Journal of Neurochemistry, 2002, 82, 903–912
Peroxynitrite enhances astrocytic volume-sensitive excitatory amino acid release via a src tyrosine kinase-dependent mechanism Rene´e E. Haskew, Alexander A. Mongin and Harold K. Kimelberg Center for Neuropharmacology and Neuroscience, Albany Medical College, Albany, New York, USA
Abstract Volume-regulated anion channels (VRACs) are critically important for cell volume homeostasis, and under pathological conditions contribute to neuronal damage via excitatory amino (EAA) release. The precise mechanisms by which brain VRACs are activated and/or modulated remain elusive. In the present work we explored the possible involvement of nitric oxide (NO) and NO-related reactive species in the regulation of VRAC activity and EAA release, using primary astrocyte cultures. The NO donors sodium nitroprusside and spermine NONOate did not affect volume-activated D-[3H]aspartate release. In contrast, the peroxynitrite (ONOO–) donor 3-morpholinosydnomine hydrochloride (SIN-1) increased volumedependent EAA release by approx. 80–110% under identical conditions. Inhibition of ONOO– formation with superoxide dismutase completely abolished the effects of SIN-1. Both the volume- and SIN-1-induced EAA release were sensitive to the
VRAC blockers NPPB and ATP. Further pharmacological analysis ruled out the involvement of cGMP-dependent reactions and modification of sulfhydryl groups in the SIN-1induced modulation of EAA release. The src family tyrosine kinase inhibitor 4-amino-5-(4-chlorophenyl)-7-(t-butyl)pyrazolo [3,4-d]pyrimidine (PP2), but not its inactive analog PP3, abolished the effects of SIN-1. A broader spectrum tyrosine kinase inhibitor tyrphostin A51, also completely eliminated the SIN-1-induced EAA release. Our data suggest that ONOO– up-regulates VRAC activity via a src tyrosine kinase-dependent mechanism. This modulation may contribute to EAAmediated neuronal damage in ischemia and other pathological conditions favoring cell swelling and ONOO– production. Keywords: anion channels, cell swelling, excitotoxicity, glutamate, ischemia, nitric oxide. J. Neurochem. (2002) 82, 903–912.
Volume-regulated anion channels (VRACs) are a group of ubiquitously expressed channels found in animal cells that are responsible for generating swelling-induced whole-cell Cl– currents (ICl,vol). Several common characteristics of VRACs include Eisenman series I anion selectivity (I–> NO3–> Br–> Cl–> F–), moderate outward rectification and intermediate single channel conductance (10–100 pS; Strange et al. 1996; Nilius et al. 1997; Okada 1997). In addition to inorganic anion permeability, VRACs are also permeable towards small organic anions and uncharged molecules collectively termed Ôorganic osmolytesÕ or Ôcompatible osmolytesÕ (reviewed in Kirk 1997; Kirk and Strange 1998). One well established role for VRACs is their involvement in cell volume regulation via the release of osmolytes. In addition, these channels are also thought to contribute to such diverse processes as cell cycle progression, organic osmolyte transport, regulation of membrane potential, and endothelial mechanosensitivity (Kirk 1997; Voets et al. 1997; Kirk and Strange 1998; Lang et al. 1998; Nakao et al. 1999; Shen et al. 2000). In the hypothalamo-
neurohypophysial system, VRACs found in astrocytes mediate tonic taurine release, which in turn regulates the electrical activity of magnocellular neurons (Hussy et al. 1997; Deleuze et al. 1998). Under pathological conditions associated with cell swelling such as trauma, ischemia or spreading depression, VRACs contribute to the pathological release of excitatory amino acids (EAAs) and subsequent Received March 14, 2002; revised manuscript received May 14, 2002; accepted May 14, 2002. Address correspondence and reprint requests to H. K. Kimelberg, Center for Neuropharmacology and Neuroscience, MC-60, Albany Medical College, 47 New Scotland Avenue, Albany, New York 12208, USA. E-mail:
[email protected] Abbreviations used: db-cGMP, dibutyryl-cGMP; EGF, epidermal growth factor; NEM, N-ethylmaleimide; NPPB, 5-nitro-2-(3-phenylpropylamino)-benzoate; ODQ, 1H-[1,2,4]oxalidazolo[4,3-a]quinoxalin1-one; PP2, 4-amino-5-(4-chlorophenyl)-7-(t-butyl)pyrazolo[3,4-d]pyrimidine, specific inhibitor of src kinase family; RVD, regulatory volume decrease; SIN-1, 3-morpholinosydnomine hydrochloride; SNP, sodium nitroprusside; SOD, superoxide dismutase; SpNO, spermine NONOate; TKs, tyrosine kinases; VRACs, volume-regulated anion channels.
