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NEUROSCIENCE |
Departments of
1 Pharmacology and Toxicology
2 Anaesthesia, Indiana University School of Medicine, Indianapolis, IN 46202, USA
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Abstract |
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(Received 30 April 2006;
accepted after revision 30 May 2006;
first published online 1 June 2006)
Corresponding author G. D. Nicol: Department of Pharmacology and Toxicology, 635 Barnhill Drive, Indiana University School of Medicine, Indianapolis, IN 46202, USA. Email: gnicol{at}iupui.edu
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Introduction |
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It is well established that a family of receptors, such as the p75 neurotrophin receptor (p75NTR) and the p55 TNF-
receptor, are coupled to the activation of a sphingomyelinase that cleaves ceramide from membrane sphingomyelin (Dobrowsky et al. 1994, 1995; Dobrowsky & Carter, 1998; Brann et al. 1999). Nerve growth factor (NGF), which can activate p75NTR and the tyrosine kinase receptor TrkA when injected into the paw of a rat, produces hyperalgesia to thermal and mechanical stimulation (Lewin et al. 1993). Similar sensitizing actions of NGF are observed in an isolated skin–nerve-type preparation wherein NGF increases the firing frequency of isolated saphenous nerve in response to thermal stimulation (Rueff & Mendell, 1996). The effect of NGF is directly on the neuron because NGF augments the capsaicin-evoked current in small-diameter sensory neurons (Shu & Mendell, 1999, 2001). Acute exposure to NGF also enhances AP firing evoked by a ramp of depolarizing current in adult rat sensory neurons maintained in culture. This effect of NGF appears to result from activation of the sphingomyelin signalling cascade via p75NTR to liberate ceramide (Zhang et al. 2002; Zhang & Nicol, 2004). These findings raise the question as to whether this increased excitability is a direct result of the action of ceramide or secondary to the metabolism of ceramide to Sph and/or S1P. In this report, we show that internally perfused Sph or S1P augments the number of APs evoked by a ramp of current in small-diameter capsaicin-sensitive sensory neurons. Inhibition of Sph kinase blocks the capacity of NGF, ceramide and Sph to augment AP firing; however, the sensitization produced by internal S1P is not affected. These results indicate that S1P is the active second messenger by which the liberation of ceramide augments the firing of sensory neurons.
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Methods |
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Primary cultures of dissociated adult rat dorsal root ganglion (DRG) neurons were prepared as previously described (Lindsay, 1988) with slight modification (Jiang et al. 2003). Briefly, male Sprague-Dawley rats (100–150 g) were killed by placing them in a chamber that was then filled with CO2. DRGs 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–1 of papain II, and incubated for 10 min at 37°C. The tube was centrifuged for 30 s at low speed (approximately 1850 g) and the pellet was resuspended in Puck's solution containing collagenase (1 mg ml–1, type 1A) and dispase II (2.5 mg ml–1). After 10 min incubation at 37°C, the tube was centrifuged for 30 s before the enzyme-containing supernatant was removed. The pellet was resuspended in F-12 medium supplemented with 250 ng ml–1 7S nerve growth factor (Harlan Bioproducts, Indianapolis, IN, USA), 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 nerve growth factor at 37°C and 3% CO2 and used within 6–24 h for electrophysiological recordings. All procedures have been 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. 2002). Briefly, a coverslip with the sensory neurons was placed in a recording chamber where the neurons were bathed in normal Ringer solution containing (mM): 140 NaCl, 5 KCl, 2 CaCl2, 1 MgCl2, 10 Hepes and 10 glucose, pH was adjusted to 7.4 with NaOH. Patch pipettes were pulled from borosilicate glass tubing and fire-polished. Whole-cell voltages or currents were recorded with an Axopatch 200 patch-clamp amplifier (Axon Instruments, Union City, CA, USA). The data were acquired and analysed using pCLAMP 6.04 or pCLAMP 8.2 (Axon Instruments). The whole-cell recording configuration was established in normal Ringer solution. Both capacitance and series resistance compensation (typically 80%) were used. The mean series resistance before compensation was 2.1 ± 0.04 M
(n
= 106). Leak subtraction was not used for the measurement of the K+ current (IK) so that any effects of these agents on the holding current could be determined. To assess excitability in the current-clamp experiments, neurons were held at their resting potentials (range –50 to –65 mV) and a depolarizing ramp of current (1 s in duration) was applied. The amplitude of the ramp was adjusted to produce two to four action potentials (APs) under control conditions then the same ramp was used throughout the recording period for each individual neuron.
