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Voltage-gated sodium channels (VGSCs) are important channels which participate in many physiological functions. Whether VGSCs can be modulated by changes in osmolality in trigeminal ganglion (TG) neurons remains unknown. In this study, by using whole-cell patch clamp techniques, we tested the effects of hypo- and hypertonicity on VGSCs in cultured TG neurons. Our data show that tetrodotoxin-resistant sodium current (TTX-R current) was inhibited in the presence of hypo- and hypertonic solutions. In hypertonic solutions both voltage–dependent activation and inactivation curves shifted to the hyperpolarizing direction, while in hypotonic solutions only inactivation curve shifted to the hyperpolarizing direction. Transient Receptor Potential Vanilloid 4 receptor (TRPV4) activator mimicked the inhibition of TTX-R current by hypotonicity and the inhibition by hypotonicity was markedly attenuated by TRPV4 receptor blocker and in TRPV4-/- mice TG neurons. We also demonstrate that inhibition of PKA selectively attenuated hypotonicity-induced inhibition, whereas antagonism of PLC and PI3K selectively attenuated hypertonicity-induced inhibition. We conclude that although hypo- and hypertonicity have similar effect on VGSCs, receptor and intracellular signaling pathways are different for hypo- versus hypertonicity-induced inhibition of TTX-R current.
Osmolality may play an important role in regulating neuron excitability in the nervous system (Schwartzkroin et al., 1998). It has been reported that hippocampal excitability can be modulated by the complex actions exerted by changes in extracellular osmolality (Baraban and Schwartzkroin, 1998). Recently, in nociceptors throughout the body there is evidence that both hypo- and hypertonic stimuli can induce nociceptor hyperexcitability, increase its activity or sensitize nociceptors, indicating that changes in osmolality could be involved in the nociception by modulating neurons excitability in the peripheral nervous system (Alessandri-Haber et al., 2003; Alessandri-Haber et al., 2004; Alessandri-Haber et al., 2005). Voltage-gated sodium channels (VGSCs), which underlie the depolarization of action potential (AP) and determine the threshold and amplitude of AP, are important ion channels regulating the biophysical properties of nociceptive neurons (Amir et al., 2006; Waxman et al., 1999). Studies have demonstrated that VGSCs participate in nociceptive signal transduction in primary sensory neurons (Chahine et al., 2005; Wood et al., 2004). In vivo, hyperosmolar solutions selectively block APs in rat sensory myelinated fibers (Matsuka and Spigelman, 2004), implying that VGSCs may be involved in anisotonicity-induced nociception. However, whether VGSCs can be modulated by anisotonicity in sensory neurons is still unknown.
Transient Receptor Potential Vanilloid 4 (TRPV4) receptor is an important osmotic cellular sensor (Liedtke and Kim, 2005; Ramsey et al., 2006) and expressed in brain as well as in dorsal root ganglion (DRG) and trigeminal ganglion (TG) (Alessandri-Haber et al., 2003; Suzuki et al., 2003). Besides hypotonic stimulus, TRPV4 can also be activated by heat, phorbol ester compounds (4α-PDD), and the metabolite of arachidonic acid (Liedtke and Kim, 2005; Voets et al., 2002; Vriens et al., 2004). Both in vivo and in vitro studies have shown that activation of TRPV4 participates in nociceptive behavior evoked by anisotonic stimuli. For example, TRPV4 has been reported to be necessary for nociceptive behavior elicited by subcutaneous injection of mild hypertonic and hypotonic saline solutions (Alessandri-Haber et al., 2005; Alessandri-Haber et al., 2004; Alessandri-Haber et al., 2003).