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neuronal damage (Kimelberg et al. 1990; Phillis et al. 1997; Kimelberg and Mongin 1998; Basarsky et al. 1999; Seki et al. 1999). Although the biophysical properties of VRACs are well studied, their molecular identity and the precise mechanisms of their activation remain controversial and uncertain. In astrocytes and other cell types, changes in intracellular ionic strength (Nilius et al. 1998; Voets et al. 1999), and several enzymes of intracellular signalling, including tyrosine kinases (Tilly et al. 1993; Crepel et al. 1998; LeppleWienhues et al. 1998; Voets et al. 1998), myosin light chain kinase (Nilius et al. 2000), protein kinase C (Robson and Hunter 1994), p21Rho small GTPase (Tilly et al. 1996; Nilius et al. 1999) and calmodulin (Mongin et al. 1999a; Li et al. 2002), have been implicated in VRAC activation. In astrocytes, VRAC activity and EAA release is potently modulated by certain neurotransmitters such as ATP (Mongin and Kimelberg 2000; Jeremic et al. 2001). Recently, Ellershaw et al. (2000) have found that in rabbit vein myocytes, nitric oxide (NO) can both activate VRACs via a cGMP-dependent mechanism and inhibit their activity via an unidentified pathway. In the CNS, NO acts as a neurotransmitter and neuromodulator (Dawson and Snyder 1994). Astrocytes, thus, are exposed to significant levels of NO during neuronal excitation, and also express high levels of soluble guanylyl cyclase (sGC), an enzyme that is activated by NO and produces cGMP (Baltrons et al. 1997; Baltrons and Garcia 1999; Teunissen et al. 2001). In addition to its direct actions, many NO effects are mediated by NO-related reactive nitrogen species. For instance, NO rapidly reacts with the superoxide anion (O2–), forming one of its most reactive products, peroxynitrite (ONOO–). This reaction is most prominent under pathological conditions and is a major cause of oxidative stress, mitochondrial dysfunction and DNA damage, as seen in ischemia and several other neurological disorders (Beckman and Koppenol 1996). While ONOO– is known to be cytotoxic, recent emerging evidence suggests that it may also modulate the activity of several intracellular signalling enzymes, including src kinases (Mallozzi et al. 1999), mitogenactivated protein kinases (Go et al. 1999; Schieke et al. 1999), and protein kinase C (Balafanova et al. 2002), some of which may be involved in the control of VRAC activity (see above). Because NO has been found to modify VRAC activity in myocytes (Ellershaw et al. 2000), in the present study we explored the possibility that astrocytic VRACs are activated or modulated by NO directly or via NO-related reactive nitrogen species such as ONOO–. To address this hypothesis we measured the effects of NO and peroxynitrite donors on the volume-dependent release of EAA in primary astrocyte cultures. In addition, we used a number of selective inhibitors to reveal the intracellular mechanisms involved in VRAC modulation by reactive nitrogen species.