To isolate IK, neurons were superfused with a Ringer solution wherein NaCl was substituted with equimolar N-methyl-glucamine chloride (NMG-Cl, 140 mM); pH was adjusted to 7.4 with KOH. Recording pipettes typically had resistances of 2–4 M when filled with the following solution (mM): 140 KCl, 5 MgCl2, 4 ATP, 0.3 GTP, 2.5 CaCl2, 5 EGTA (calculated free Ca2+ concentration of 
100 nM; MaxChelator) and 10 Hepes; pH was adjusted to 7.3 with KOH. This pipette solution was used in the current-clamp recordings as well. The membrane was held at –60 mV; this value was chosen so that current measurements could be ascertained at a voltage that reflected the normal resting potential in these sensory neurons. Activation of IK was determined by voltage steps of 350 ms, which were applied at 5 s intervals in +10 mV increments to +60 mV. After obtaining the control response, the bath solution was changed to the appropriate Ringer solution, and cells were superfused continuously for the appropriate times. In a separate series of time control experiments, the peak amplitudes for IK did not vary significantly over a 20 min time period indicating that there was little run-down of this current over this time. At the end of each recording, the neuron was exposed to 100 nM capsaicin. This neurotoxin was used to distinguish capsaicin-sensitive sensory neurons, as these neurons are believed to transmit nociceptive information (Holzer, 1991). The results reported below were obtained from capsaicin-sensitive neurons only. All experiments were performed at room temperature (22°C).
Data analysis
Data are presented as the means ± S.E.M. The excitability parameters described in Tables 1 and 2 were determined, in part, by differentiating the voltage trace (dV/dt) in the current-clamp recordings (sampling frequency of 250 Hz). The voltage 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 over 100 ms that began with the onset of the current ramp (134–234 ms). The rheobase was measured as the amount of ramp current at the firing threshold. The membrane resistance was calculated as the difference between the firing threshold and the resting membrane potential divided by the rheobase current. The voltage dependence for activation IK was fitted with the Boltzmann equation where G/Gmax = 1/[1 + exp(V0.5 – Vm)/k], where G is the conductance (G = I/(Vm – EK)), Gmax is the maximal conductance obtained from the Boltzmann fit, V0.5 is the voltage for half-maximal activation, Vm 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 ± S.E.M. Statistical differences between the control recordings and those obtained under various treatment conditions were determined by using a t test, a paired t test, an analysis of variance (ANOVA), or a repeated measures ANOVA (RMANOVA). When a significant difference was obtained with an ANOVA, post hoc analyses were performed using a Tukey test. Values of P < 0.05 were judged to be statistically significant.
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C2-ceramide (N-acetyl sphingosine, hereafter referred to as ceramide), Sph and S1P were obtained from Avanti Polar Lipids, Inc. (Alabaster, AL, USA). All other chemicals were obtained from Sigma Chemical Corp. (St Louis, MO, USA). Tissue culture supplies were purchased from Invitrogen (Carlsbad, CA, USA). Ceramide, Sph and capsaicin were dissolved in 1-methyl-2-pyrrolidinone to obtain concentrated stock solutions. S1P was dissolved according to instructions provided by the supplier (http://www.avantilipids.com/SyntheticSphingosine-1-phosphate.asp). These stock solutions were then diluted with Ringer solution or the pipette solution to yield the appropriate concentration. We have demonstrated previously that the vehicle 1-methyl-2-pyrrolidinone has no effect on AP firing or the activation of IK (Zhang et al. 2002).