Based on the sensitivity to tetrodotoxin (TTX), VGSCs are divided into TTX-resistant (TTX-R) and TTX-sensitive channels (TTX-S). TTX-R channels, including NaV1.8, and NaV1.9 isoforms, are preferentially expressed in small- and medium-sized TG and DRG neurons and involved in pain transduction (Amaya et al., 2000; Kim and Chung, 1999; Novakovic et al., 1998). VGSCs are modulated by several intracellular signaling pathways (Chahine et al., 2005). In primary sensory neurons, PKA and PKC systems are two pathways which have been extensively studied. The two pathways are activated by many biological compounds, such as prostaglandin E2, bradykinin and serotonin, and then enhance the function of TTX-R channels inducing inflammatory hyperalgesia (Cardenas et al., 1997; Gold et al., 1998; Ikeda et al., 2005; Smith et al., 2000). Other pathways such as PKG (Renganathan et al., 2002) and lipid cascade (Hourez et al., 2005) have also been reported to participate in modulating the function of VGSCs. Our previous study shows that anisotonicity can sensitize capsaicin evoked current (Icaps) via different intracellular pathways (Liu et al., 2007). In the present study, we tested the effect of anisotonicity on TTX-R current in TG neurons and then explored whether TRPV4 receptor and specific intracellular pathways were involved in the effect of hypo- and hypertonicity.
Trigeminal ganglion (TG) neurons from male Sprague–Dawley rats (180–200 g) and mice (C57BL/6 wild type and TRPV4 knockout) were cultured as described previously (Liu et al., 2006). Briefly, trigeminal ganglia were dissected aseptically and collected in modified Hank's Balanced Salt solution (mHBSS). After being washed in mHBSS, the ganglia were diced into small pieces and incubated in mHBSS for 20–40 min at 37 °C with 0.1% collagenase (Type XI-S). Individual cells were dissociated by triturating them through a fire-polished glass pipette, followed by a 10 min incubation at 37 °C with 10 μg/ml DNase I (Type IV) in F-12 medium (Life Technologies, Gaithersburg, MD) and centrifuged for 5 min at 1500 rpm/min. After centrifuging three times, the cells were cultured in F-12 supplemented with 10% fetal bovine serum. The cells were plated on poly-D-lysine coated glass coverslips (15 mm diameter) and cultured for 24 h at 37 °C in a water saturated atmosphere with 5% CO2. The cell diameter (μm) was measured with a calibrated eyepiece under phase contrast illumination.
Care of animals conformed to standards established by the National Institutes of Health. All animal protocols were approved by the Duke University Institutional Animal Care and Use Committee. All efforts were made to minimize animal suffering and to reduce the number of animals used.
All experiments were carried out at room temperature (22–23 °C). Whole-cell patch clamp recordings were obtained using an Axopatch-200B patch clamp amplifier (Axon Instruments, Foster City, CA) and the output was digitized with a Digidata 1322A converter (Axon Instruments). The sampling rate was 10 kHz and filtered at 5 kHz. The capacitance and series resistance were compensated ≥ 90%. Data obtained from neurons in which uncompensated series resistance resulted in voltage-clamp errors > 5 mV were not taken in further analysis. Liquid junction potentials were compensated before patching. When the osmolality of external solutions was changed from 300mOsm to hypotonicity or to hypertonicity, measurements of the changes in liquid junction potentials were less than 2 mV and were not corrected.
The voltage-dependent activation curve (G–V curve) was measured by 20 ms depolarizing pulses from −80 to +40 mV stepping by 5 mV with interval of 2s. The voltage-dependent inactivation curve (inactivation–voltage curve) was obtained by double pulses: precondition pulses (20ms) were from −100 to +20 mV in 5 mV steps and following 0 mV test pulse (20 ms) with interval of 4s. For recording 4α-PDD-induced current, the holding potential was −60 mV. Other experiments, unless stated, the holding potential was −80 mV.