Materials and methods Chemicals 3 D-[ H]aspartate (specific activity 18 Ci/mM) was obtained from Du Pont-NEN Research Products (Boston, MA, USA). Dispase (neutral protease Dispase Grade II) was purchased from Boehringer Mannheim (Indianapolis, IN, USA). All cell culture reagents were from Gibco (Grand Island, NY, USA). 3-Morpholinylsydnoneimine chloride (SIN-1) was purchased from Tocris Cookson (Ballwin, MO, USA). 4-Amino-phenylpyrazolo [3,4-d]pyrimidine (PP3) and tyrphostin A51 were from Calbiochem (La Jolla, CA, USA). 4-Amino-5-(4-chlorophenyl)-7-(t-butyl)pyrazolo[3,4-d]pyrimidine (PP2) was from BIOMOL Research Laboratories (Plymouth Meeting, PA, USA). ATP disodium salt, dibutyryl-cGMP (db-cGMP), N-ethylmaleimide (NEM), 5-nitro-2-(3-phenylpropylamino)-benzoate (NPPB), 1H-[1,2,4]oxadiazolo[4,3-a]quinoxalin-1one (ODQ), sodium nitroprusside (SNP), spermine NONOate (SpNO), superoxide dismutase (SOD), and other chemicals, unless otherwise specified, were from Sigma Chemical (St Louis, MO, USA). Cell culture Confluent primary astrocyte cultures were prepared from the cerebral cortex of newborn Sprague–Dawley rats, according to Frangakis and Kimelberg (1984), with minor modifications as described below. All animal procedures were approved by the institutional animal care committee and adhered to the National Institutes of Health guide for animal care. The cerebral cortices were separated from the meninges and basal ganglia, and tissue was dissociated using the neutral protease Dispase. Dissociated cells were seeded on poly-D-lysine coated 18 · 18-mm glass coverslips (Caroline Biological Supply Co, Burlington, NC, USA) and grown for 3–4 weeks in minimal essential medium (MEM) supplemented with 10% heat-inactivated horse serum (HIHS), 50 U/mL penicillin and 50 lg/mL streptomycin at 37C in a humidified 5% CO2/95% air atmosphere. Culture medium was replaced twice a week. After 2 weeks of cultivation, penicillin and streptomycin were removed from the culture medium. Immunocytochemistry showed ‡ 98% of the cells stained positively for the astrocytic marker glial fibrillary acid protein. Excitatory amino acid release EAA efflux measurements were performed as following. Astrocytes grown on glass coverslips were loaded overnight with D-[3H]aspartate (4 lCi/mL, final concentration 220 nM) in 2.5 mL of MEM containing 10% HIHS in a CO2 incubator set for 5% CO2/95% air at 37C. Before the start of the efflux measurements, the cells were washed of extracellular isotope and serum-containing medium in HEPES-buffered solution. The basal HEPES-buffered medium contained: 122 mM NaCl, 3.3 mM KCl, 0.4 mM MgSO4, 1.3 mM CaCl2, 1.2 mM KH2PO4, 10 mM D-glucose, 25 mM HEPES. pH was adjusted to 7.4 with NaOH (15 mM). The coverslips were inserted into a Lucite perfusion chamber which had a depression precisely cut in the bottom to accommodate the coverslip and a Teflon screw top leaving a space above the cells of around 100 lm in height. The cells were superfused at a flow rate of 1.0 mL/min in an incubator set at 37C with HEPES-buffered media. In hypo-osmotic media, NaCl concentration was reduced to 72 mM () 50 mM NaCl, a 35% decrease in medium osmolarity). The osmolarities of all buffers
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were checked using a freezing point osmometer (Advanced Instruments, Needham Heights, MA, USA). Superfusate fractions were collected at 1-min intervals. At the end of each experiment, the isotope remaining in the cells was extracted with a solution containing 1% sodium dodecyl sulfate (SDS) plus 4 mM EDTA. Four milliliters of Ecoscint scintillation cocktail (National Diagnostics, Atlanta, GA, USE) were added and each fraction was counted for [3H] in a Packard Tri-Carb 1900TR Liquid Scintillation Analyzer (Packard Instrument Co., Meriden, CT, USA). Percent fractional isotope release for each time point was calculated by dividing radioactivity released in each 1-min interval by the radioactivity left in the cells (the sum of all the radioactive counts in the remaining fractions up to the beginning of the fraction being measured, plus the radioactivity left in the cell digest) using a custom computer program. Intracellular ATP measurements ATP was assayed by the luciferin-luciferase light reaction using the standard manufacturer kit (Sigma Chemical) and a liquid scintillation counter for the light production quantification. The ATP assay mixture was diluted 200-fold with ATP dilution buffer (Sigma Chemical). Cells were treated with NO donors for 15 min and then lysed with 1 mL of 0.1% Triton X-100 plus 50 lM EGTA. The lysates were placed for 10–15 s in a boiling water bath to inactivate exo- and endo-ATPases and were stored on ice until assayed for not more than 30 min. Fifty microliters of the cell extract were added to 1 mL of ATP assay mixture and counted within 10 s of addition after gentle swirling. The ATP values were obtained from an ATP standard curve performed on the same day and normalized for the protein content in the same wells. A bicinchoninic acid assay was used to measure protein concentration (Goldschmidt and Kimelberg 1989). Statistical analyses Statistical analysis was performed in Origin 6.0 (Microcal Software, Northampton, MA, USA). Data are presented as mean ± SEM values of five to 17 experiments performed on at least two different astrocyte preparations. Effects of all agents were always compared to the controls performed on the same day and on the same culture preparation. The data were analyzed by a one-way ANOVA followed by a post-hoc Newman–Keuls test for multiple comparisons.