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Results |
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Our previous work demonstrated that ceramide, a product of sphingomyelin hydrolysis, enhanced the excitability of small-diameter capsaicin-sensitive sensory neurons (Zhang et al. 2002). However, it is well established that ceramide can be further metabolized to Sph through the action of ceramidase and, in turn, Sph can be phosphorylated by Sph kinase to yield S1P (Spiegel et al. 1996; Pyne & Pyne, 2000; Hannun et al. 2001). This raises the question whether the ceramide-induced augmentation of excitability was due directly to ceramide or to its metabolic products. To address this question, the effect of Sph on excitability was investigated by using a ramp of depolarizing current to evoke APs in the absence or presence of Sph. An extracellular application of 10 µM Sph to isolated sensory neurons had no effect on the number of APs, even after a 20 min exposure (summarized in Fig 1B, 3.9 ± 0.1 control versus 4.1 ± 0.2 after a 20 min treatment, n = 7). Because Sph is a polar lipid and a sufficient quantity may not cross the neuronal membrane over this 20 min time period, in another series of experiments Sph (1 µM) was perfused internally into sensory neurons by diffusion from the recording pipette. As shown in a recording from a representative neuron (see Fig. 1A), the number of APs evoked by the ramp increased from a control value of 3 APs to 10 APs after 20 min of internal perfusion. The results obtained from seven sensory neurons under these conditions are summarized in Fig. 1C and show that the number of APs increased in a time-dependent manner from an average control value of 3.1 ± 0.3 to a value 10.9 ± 1.6 (RMANOVA) after a 20 min exposure to internal Sph. The effects of internal Sph on the parameters of excitability for seven neurons are summarized in Table 1. The increased excitability resulting from the 20 min exposure to Sph was not accompanied by depolarization of the membrane or a change in the firing threshold of the AP. However, in these same neurons, the latency for the generation of the first AP was significantly decreased by 2.2 ± 0.8 fold (n = 7, RM ANOVA), the rheobase was reduced by 2.8 ± 0.5 fold, and the membrane resistance was increased by 2.5 ± 0.5 fold. These results demonstrate that elevations in internal Sph significantly enhance the excitability of these sensory neurons without altering the resting Vm or the firing threshold. These observations are similar to those reported previously by our laboratory for the effects of NGF and ceramide in augmenting the excitability in capsaicin-sensitive sensory neurons (Zhang et al. 2002).
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Although all three sphingolipids, ceramide, Sph and S1P, augment the excitability of sensory neurons, ceramide and Sph can be converted to S1P by a series of enzymatic reactions. This raises the question as to which sphingolipids are effective second messengers that enhance the excitability of sensory neurons. To examine this idea, dimethylsphingosine (DMS), a specific competitive inhibitor of Sph kinase, was used to block the conversion of ceramide and Sph to S1P (Olivera & Spiegel, 1993; Yatomi et al. 1996; Edsall et al. 1998). Figure 3A demonstrates that treatment with 20 µM DMS alone for 20 min had no significant impact on the excitability of sensory neurons (n = 7). In a separate series of experiments, 20 min pretreatment with 20 µM DMS blocked the capacity of internally perfused Sph (1 µM, Fig. 3B) to augment the number of APs evoked by the depolarizing ramp. Pretreatment with DMS also blocked the ability of ceramide to increase AP firing in five sensory neurons (data not shown). In contrast to the above, pretreatment with DMS did not alter the capacity of internal S1P to augment the number of evoked APs in six neurons and increased the number from a control value of 4.2 ± 0.5 to 10.3 ± 1.3 after 20 min (see Fig. 3C). In the presence of DMS, S1P decreased the rheobase and the latency, increased the membrane resistance, and did not alter the firing threshold. This excitatory effect of S1P in the presence of DMS is similar to the results described above for S1P alone. These results support the idea that the sensitizing action of Sph is secondary to its metabolism to S1P.