For voltage-clamp experiments, the resistance of the glass pipettes (No. 64-0817(G85150T-3), Warner Instruments Inc., Hamden, CT, USA) was 1–2 MΩ when filled with pipette solution composed of (in mM): CsF 135, NaCl 10, CaCl2 1, MgCl2 2, EGTA 10, HEPES 10, Tris-ATP 5 at pH 7.3 and osmolality 300mOsm. The external solution was composed of (in mM): NaCl 30, KCl 5, MgCl2 3, TEA-Cl 20, Choline-Cl 35, 4-AP 3, D-Mannitol 106, HEPES 10, TTX 0.0002 at pH 7.4 and osmolality 300mOsm. The use of this low Na solution was to reduce the magnitude of the rapidly activating sodium currents to overcome possible voltage errors that may arise due to poor space clamp during the voltage command. Hypo- and hypertonic external solutions were obtained by adjusting the concentration of D-Mannitol. When recording 4α-PDD-induced current the external solution was “standard external solution” composed of (in mM): NaCl 147, KCl 5, MgCl2 1, CaCl2 2, D-glucose 10, HEPES 10 at pH 7.4 and osmolality 300mOsm. The osmolality was measured using a vapor pressure osmometer (Model 3300, Advanced Instruments, Norwood, MA).
The amplitude of TTX-R current was calculated as peak current. Data were analyzed using pClamp (Axon Instruments) and SigmaPlot (SPSS Inc., Chicago, IL, USA) software. All of data were presented as mean ± S.E.M. and the significance was indicated as P<0.05 (*) and P<0.01(**) tested by paired or unpaired Student's t-tests. G–V curve and inactivation–voltage curve were fitted by Boltzmann functions, which G/Gmax= 1/(1 + exp (V0.5 − Vm)/k) or I/Imax =1/(1 + exp (V0.5 − Vm)/k), with V0.5 being membrane potential (Vm) at which 50% of activation or inactivation was observed and k being the slope of the function. The dose-response curve was fitted by Hill equation, in which Ipeak= Ipeakmax /[1+(IC50/C)n], with n as the Hill coefficient, and IC50 as the concentration producing 50% inhibition.
Cell culture materials were purchased from GIBCO (Life Technologies, Rockville, MD, USA). KT-5823, U73122 (1-[6-((17β-3-Methoxyestra-1,3,5(10)-trien-17-yl) amino) hexyl]-1H-pyrrole-2,5-dione), D(-)Mannitol and 4α-PDD (4α-phorbol-12,13-didecanoate) were obtained from CALBIOCHEM (San Diego, CA, USA) and others, unless stated, all came from Sigma Chemical Company.
4α-PDD, KT-5823, U73122, Wortmannin, pCPT-cAMP (8-(4-Chlorophenylthio)-adenosine 3′,5′-cyclic monophosphate sodium salt), pCPT-cGMP (8-(4-Chlorophenylthio)-guanosine 3′,5′-cyclic monophosphate sodium salt), BIM (Bisindolylmaleimide II), PMA (Phorbol 12-myristate 13-acetate), LY294002 (2-(4-Morpholinyl)-8-phenyl-1(4H)-benzopyran-4-one hydrochloride), H-89 (N-[2-(p-Bromocinnamylamino)ethyl]-5-isoquinolinesulfonamide dihydrochloride) and KT5720 ((9S,10S,12R)-2,3,9,10,11,12-Hexahydro-10-hydroxy-9-methyl-1-oxo-9,12-epoxy-1H-di indolo[1,2,3-fg:3′,2′,1′-kl]pyrrolo[3,4-i][1,6]benzodiazocine-10-carboxylic acid hexyl ester) were prepared as stock solutions in DMSO. The final concentration of DMSO in the bath chamber or pipette solution was <0.1%.
KT5823, H-89, U73122, Wortmannin, LY294002 were present in the pipette solution, while 4α-PDD, pCPT-cAMP, pCPT-cGMP, BIM, PMA, KT5720 were applied in the bath solution.
In this study, rat TG neurons having projected soma diameters ranging between 15 and 30 μm were used to explore the effect of anisotonic stimuli on TTX-R current.
Fig. 1A shows when the external solution was changed from isotonicity (300mOsm) to hypotonicity (260mOsm) TTX-R current was inhibited by 31.7 ± 7.2% from −137.4 ± 20.0 pA/pF to −92.9 ± 17.6 pA/pF (n=17, paired t-test, P<0.05). The inhibition by hypotonicity was largely reversible and recovered to −131.5 ± 22.3 pA/pF after hypotonicity was washed out for 3 min. We also found that G–V curve did not markedly shift before and during hypotonic treatment (paired t-test, P>0.05) (Fig. 1C).