Fig. 1 Nitric oxide donors and dibutyryl-cGMP do not affect volumedependent excitatory amino acid release in astrocyte cultures. Effects of 100 lM sodium nitroprusside (d) (a), 10 lM (d) and 100 lM (m) spermine NONOate (b), and 100 lM (d) dibutyryl-cGMP (c) on astrocytic D-[3H]aspartate release under iso-osmotic and hypo-osmotic
Results
Effects of NO donors on basal and volume-dependent EAA release To evaluate the effects of NO donors on VRAC activity we exposed cortical astrocyte cultures to hypo-osmotic medium (35% reduction in osmolarity) to induce significant cell swelling and VRAC-mediated D-[3H]aspartate efflux. Under these conditions practically all swelling-activated EAA release is mediated by anion channels (Rutledge et al. 1998). Cells were treated with NO donors for 10 min before and during superfusion with hypo-osmotic medium. Sodium nitroprusside (SNP) 100 lM, or the more potent NO donor spermine NONOate (SpNO) 10 lM and 100 lM, did not alter swelling-induced D-[3H]aspartate efflux (Figs 1a and b). Although the NO donors tested had no overall effect on the volume-dependent D-[3H]aspartate release in our experiments, Ellershaw et al. (2000) found both NO-induced inhibition and stimulation of the VRAC-mediated Cl– currents in different cells from the same myocyte cell preparation. Therefore, it is possible that superimposition of inhibitory and stimulatory NO effects in an entire astrocyte population obscured the NO effects on individual cells. To address this possibility we attempted to isolate the cGMPdependent mechanism of NO action, which is responsible for up-regulation of VRAC activity in myocytes (Ellershaw et al. 2000), by using the cell-permeable cGMP analog dibutyryl-cGMP (db-cGMP). One hundred lM of db-cGMP had no effect on D-[3H]aspartate release under hypo-osmotic conditions (Fig. 1c). Stimulation of EAA release by the peroxynitrite donor SIN-1 In addition to the activation of sGC and subsequent cGMPdependent effects, NO also directly regulates the activity of various ion channels typically through the modification of sulfhydryl (SH) groups (Bolotina et al. 1994; Koivisto and
conditions. Cells were treated with the nitric oxide donors or db-cGMP for 10 min before and during exposure to hypo-osmotic medium. Data are presented as the mean ± SEM of 5–9 experiments performed on two to three different cell culture preparations.
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Nedergaard 1995; Shin et al. 1997; Xu et al. 1998). In many instances, this modification of SH-groups is mediated by peroxynitrite rather than NO itself (Beckman and Koppenol 1996). We therefore tested the effect of SIN-1, which simultaneously generates NO and O2– followed by immediate ONOO– formation, on the volume-dependent EAA release. 3 D-[ H]aspartate efflux by SIN-1 potentiated 80–110% at the concentration of 500 lM (Fig. 2a) but had no statistically significant effect at a 100-lM concentration (data not shown). This lower sensitivity to ONOO– is characteristic of astrocytes, as compared to other cell types, and is likely due to their effective cytoplasmic antioxidant protection (Bolanos et al. 1995). Because the effective
Fig. 2 Peroxynitrite (ONOO–) potently up-regulates astrocytic excitatory amino acid release via volume-regulated anion channels. (a) Effects of the ONOO– donor SIN-1 (500 lM, d) or a combination of SIN-1 and the O2– scavenger superoxide dismutase (SOD, 200 U/mL, m) on volume-dependent D-[3H]aspartate release. Astrocytes were treated with SIN-1 or SIN-1 plus SOD for 10 min before and during exposure to hypo-osmotic medium. Data are the means ± SEM of 7– 17 experiments performed on four different cell culture preparations. **p < 0.001, SIN-1 versus control (s); ##p < 0.01 SIN-1 versus SIN1 + SOD. (b) Effects of the anion channel blockers, 10 mM ATP and 100 lM NPPB on maximum volume-dependent D-[3H]aspartate release with or without SIN-1 treatment. Astrocytes were exposed to 500 lM SIN-1 for 10 min prior to and during hypo-osmotic medium exposure. Anion channel blockers were added to hypo-osmotic medium only. h, percentage inhibition of the control volume-dependent release; j, percentage inhibition of the volume-dependent release in SIN-1-treated cells. Data are the means ± SEM of the values for 3–5 experiments performed on two different cultures. **p < 0.01, ATP or NPPB versus control, ##p < 0.01, ATP + SIN-1 or NPPB + SIN-1 versus SIN-1 alone.