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It is well established that a reduction in K+ currents enhances the excitability of sensory neurons (Weinreich & Wonderlin, 1987; Gold et al. 1996; Nicol et al. 1997; Cordoba-Rodriguez et al. 1999; Zhang et al. 2002). Therefore, we explored the idea that the sensitizing action of Sph or S1P observed in the current-clamp experiments results from the inhibition of an outward K+ current (IK). Internal perfusion with Sph (1 µM) produced a time-dependent decrease in IK in capsaicin-sensitive adult sensory neurons (see Fig. 5A). As shown for a representative neuron, under control conditions (left panel) the peak IK obtained for the step to +60 mV was 8.43 nA. After internal perfusion with 1 µM Sph for 20 min, IK was reduced to 6.02 nA (middle panel) and corresponds to 30% inhibition. The Sph-sensitive IK (right panel) was obtained by subtraction of the traces in the middle panel from those in the left panel. This Sph-sensitive current exhibits little time-dependent decrease in amplitude during the voltage step suggesting that it may be a delayed rectifier type of IK. The Sph-induced inhibition of IK obtained from five neurons is summarized in Fig. 5B. The left panel of Fig. 5B shows the current–voltage relations for the time-dependent suppression of IK where Sph significantly reduced IK from an average control value of 8.90 ± 1.88 nA to a value of 6.47 ± 1.35 nA (step to +60 mV) 20 min after attaining the whole-cell configuration. The inhibition of IK measured at +60 mV after 20 min corresponded to a reduction by 26 ± 5% (range 17–42% inhibition). In another series of experiments, internal perfusion with 50 µM Sph reduced IK by 57 ± 2% (n
= 4, data not shown) after a 20 min exposure. The effects of Sph on the conductance–voltage relation are summarized in the right panel of Fig. 5B, and the Boltzmann fitting parameters are summarized in Table 3. The values of G/Gmax were reduced in a time-dependent manner; however, there was no significant change in the average values for V0.5 and k. In a separate series of studies, the inhibition of IK produced by internally perfused Sph (1 µM) was blocked by pretreatment with 20 µM DMS (5.83 ± 0.53 for the control versus 5.97 ± 0.60 nA after 20 min perfusion with Sph, n
= 5). In addition, IK was recorded from four small-diameter sensory neurons before and after exposure to 20 µM DMS. Under control conditions, the peak IK measured at +60 mV was 7.62 ± 1.18 nA; these values were not significantly different (P
= 0.07, t test) from the control IK for those experiments examining the internal perfusion of Sph (as shown in Fig. 5B). After 5 and 10 min exposures to 20 µM DMS, the peak IK values at +60 mV were 7.71 ± 1.11 and 7.58 ± 0.95 nA, respectively. These results suggest that DMS by itself does not inhibit IK, and that the low values of IK for that particular series of experiments wherein DMS blocked the Sph-induced decrease in IK were within biological variation. Figure 5C illustrates the current–voltage relations for the Sph-sensitive IK (left panel) and their normalization to the maximal peak current obtained at +60 mV (I/Imax, right panel). It appears that this current begins to activate at about –10 mV because the values of the Sph-sensitive IK were significantly different at this voltage compared with those obtained at –60 mV (ANOVA). These results indicate that internally perfused Sph can suppress IK without significantly shifting V0.5 and that this inhibition depends on the conversion of Sph to S1P.
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30% inhibition). The S1P-sensitive IK obtained by subtraction has a peak amplitude of 4.85 nA and is illustrated in the right panel. Figure 6B (left panel) summarizes the current–voltage relations for the suppression of IK by S1P observed in seven neurons at the different time points wherein IK was reduced significantly from a control value of 9.57 ± 1.81 nA to a value of 6.28 ± 1.41 nA after a 20 min exposure to S1P (RM ANOVA). This corresponds to a reduction of 32 ± 8% (range 0–56%) of the peak IK obtained at +60 mV and is similar to the 26 ± 5% inhibition by Sph. One of the seven neurons exposed to S1P exhibited a reversal of inhibition after 20 min, even though IK was suppressed by 16% at 10 min. If this neuron is excluded, then the suppression produced by S1P ranged from 15 to 56%. Figure 6B (right panel) demonstrates the conductance–voltage relation wherein the Boltzmann fitting parameters are summarized in Table 3. As for Sph, S1P reduced the value of G/Gmax in a time-dependent manner but did not alter V0.5 or k. For the control recordings, the values of V0.5 for Sph- and S1P-treated neurons were not different (P
= 0.13, t test). The current–voltage relation for the S1P-sensitive IK is shown in Fig. 6C (left panel). This current begins to exhibit significant activation at about –10 mV (compared to –60 mV, ANOVA) and is the same voltage as determined for activation of the Sph-sensitive IK. The normalization of the S1P-sensitive IK to their respective maximal peak currents obtained at +60 mV (I/Imax) is shown in the right panel of Fig. 6C. Here, the S1P-sensitive current exhibits a significant increase at about –30 mV, perhaps resulting from the reduced variability upon normalization. The Sph-sensitive current–voltage relation is overlaid on that for the S1P-sensitive current and demonstrates that these two currents are essentially the same (P > 0.05, t test for each respective voltage). Taken together with the findings for DMS, these many points of similarity between the Sph- and S1P-sensitive currents would strongly indicate that S1P is the active agent that leads to the suppression of IK.