Fig. 1D shows the effect on inactivation–voltage curve when the extracellular solution was changed from 300mOsm to 260mOsm. In the presence of hypotonicity, the inactivation–voltage curve markedly shifted to the hyperpolarizing direction (paired t-test, P<0.05).
As seen in Fig. 2A, TTX-R current was also inhibited when the external solution was changed from isotonicity (300mOsm) to hypertonicity (330mOsm). On the average, TTX-R current was inhibited by 38.8 ± 8.5% from −140.1 ± 20.7 pA/pF to −83.1 ± 10.7 pA/pF (n=16, paired t-test, P<0.01). Different from the inhibition by hypotonicity, the inhibition of TTX-R current by hypertonicity was irreversible and the amplitude was −90.4 ± 9.2 pA/pF after a 3 min washout. Also different from the inhibition by hypotonicity, both G–V curve (Fig. 2C) and inactivation–voltage curve (Fig. 2D) shifted to the hyperpolarizing direction during hypertonic treatment (paired t-test, P<0.05).
The dose–response curve of the inhibition by anisotonic stimuli was presented in Fig. 2E. Since 260mOsm and 330mOsm exhibited maximal or near maximal inhibition of TTX-R current and since these changes in tonicity were modest, these concentrations were used in all subsequent experiments.
TRPV4 receptors are osmosensitive receptors and have been shown to be activated in anisotonic media (Liedtke et al., 2000; Liedtke, 2005). To test the hypothesis that TRPV4 receptors may be involved in the modulation of TTX-R current by anisotonicity, we tested whether 4α-PDD, the agonist of TRPV4 receptor (Watanabe et al., 2003), could mimic the inhibition by hypotonicity or hypertonicity. As shown in Fig. 3A, after exposed to 4α-PDD (3 μM) for 3 min, TTX-R current was reduced by 50.3 ± 7.2% from −138.3 ± 24.7 pA/pF to −65.0 ± 9.5 pA/pF (n=11, paired t-test, P<0.01). Moreover, the response was recoverable after washout. In the presence of 3 μM 4α-PDD, there was no shift in G–V curve (paired t-test, P>0.05) (Fig. 3C). However, 4α-PDD caused a hyperpolarizing shift in the inactivation–voltage curve (paired t-test, P<0.05) (Fig. 3D). The concentration–dependent inhibition of TTX-R current by 4α-PDD is shown in Fig. 3E. TTX-R current was inhibited 6.1 ± 1.9% (from −140.1 ± 11.7 pA/pF to −131.7 ± 10.5 pA/pF, n=6, paired t-test, P>0.05) and 69.2 ± 6.8% (from −139.9± 13.3 pA/pF to −43.9 ± 7.1 pA/pF, n=6, paired t-test, P<0.01) by 0.01 and 100 μM 4α-PDD, respectively. The dose-response curve was fitted by Hill equation with IC50 being 2.4 μM.
To further determine whether TRPV4 receptor was involved in the effects of hypo and hypertonicity, ruthenium red (RR), a non-specific TRPV4 receptor antagonist (Voets et al., 2002) was used to determine how it would affect TTX-R current under isotonic (300mOsm), hypo- (260mOsm) and hypertonic conditions (330mOsm). In isotonic condition, after exposure to 10μM RR for 3 min, TTX-R current was reduced by 12.1±3.7% from −136.9±16.1 pA/pF to −120.9±15.0 pA/pF (n=8, paired t-test, P<0.05). The inhibition of TTX-R by hypotonicity was reduced from 31.7 ± 7.2% (n=17) to 9.0±2.1% (n=13) upon pre-incubation with 10μM RR (unpaired t-test, P<0.05). In contrast, the inhibition by hypertonicity was not affected in the presence of RR (330mOsm: 38.8 ± 8.5%, n=16; 330+RR: 40.3±7.1%, n=13) (unpaired t-test, P>0.05) (Fig. 4A).