concentration of SIN-1 was higher compared to the concentrations of NO donors used, we additionally tested 500 lM SpNO. Unlike 500 lM SIN-1, SpNO at the same concentration showed a tendency to reduce volume-dependent EAA release, which was not statistically significant (approx. 40% inhibition, n ¼ 4, p ¼ 0.28, data not shown). To confirm that ONOO– was responsible for SIN-1-induced enhancement of volume-dependent EAA release, we superfused astrocyte cultures with 500 lM SIN-1 in combination with 200 U/mL superoxide dismutase (SOD), an enzyme that scavenges and metabolizes O2– and therefore prevents the formation of ONOO–. Application of SOD completely eliminated the SIN-1 effect on D-[3H]aspartate release (Fig. 2a). In order to establish that the SIN-1-induced potentiation of volume-sensitive EAA release was via VRAC, we tested a known VRAC inhibitor ATP (which blocks the channel activity at extracellular millimolar concentrations) and a broad spectrum anion channel blocker NPPB (Okada 1997). In our experiments, 10 mM ATP inhibited D-[3H]aspartate release by 79% in SIN-1-stimulated cells as compared to 80% inhibition under control hypo-osmotic conditions (Fig. 2b). 100-lM NPPB reduced the maximal D-[3H]aspartate efflux by 62% in SIN-1-treated cells as compared to 63% inhibition under control hypo-osmotic conditions (Fig. 2b). To determine whether the peroxynitrite-induced stimulation of astrocytic VRACs was due to the modification of SH-groups, we compared the effects of SIN-1 to those of the membrane-permeable SH-modifying reagent, N-ethylmaleimide (NEM). In contrast to SIN-1, 50 lM NEM potently inhibited EAA release under hypo-osmotic conditions (Fig. 3a). Similarly, when co-applied with SIN-1, NEM inhibited D-[3H]aspartate release to the same extent as under control hypo-osmotic conditions (80% inhibition in both cases, Fig. 3a). The equal sensitivity of EAA release to NEM in cells treated and untreated with SIN-1 is consistent with a similar transport mechanism. Although EAA release in NEM-treated cells recovered to near basal levels in iso-osmotic media, we always observed a secondary potent up-regulation of EAA release (Fig. 3a). This secondary up-regulation likely has a non-specific nature and several potential causes include cell lysis and detachment from substrate, or massive reversal of glutamate transporters upon energy failure (Longuemare and Swanson 1995). We did not examine this phenomenon further because the secondary EAA release was only seen in NEM-treated cells after exposure to hypo-osmotic medium and not found in cells treated with SIN-1 alone (Fig. 2a). At high concentrations, ONOO– may activate sGC (Beckman and Koppenol 1996). Therefore to completely rule out the involvement of sGC in mediating the effect of ONOO– on VRAC activity, we used the sGC inhibitor ODQ in conjunction with SIN-1. ODQ did not significantly affect the ONOO–-induced VRAC potentiation (Fig. 3b). These results are consistent with our db-cGMP data and confirm
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Fig. 4 Effect of nitric oxide donors, SIN-1 and NEM on intracellular ATP levels. Astrocyte cultures were treated with 100 lM SNP, 100 lM spermine NONOate, 500 lM SIN-1, 500 lM SIN-1 plus 200 U/mL SOD, or 50 lM NEM for 20 min. After extracellular medium aspiration, cells were lysed, and intracellular ATP levels were measured using a luciferin-luciferase reaction as described in Materials and methods. Each data set represents the mean ± SEM of 5–9 measurements of intracellular ATP levels normalized to control values measured in the same cell culture.
Fig. 3 The modification of SH-groups and activation of soluble guanylyl cyclase are not responsible for the SIN-1 effects on excitatory amino acid release. (a) Effect of the cell-permeable SH-group modifying reagent N-ethylmaleimide (NEM) on the volume-dependent 3 D-[ H]aspartate release with or without SIN-1 treatment. Cells were treated with 50 lM NEM (d), 500 lM SIN-1 (n), or a combination of SIN-1 + NEM (m) for 10 min before and during exposure to hypoosmotic medium. Each data set represents the means ± SEM of 4–7 experiments performed on three culture preparations. **p < 0.01, NEM versus control (s), ##p < 0.01, SIN-1 versus SIN-1 + NEM. (b) Effect of the specific soluble guanylyl cyclase inhibitor, ODQ (10 lM), on the volume-dependent D-[3H]aspartate release in cells treated with 500 lM SIN-1. Each data set represents the mean ± SEM of 6–10 experiments performed on two to three different culture preparations. **p < 0.01 SIN-1 versus control, #p < 0.05 SIN-1 + ODQ versus control.