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Discussion |
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The idea that S1P acts as an intracellular signalling agent is supported by the observations that treatment with the Sph kinase inhibitor DMS blocked the ability of NGF, ceramide and internal Sph to augment excitability. In contrast, exposure to DMS did not affect the sensitization produced by internally applied S1P. We assume that DMS is preventing the conversion of exogenous or endogenous ceramide and Sph to S1P since previous work demonstrated that exposing cells to this sphingosine kinase inhibitor decreases the levels of cellular S1P and increases Sph and/or ceramide levels (Meyer zu Heringdorf et al. 1998; Hobson et al. 2001). Furthermore, our previous work suggests that a component of NGF-induced excitability in sensory neurons is mediated by the p75 neurotrophin receptor (Zhang & Nicol, 2004) and activation of this receptor initiates the sphingolipid signalling cascade (Dobrowsky et al. 1994; Brann et al. 1999). Thus, our current results, when taken with previous findings, indicate that NGF-induced modulation of excitability in sensory neurons involves activation of a signalling pathway that ultimately liberates S1P as an intracellular second messenger.
Previous studies by various investigators using different endpoints suggest that other signalling pathways also contribute to the NGF-induced peripheral sensitization. At the receptor level, much indirect evidence supports the notion that activation of the TrkA receptor mediates the sensitizing actions of NGF. Co-expression of TrkA and TRPV1 receptors in oocytes and HEK cells results in an increase in capsaicin-sensitivity after NGF treatment (Chuang et al. 2001), suggesting that TrkA activation is critical for NGF-induced increases in currents conducted by TRPV1. In addition, systemic injection of NGF results in hyperalgesia in transgenic mice with a presumed p75 receptor knock-out (Lee et al. 1992), suggesting that the nociceptive actions of NGF are mediated by TrkA (Bergmann et al. 1998). However, recent work with this transgenic strain shows that the animals express a protein isoform of the p75 receptor (von Schack et al. 2001; Paul et al. 2004), thus complicating the interpretation of these behavioural results. Activation of the TrkA receptor will increase the activity of a number of downstream signalling pathways (Kaplan & Miller, 2000) and inhibiting these cascades attenuates NGF-induced sensitization of sensory neurons. Although this implies that TrkA activation mediates NGF-induced sensitization, results vary depending on the preparation used and the endpoints examined. For example, NGF-induced augmentation of capsaicin-evoked currents in expression systems are attenuated by blocking phospholipase C (PLC) activity (Chuang et al. 2001), whereas NGF actions on capsaicin currents in isolated sensory neurons are reduced by inhibition of protein kinase A (PKA) and protein kinase C (PKC) (Shu & Mendell, 2001). Augmentation of the capsaicin-induced increase in intracellular Ca2+ concentration by NGF are reversed by inhibition of phosphatidylinositol 3 kinase and PKC, but not by inhibitors of MAP kinase or PKA (Bonnington & McNaughton, 2003). Thus, controversy remains as to which signalling cascades mediate the acute sensitizing actions of NGF. It is possible that different NGF signalling pathways mediate various effectors that can contribute to the sensitization of sensory neurons. Furthermore, both TrkA and p75 receptors may be necessary for the NGF-induced peripheral sensitization since there is precedent in the literature for an interaction between these TrkA and p75 receptors (Bilderback et al. 2001; Lad et al. 2003). Additional studies are warranted to determine which of these possibilities can account for the variable results seen by different investigators when examining the acute effects of NGF on sensory neurons.