Here, it was noted that after pre-application of RR for 3 min, TTX-R current was not significantly changed before and during 3 μM 4α-PDD treatment (RR: −121.1 ± 11.9 pA/pF, RR+4α-PDD: −123.7 ± 12.4 pA/pF; n=10, paired t-test, P>0.05).
We further explored the possible role of TRPV4 receptor by measuring the effect of anisotonicity on TTX-R current in TRPV4+/+ and TRPV4-/- mice TG neurons with soma diameters ranging between 10 and 25 μm.
Effects of hypotonicity (260mOsm) on TTX-R current obtained from TRPV4+/+ and TRPV4-/- mice are shown in Fig. 4B. Similar to the results found in rat TG neurons, TTX-R current was reversibly reduced by 32.1 ± 8.4% from −147.1 ± 36.3 pA/pF to −99.8 ± 14.9 pA/pF (n=13, paired t-test, P<0.01) during hypotonic treatment in TRPV4+/+ mice TG neurons. In contrast, TTX-R current was only reduced by 14.1 ± 6.7% from −144.7 ± 24.0 pA/pF to −128.3 ± 12.1 pA/pF (n=13, paired t-test, P<0.05) in TG neurons from TRPV4-/- mice when exposed to hypotonicity. There was marked difference between the inhibition of TTX-R current by hypotonicity in TRPV4+/+ and in TRPV4-/-mice (unpaired t-test, P<0.05).
Here, TTX-R current was not markedly affected by 3 μM 4α-PDD in TRPV4-/- mice TG neurons (300mOsm: −142.1 ± 13.4 pA/pF, 4α-PDD: −143.1 ± 22.7 pA/pF; n=8, paired t-test, P>0.05).
TTX-R current was inhibited irreversibly when the extracellular osmolality was changed from 300mOsm to 330mOsm. In the presence of hypertonicity, TTX-R current was reduced by 38.1 ± 9.1% from −150.2 ± 11.4 pA/pF to −92.3 ± 19.6 pA/pF (n=14, paired t-test, P<0.01) in TG neurons from TRPV4+/+ mice. After exposure to hypertonicity, TTX-R current was decreased by 41.5 ± 9.7% from −152.4 ± 16.3 pA/pF to −90.9 ± 23.6 pA/pF (n=15, paired t-test, P<0.01) in TG neurons from TRPV4-/- mice. No statistical difference was found in the reduction of TTX-R current by hypertonicity between TRPV4+/+ and TRPV4-/- mice (unpaired t-test t, P>0.05) (Fig. 4B).
Taken together, these data demonstrated that anisotonicity had similar effect on TTX-R current in TG neurons from rats and TRPV4+/+ mice. However, it is noticeable that the inhibition by hypotonicity seen in TRPV4+/+ mice was markedly attenuated in TRPV4-/- mice, suggesting the involvement of TRPV4 receptor in hypotonic-induced inhibition.
In this study, we tested several intracellular pathways under isotonic (300mOsm), hypotonic (260mOsm) and hypertonic (330mOsm) conditions to determine which, if any, of them are involved in the effect of hypo- and hypertonicity on TTX-R current.
PKA system is one of the important intracellular signal pathways modulating VGSCs. The effects of PKA are various depending on different types of cells, subtypes of VGSCs and phosphorylation sites (Chahine et al., 2005). In sensory neurons, activation of PKA by pro-inflammatory hyperalgesic agents such as serotonin and prostaglandin E2 results in an increase of TTX-R currents (Gold et al., 1996; Gold et al., 1998). Here, under isotonic condition, TTX-R current was increased from −137.4 ± 7.9 pA/pF to −156.2 ± 22.1 pA/pF (n=8, paired t-test, P<0.05) by application of 1 mM pCPT-cAMP (agonist of PKA). TTX-R current was decreased from −140.3 ± 12.8 pA/pF to −119.9 ± 9.6 pA/pF (n=17, paired t-test, P<0.05) and from −138.8 ± 9.9 pA/pF to −120.3 ± 11.8 pA/pF, (n=15, paired t-test, P<0.05) by pre-incubation of PKA antagonists, H-89 (10 μM) or KT5720 (1 μM) 10 min, respectively. Fig. 5A shows that following pre-incubation with H-89 or KT5720 for 10 min, the inhibition by hypotonicity was markedly attenuated (unpaired t-test, P<0.05), whereas the inhibition by hypertonicity was unaffected (unpaired t-test, P>0.05).