that cGMP is not involved in enhancing VRAC activity in cultured astrocytes. Intracellular ATP levels The NO donors and other agents used in this study are capable of inhibiting energetic metabolism and mitochondrial function (Brown 2001). A decrease in intracellular ATP levels, on one hand, is known to suppress VRAC activity and VRAC-mediated EAA release (Jackson et al. 1994; Rutledge et al. 1999), and on the other hand, diminished intracellular ATP levels may induce EAA release unrelated to the VRAC activity (Longuemare and Swanson 1995). We therefore measured intracellular ATP levels in astrocytic cultures treated for 20 min (time corresponding to 10-min preincubation plus 10-min exposure to hypo-osmotic medium during efflux experiments) with the NO donors, SIN-1 and NEM. Within this time frame 100 lM SpNO, 500 lM SIN-1, or SIN-1 plus 200 U/mL SOD did not alter intracellular ATP
levels as compared to the ATP content measured from untreated astrocytes from the same multiwell plate (Fig. 4). In contrast, cells treated with 100 lM SNP or 50 lM NEM showed a 30–35% reduction of ATP levels (Fig. 4). Effect of tyrosine kinase inhibitors on the modulation of EAA release by SIN-1 Recent findings suggest that ONOO– can activate nonreceptor tyrosine kinases of the src family via the nitration of tyrosine residues (Mallozzi et al. 1999, 2001) and p56LCK kinase, a member of the src family, has been implicated in VRAC activation (Lepple-Wienhues et al. 1998). Therefore, we used the src kinase family inhibitor PP2 to test for the potential involvement of src in the effects of SIN-1. Pretreatment with 10 lM PP2 inhibited the peak SIN-1-induced increment in astrocytic D-[3H]aspartate release by 70% (Fig. 5a). In contrast, control EAA release under hypoosmotic conditions was not significantly affected by PP2 treatment alone (Fig. 5a; 19% inhibition, p ¼ 0.364). To verify the specificity of the PP2 effects we used PP3, an inactive structural analog of PP2. In contrast to PP2, 10 lM PP3 had no significant effect on the SIN-1-induced increment in EAA release (n ¼ 5, p ¼ 0.442, data not shown). Src participates in signalling transduction pathways involving other tyrosine kinases (TKs) that have been implicated in the activation or positive modulation of VRAC function in astrocytes (Crepel et al. 1998; Deleuze et al. 2000). Therefore, we also tested the effect of a broader spectrum TK inhibitor, tyrphostin A51, on both volumedependent EAA release and its modulation by SIN-1. Twenty lM of tyrphostin A51 completely eliminated SIN1-induced enhancement of the volume-dependent EAA release but, when used alone, was ineffective under control hypo-osmotic conditions (Fig. 5b).
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smooth muscle cells, the intercellular signalling molecule NO positively modulates VRAC activity via a cGMPdependent mechanism (Ellershaw et al. 2000). The latter finding, and the fact that astrocytes have high levels of sGC, prompted us to explore whether NO may also act as an activator or modulator of VRACs in astroglial cells. Our data show that peroxynitrite, rather than NO itself, potently modulates astrocytic VRACs via a mechanism involving non-receptor tyrosine kinases of the src family.
Fig. 5 SIN-1 enhances excitatory amino acid release via a tyrosine kinase-dependent mechanism. (a) Effect of the specific src kinase family inhibitor, 10 lM PP2 either alone (n) or in combination with 500 lM SIN-1 (m) on the volume-dependent D-[3H]aspartate release. Cells were pre-treated with PP2 for 20 min before exposure to hypoosmotic medium. Data are the means ± SEM of 6–12 experiments performed on three to four different culture preparations. **p < 0.01 SIN-1 (d) versus control (s); #p < 0.05, SIN-1 versus SIN-1 plus PP2 (m). (b) Effect of the tyrosine kinase inhibitor 20 lM tyrphostin A51 either alone (n) or in combination with 500 lM SIN-1 (m) on the volume-dependent D-[3H]aspartate release. Data are the means ± SEM of 6–10 experiments performed on two different culture preparations. **p < 0.01 SIN-1 (d) versus control (s); ##p < 0.01, SIN-1 versus SIN-1 plus tyrphostin A51 (m).
