton 2003; 1 μM, Zhu and Oxford 2007), whereas in other studies BIM (500 nM, Shu and Mendell 2001) or stauorosporine (a conventional/novel PKC inhibitor, Ling et al. 2002) had no effect on NGF sensitization of TRPV1 (Chuang et al. 2001). These differences remain to be resolved. In our studies, treatment with BIM had no effect on increased AP firing produced by NGF; however, inhibition of aPKC suppressed this enhanced excitability. These findings indicate that although NGF can enhance the sensitivity of small-diameter sensory neurons, this sensitization is accomplished through activation of different intracellular signaling pathways. These different isoforms of PKC may act on different targets, i.e., the conventional/novel isoforms may target TRPV1, whereas the atypical PKMζ may target voltage-dependent channels. Recent studies have demonstrated that activation of individual isoforms of PKC can produce differential outcomes. For example, different isoforms of PKC give rise to differential phosphorylation of distinct regions of the α-subunit of Cav1.2, which can then regulate activity (Yang et al. 2005, 2009). In addition, activation of different isoforms can alter the outcomes of downstream activation patterns. In bovine aortic endothelial cells, siRNA knockdown of novel PKCε blocked the VEGF-stimulated phosphorylation of Akt, whereas knockdown of conventional PKCα enhanced VEGF-induced Akt phosphorylation (Rask-Madsen and King 2008). Thus these observations support the idea that receptor-mediated activation of particular isoforms of PKC can modulate specific downstream targets that alter cellular activities.

NGF and PKMζ activation.

Our results indicate that NGF via a PI3K- and PKMζ-dependent pathway leads to the increased excitability. On the basis of current literature, one could speculate that NGF somehow initiates the synthesis of PKMζ from repressed mRNA localized in the neuronal cell body. Consistent with this idea are recent findings showing that after a 15-min treatment with 10 ng/ml NGF, there was significant activation of eukaryotic initiation factors, i.e., the translation machinery, in isolated adult sensory neurons (Melemedjian et al. 2010). If, in fact, this is the mechanism of action, then our results would suggest the translational events can occur rapidly given that significant increases in AP firing were observed within 2 min after exposure to NGF or BDNF (Figs. 1–4 in the present study; Zhang et al. 2008). Alternatively, NGF by a presently undefined pathway leads to the activation of existing PKMζ in the neuronal cell body and does not require protein synthesis. Recent evidence suggests that tetanus-induced long-term LTP in the presence of BDNF leads to increased expression of PKMζ in the absence of protein synthesis; however, this occurred over a much longer time scale (3 h) than our present studies (Mei et al. 2011). These findings raise critical questions regarding the specific cellular mechanisms whereby NGF activates or generates PKMζ in sensory neurons and what are the potential targets of PKMζ modulation. Thus our results demonstrate that NGF increases the excitability of small-diameter capsaicin-sensitive sensory neurons through a PI3K-mediated and perhaps a ceramide-dependent activation/synthesis of the atypical PKC, PKMζ. Important future studies need to focus on whether the increased AP firing mediated by NGF requires protein synthesis and what other effectors molecules may be necessary, e.g., what role ceramide has in the activation of generation of PKMζ.

GRANTS

This investigation was conducted in a facility constructed with support from National Institutes of Health (NIH) Research Facilities Improvement Program Grant C06 RR015481-01. This work was supported by NIH Grant NS46084 and in part by funding from the Ralph W. and Grace M. Showalter Trust (to G. D. Nicol) and NIH Grants R01 MH53576 and MH 57068 (to T. C. Sacktor).

DISCLOSURES

No conflicts of interest, financial or otherwise, are declared by the author(s).

AUTHOR CONTRIBUTIONS

Author contributions: Y.Z., J.K., and K.E.H. performed experiments; Y.Z., J.K., K.E.H., and G.D.N. analyzed data; Y.Z., J.K., K.E.H., T.C.S., and G.D.N. interpreted results of experiments; Y.Z., J.K., K.E.H., T.C.S., and G.D.N. edited and revised manuscript; Y.Z., J.K., K.E.H., T.C.S., and G.D.N. approved final version of manuscript; J.K., K.E.H., and G.D.N. conception and design of research; G.D.N. prepared figures; G.D.N. drafted manuscript.