We further tested the effect of PKA antagonists on the inhibition of TTX-R current by 4α-PDD. It was found that after pre-application of H-89 or KT5720, the inhibition by 3 μM 4α-PDD was reduced to 11.7 ± 1.3% (n=11) and 13.6 ± 2.7% (n=11) respectively, which was markedly different from that in normal pipette solution (50.3 ± 7.2%) (unpaired t-test, P<0.05). These data indicated that PKA system selectively contributed to the inhibition of TTX-R current by hypotonicity.
PKC system is another important intracellular messenger pathway modulating VGSCs. In DRG neurons, activation of PKC causes a dose-dependent increase in the amplitude of TTX-R current (Baker, 2005; Gold et al., 1996;Gold et al., 1998). In the present study, under isotonic condition, TTX-R current was enhanced from −137.5 ± 11.0 pA/pF to −153.8 ± 8.8 pA/pF (n=10, paired t-test, P<0.05) in the presence of 1 μM PMA (PKC agonist) and decreased from −136.5 ± 14.3 pA/pF to −106.6 ± 8.7 pA/pF (n=11, paired t-test, P<0.05) in the presence of 1 μM BIM (PKC antagonist). As shown in Fig. 5B, TTX-R current by hypo- or hypertonicity was not altered by pre-application of BIM, implying that PKC system did not underlie the modulation of TTX-R current by anisotonicity.
Consistent with previous study (Liu et al., 2004), it was found that activation of PKG reduced the amplitude of TTX-R current. In this study, TTX-R current was reduced from −139.5 ± 14.9 pA/pF to −111.9 ± 10.6 pA/pF (n=11, paired t-test, P<0.05) when extracellular application of 1 mM pCPT-cGMP (PKG activator) 3 min. TTX-R current was increased from −137.1 ± 13.8 pA/pF to −149.8 ± 15.2 pA/pF (n=11, paired t-test, P<0.05) when neurons were pre-incubated with 10 μM KT5823 (PKG antagonist) 10 min. As shown in Fig. 5C, following pre-incubation with KT5823, the inhibition by neither hypotonicity nor hypertonicity was markedly affected (unpaired t-test, P>0.05), which eliminated the involvement of PKG in inhibition of TTX-R current by anisotonic stimuli.
It is reported that in striatal neurons inositol 1, 4, 5-triphosphate receptors antagonistically modulate VGSCs (Hourez et al., 2005). Here PI3K inhibitor (Wortmannin and LY294002) and PLC inhibitor (U73122) were used to test whether lipid cascade participated in the inhibition of TTX-R current by anisotonicity.
TTX-R current was reduced from −135.1 ± 16.6 pA/pF to −91.3 ± 14.5 pA/pF (n=15, paired t-test, P<0.05) and from −139.1 ± 15.1 pA/pF to −112.7 ± 9.5 pA/pF (n=17, paired t-test, P<0.05) by pre-application of 2 μM Wortmannin or 50 μM LY294002 for 10 min in isotonic solution, respectively. Fig. 5D shows the different effects of PI3K inhibitors on the inhibitions of TTX-R current by hypo- and hypertonicity. In the presence of Wortmannin or LY294002, the inhibition of TTX-R current by hypertonicity was 14.9 ± 3.1% and 16.6 ± 7.8% respectively, which was markedly different from that in normal pipette solution (unpaired t-test, P<0.05). In contrast, the inhibition by hypotonicity was unaffected by either Wortmannin or LY294002 (unpaired t-test, P>0.05).