Discussion
There is a general agreement that volume-regulated anion channels (along with volume-regulated cation channels) are critical for regulatory volume decrease (RVD) in virtually all animal cells when cells are subjected to significant osmotic gradients in vitro (Strange et al. 1996; Lang et al. 1998). Because in healthy brain tissue significant degrees of cell swelling are unlikely to exist, it is not clear whether VRACs are functional in the CNS under normal physiological conditions. This uncertainty was first resolved in the hypothalamo-neurohypophysial system. In the hypothalamus, astrocytic VRACs are tonically active in non-swollen cells and mediate taurine release, which via glycine receptor activation regulates the secretion of vasopressin by magnocellular neurons (Hussy et al. 1997, 2000, 2001; Deleuze et al. 1998). Recent in vitro data suggest that certain neurotransmitters, such as ATP, can potently modulate astrocytic VRACs in non-swollen or moderately swollen cells (Mongin and Kimelberg 2000; Jeremic et al. 2001). In
Lack of cGMP-dependent modulation of VRACs in astrocytic cells In various cell types, VRAC activity is modulated by several serine/threonine protein kinases including protein kinases A and C and the myosin light chain kinase (Jackson and Strange 1993; Robson and Hunter 1994; Oz and Sorota 1995; Nilius et al. 2000). The cGMP-dependent modulation found in acutely isolated vein myocytes (Ellershaw et al. 2000) does not exist in cultured astrocytes because we found no effect of the NO donors or membrane-permeable cGMP analog on VRAC activity measured as volume-dependent EAA release. The stimulation of VRACs by the ONOO– donor SIN-1 was as well insensitive to the sGC inhibitor ODQ. The lack of cGMP involvement is rather unexpected as astrocytes in culture and in situ express both the NO-activated enzyme, soluble guanylyl cyclase, and cGMP-dependent protein kinases (Baltrons et al. 1997; de Vente et al. 2001; Takuma et al. 2001; Teunissen et al. 2001). Therefore VRACs responsible for EAA release in astrocytes or VRAC-associated regulatory proteins either have no sites for cGMP-dependent phosphorylation or are inaccessible to protein kinase G. Peroxynitrite enhances VRAC activity via a tyrosine kinase-dependent mechanism In our experiments, the NO donors did not affect VRAC activity at concentrations sufficient to up-regulate cGMP production and other NO-related reactions. In contrast, the peroxynitrite donor SIN-1 strongly enhanced astrocytic volume-dependent EAA release. The O2– scavenger SOD abolished this effect verifying the involvement of ONOO–. In order to confirm that the SIN-1-induced EAA release was mediated by VRACs, we tested two commonly used VRAC inhibitors, NPPB and ATP. While NPPB is a broad-spectrum anion channel inhibitor, extracellular ATP at high concentrations selectively inhibits VRACs (Strange et al. 1996; Okada 1997). Both of the inhibitors equally attenuated volume-dependent EAA release with or without SIN-1 treatment, supporting VRAC involvement. We next considered several potential intracellular mechanisms responsible for the peroxynitrite-induced VRAC activation. Among them, a cGMP-dependent mechanism seemed least likely and was ruled out by the ODQ experiments (see previous section on cGMP). Several types
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of ion channels are directly activated by NO and related species via modification of protein sulfhydryl groups (Bolotina et al. 1994; Koivisto and Nedergaard 1995; Shin et al. 1997; Xu et al. 1998). However, VRACs as well as voltage-dependent Cl– channels are reportedly inhibited by SH-group oxidation/modification (Gonzalez et al. 1995; Nagel and Katz 1997). Consistent with the literature data and in contrast to SIN-1 effects, we found potent inhibition of the volume-dependent EAA release by the membranepermeable SH-group-modifying reagent NEM. NEM also partially decreased ATP levels and intracellular ATP binding is required for VRAC activation (Jackson et al. 1994; Rutledge et al. 1999). However, the inhibitory effect on EAA release was unlikely due to this mechanism, because SNP caused comparable reduction in ATP levels but no inhibition of EAA release. Therefore, oxidation/modification of SH-groups in VRAC or anchored regulatory proteins, inhibits VRAC activity and cannot be responsible for SIN-1 effects. Peroxynitrite oxidizes and nitrates many intracellular proteins. It has been found to activate non-receptor tyrosine kinases of the src family through the displacement of the Y527 phosphotyrosine in the SH2 domain (Mallozzi et al. 1999, 2001), as well as potentiate several members of the MAPK signalling cascade (Go et al. 1999; Schieke et al. 1999). Tyrosine kinase signalling cascades have long been considered a critical element of VRAC