Although our results suggest that S1P acts as an intracellular second messenger, it also is possible that in cells, S1P can be dephosphorylated to Sph, which in turn can be catabolized to ceramide, and ceramide can be phosphorylated by ceramide kinase to produce ceramide 1-phosphate (C1P), which recent evidence indicates can act as an intracellular second messenger much like S1P (Colombaioni & Garcia-Gil, 2004; Lamour & Chalfant, 2005). In non-neuronal cells, exposure to S1P increases COX2 expression (Pettus et al. 2003, 2005) and increases the accumulation of cAMP (Damirin et al. 2005), whereas C1P can increase phospholipase A2 activity that results in increased levels of arachidonic acid (Pettus et al. 2005). These results suggest that activation of the sphingolipid cascade could increase the production of prostaglandins in cells. This could, in part, account for the sensitizing actions of ceramide and S1P since the prostaglandins E2 and I2 increase intracellular cAMP and consequently sensitize sensory neurons (Hingtgen et al. 1995; Nicol et al. 1997). In addition, S1P can be released from a variety of cell types (see below) and acts as a first messenger by binding to G-protein-coupled receptors (S1P receptors; also known as EDG receptors; Taha et al. 2004; Rosen & Goetzl, 2005). The binding of S1P to its receptors activates a number of signal transduction cascades (Spiegel & Milstien, 2002, 2003; Rosen & Goetzl, 2005) and results in an increase in excitability of sensory neurons (Zhang et al. 2006). S1P increases the intracellular Ca2+ concentration independent of the production of inositol trisphoshate (Ghosh et al. 1990, 1994), and potentially could increase excitability as well. However, in sensory neurons this seems unlikely for two reasons. First, treatment with external S1P did not elevate intracellular Ca2+ levels in capsaicin-sensitive sensory neurons (Y. H. Zhang & G. D. Nicol, unpublished observations). Second, acute exposure to NGF did not increase intracellular Ca2+ levels (Bonnington & McNaughton, 2003). At present, other potential effectors or second messengers activated by S1P in neurons are unknown (see review by Colombaioni & Garcia-Gil, 2004) and this remains an area of future investigation.
In contrast to our observations in sensory neurons demonstrating that external Sph is ineffective at altering the generation of APs, previous studies have shown that external Sph is capable of modulating membrane currents. When applied externally to rat ventricular myocytes, Sph reduced both INa and ICa (McDonough et al. 1994; Yasui & Palade, 1996). Similar results were reported for Sph-induced inhibition of ICa in GH4C1 cells (Titievsky et al. 1998). Whether the action of Sph on these cells was secondary to the generation of S1P by activation of Sph kinase was not determined. In this regard, exposing human umbilical vein endothelial cells to Sph leads to the formation of S1P in the external medium (Ancellin et al. 2002). Furthermore, Sph kinase activity is detected in conditioned media from these cells, demonstrating that Sph kinase can be secreted or released from cells. This could be a mechanism leading to increased levels of S1P in the plasma with consequent activation of the G-protein-coupled S1P receptors. Our negative results with externally applied Sph suggest that this lipid is not active on sensory neurons and that it is not metabolized to S1P under our experimental conditions.
In conclusion, our current observations, when taken together with our previous findings (Zhang et al. 2002), suggest that activation of sphingolipid metabolism in sensory neurons is an important signalling pathway in modulating excitability, and that NGF-mediated sensitization involves activation of this pathway. Ceramide can be metabolized to S1P, which then modulates the activity of different ion channels by an unknown mechanism to enhance AP firing in nociceptive sensory neurons. Thus, the sphingomyelin signalling cascade is likely to play an important role in controlling the sensitivity of sensory neurons after exposure to inflammatory or neuropathic conditions.
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