Under isotonic condition, TTX-R current was not changed in the presence of 10 μM U73122 (300mOsm: −138.0 ±16.1 pA/pF, U73122: −134.1 ± 13.1 pA/pF, n=13, paired t-test, P>0.05). Although pre-application of U73122 did not affect the inhibition by hypotonicity, it did statistically attenuate the inhibition by hypertonicity (unpaired t-test, P>0.05) (Fig. 5D). Collectively, these data suggested that lipid cascade was selectively responsible for the inhibition of TTX-R current by hypertonicity.
It has been previously shown that TG and DRG neurons with small- and medium-sized somas have characteristics of nociceptors (Cardenas et al., 1995; Liu et al., 2001). That is, they have broad action potentials with humps on their repolarization phase and are activated by capsaicin (Liu et al., 2001). Therefore, in the present study, small- to medium-sized TG neurons were used to study the role of TTX-R current in the anisotonicity-induced nociception. As shown in Fig. 2E, TTX-R current was inhibited by hypo- and hypertonicity and similar dose–response curve was also found in the effect of anisotonicity on capsaicin-induced current (Liu et al., 2007). However, different from the sensitization on capsaicin-induced current by changes in osmolality, the modulation of TTX-R current was marked when the external osmolality was changed within a small range (±40mOsm). Concerning the maximal responses, TTX-R current was reduced by 32% and 39% after exposure to hypo- and hypertonicity respectively. In this study, we found different effect of hypo- and hypertonicity on the activation and inactivation kinetics of TTX-R current, that is, both G–V curve and inactivation–voltage curve shifted to the hyperpolarizing direction under hypertonic stimulus, whereas only inactivation–voltage curve shifted under hypotonic stimulus (Figs.1 and and2).2). This difference indicated that different mechanisms might underlie the modulation of TTX-R current induced by hypo- versus hypertonicity (Fig. 6).
Although both hypo- and hypertonicity decreased TTX-R current, the inhibitions by hypo- and hypertonicity were mediated by different receptors. First, the inhibition by hypotonicity was well confirmed by TRPV4 activator. In the presence of 4α-PDD, TTX-R current was reversibly inhibited and only inactivation–voltage curve shifted to the hyperpolarizing direction (Fig. 3). Second, the inhibition of TTX-R current by hypotonicity was markedly attenuated by TRPV4 antagonist ruthenium red and in TRPV4-/- mice TG neurons, whereas that by hypertonicity not (Fig. 4). These data indicate that TRPV4 receptor is selectively involved in the inhibition of TTX-R current by hypotonicity.
But it is still unknown how TRPV4 receptor contributes to hypotonicity-induced modulation of TTX-R current. In native cells, at the room temperature, hypotonicity (260mOsm)-induced current is small and only seen in a small subpopulation of cells (Alessandri-Haber et al., 2003; Liu et al., 2007). In the present study, we found that TTX-R current was markedly decreased, but no detectable hypotonic-induced current was recorded (data not shown). Therefore, it is indicated that the inhibition of TTX-R current by hypotonicity is not dependent on hypotonic-induced current via TRPV4 receptor. Recently, some TRP channels such as TRPM1, TRPP1 and TRPP2 have been reported to have biological effects independent of channel opening, which is referred as “non-conduction function” (Kaczmarek, 2006) and it was possible that TRPV4 modulated TTX-R current through this mechanism under hypotonic stimuli. TRPV4 receptor is a poly-activated receptor which is also sensitive to warm temperature besides anisotonic stimulation (Guler et al., 2002). In order to exclude the activation by heat, all the experiments were performed at room temperature (22–23 °C). At present, we did not do the experiment at higher temperature, but given the fact that TRPV4 can be activated at warm temperature, it is likely that TRPV4 might be sensitized in vivo.
Here, it is noted that hypotonicity-induced inhibition was not completely blocked in TRPV4-/- mice TG neurons, whereas the inhibition by 4α-PDD was completely blocked in TRPV4-/- mice, indicating that besides TRPV4 receptor, there might be other factors underlying hypotonicity-induced modulation of TTX-R current.
VGSCs are modulated by many intracellular signaling pathways, such as PKA, PKC, PKG system and lipid cascade (Chahine et al., 2005). In the present study, we found that TTX-R current was inhibited by antagonists of PI3K, PKA and PKC systems (Wortmannin, LY294002, H-89, KT5720 and BIM) and increased by PKG antagonists (KT5823). So, it is obvious that TTX-R current can be modulated by the above intracellular signaling pathways in TG neurons. Then we tested which of them was involved in hypo or/and hypertonic response by using the antagonists because it was supposed that if certain intracellular system was involved in anistonicity-induced modulation, antagonism of it would blocked the process (Perez et al., 2006). By comparing the effects of antagonists on anisotonicity-induced inhibition we found that after pre-incubation of PKC or PKG antagonist, the inhibition of TTX-R current by neither hypo- nor hypertonicity was markedly affected, indicating that PKC or PKG systems is not responsible for the modulation of TTX-R current under anisotonic conditions (Fig. 5, B and C).
In the present study, the inhibition of TTX-R current by hypotonicity was reversible, whereas that by hypertonicity not. One of the possible reasons is that different intracellular signaling pathways are involved in the action induced by hypo- versus hypertonicity and the effective time of different signaling pathway varies. To support this hypothesis, it is notable that PKA and lipid cascade were selective for hypo- versus hypertonicity-induced inhibition of TTX-R current. In the present study, hypotonicity-induced inhibition was markedly attenuated (about 70%) by PKA antagonists, implying that PKA system is selectively involved in hypotonic-response. But we should note that TTX-R current was inhibited 32% by hypotonicity in TRPV4+/+ mice and this inhibition was reduced to 14% in TRPV4-/- mice, demonstrating that about 56% hypotonic-inhibition is mediated via TRPV4 receptor. Therefore, it is suggested that PKA antagonists affect not only the TRPV4-dependent inhibition but also the TRPV4-independent component under hypotonic conditions. In this study, hypertonicity-induced inhibition was selectively attenuated by PI3K and PLC inhibitors, indicating that lipid cascade is selectively responsible for hypertonic-response. Collectively, it is proposed that specific and different intracellular signaling pathways are required for the inhibition of TTX-R current by hypo- versus hypertonicity. Here we are also aware that inhibition of PKA and lipid cascade could not completely block the decrease of TTX-R current by anisotonicity, implying that other mechanisms might be involved in the process.
VGSCs, which produce the inward membrane current necessary for the generation of AP, play an important role in regulating neurons excitability (Amir et al., 2006; Black et al., 2004; Fang et al., 2005; Priest et al., 2005; Waxman and Hains, 2006). Many experimental and clinical observations have revealed a link between VGSCs and sensory neuron hyperexcitability producing pain (Devor, et al., 1992; Omana-Zapata, et al., 1997; Rizzo, 1997; Wood, et al., 2004). The electrical excitability of nociceptive neurons is an important component of the pain response. In the present study we found that TTX-R current was inhibited by changes in osmolality in TG neurons, which might result in decreased excitability of nociceptors. It is known that the generation, propagation and modulation of AP in nociceptors depend on the activity of a variety of receptors (such as TRPV1, P2X, 5-HT3, and GABA) and voltage-gated ion channels (such as Na+, Ca2+ and K+), and TTX-R current is not the sole target of anisotonicity (Liu et al., 2007). Therefore anisotonicity-induced nociception is likely the integrative result due, at least in part, to the complex modulation of a large number of ion channels on the membrane. Our present framework provides the new information for better understanding the role of TTX-R channels in anisotonicity-induced nociception.
We thank Dr. Wolfgang Liedtke for TRPV4-/- mice. This work was supported by National Institute of General Medical Sciences Grant GM-63577 and by grants from Philip Morris Inc. USA and Philip Morris International and National Natural Science Foundation of China (30571537 and 30271500).
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