activation (Tilly et al. 1993; Crepel et al. 1998; Voets et al. 1998) or modulation (Deleuze et al. 2000). The src family member p56lck is both necessary and sufficient for VRAC activation in lymphocytes (Lepple-Wienhues et al. 1998). Therefore, ONOO–-induced src activation may be an alternative mechanism for VRAC activation. In our experiments both the specific inhibitor of the src kinase family, PP2, and a broader spectrum TK inhibitor tyrphostin A51, potently suppressed the ONOO– effect on astrocytic VRACs but when used alone these inhibitors were ineffective in reducing swelling-activated EAA release. This is consistent with previous findings that TKs are not involved in the activation of astrocytic amino acid-permeable VRACs but instead contribute to VRAC modulation (Mongin et al. 1999b; Deleuze et al. 2000). In contrast, TK and MAPK activation are required for swellinginduced Cl– currents in cultured astrocytes (Crepel et al. 1998). The difference between volume-dependent Cl– and amino acid release warrants further exploration and may be due to multiple permeability pathways that are differentially involved in these processes (Junankar and Kirk 2000). Two outstanding questions remain: (i) whether VRAC is an immediate target for TKs and (ii) whether src regulates VRAC activity directly or via other members of TK cascades. Due to the unidentified molecular nature of VRACs, the first question is currently impossible to address. Tyrphostin A51 is a highly potent inhibitor of receptor TKs and is especially potent at the epidermal growth factor
(EGF)-receptor kinase (IC50 ¼ 800 nM; Levitzki 1990). However, to our knowledge, this compound does not act on any members of the src family. Therefore, the ONOO–induced EAA release in astrocytes likely involves additional tyrphostin-sensitive enzyme(s) of TK signalling cascades. The likelihood that the EGF receptor is involved is small because we previously found no effect of EGF on volumedependent EAA release (Mongin et al. 1999b). Possible significance of the ONOO–-induced VRAC activation Recently, astrocytic EAA release has been proposed as an important feedback signal in neuron-astrocyte communication (Carmignoto 2000; Haydon 2001). We initiated this study based on the hypothesis that NO released during neuronal excitation may trigger EAA release both in swollen and nonswollen astrocyte cultures. However, in our experiments, neither NO nor the related molecule, ONOO–, modified EAA release under basal iso-osmotic conditions. It therefore seems unlikely that NO contributes to the regulation of EAA release in astrocytes under physiological conditions. In contrast, ONOO– should be an important regulator of EAA release in various neural pathologies associated with cell swelling and ONOO– production such as trauma, ischemia and others (Kimelberg and Mongin 1998; Kimelberg 2000). Pathological glutamate release is potently inhibited by VRAC blockers in animal models of ischemia and during the onset of spreading depression in slices (Phillis et al. 1997; Basarsky et al. 1999; Seki et al. 1999). ONOO– formation in ischemic stroke has been quantitatively linked to neural damage (Eliasson et al. 1999; Osuka et al. 2001). As seen from our data, ONOO– increases VRAC activity via a src-dependent mechanism by 80–110% and, therefore, may be an important factor of elevated EAAs and excitotoxic brain damage. This action may supplement other toxic effects of ONOO–, such as the oxidative damage of cellular components and disruption of energetic metabolism (Beckman and Koppenol 1996; Brown 2001). Because VRAC activity strictly depends on intracellular ATP levels (Jackson et al. 1994; Rutledge et al. 1999), anion channel activation and modulation in the ischemic infarction core is less likely. In contrast, in the penumbra intracellular ATP content remains as high as 70% of normal tissue levels (Lipton 1999) and ONOO– formation predominantly occurs during reperfusion in this peri-infarct region (Fukuyama et al. 1998). Consistent with our suggestions, Phillis et al. (1996) have found that the inhibition of tyrosine phosphorylation with genistein significantly depresses EAA release in the ischemic rat cortex. Furthermore, src gene knockout mice show a 50% reduction in infarction volume in a permanent focal ischemia model when compared to their wild-type counterparts (Paul et al. 2001). In the same study the src inhibitor PP1 significantly reduced the infarction in wild-type animals. These data support the pathological significance of
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src activation and may in part be explained by a reduction of ischemic EAA release, as implied by our in vitro study. In summary, our data suggest that in cultured astrocytes, peroxynitrite, but not nitric oxide, potently up-regulates VRAC activity and VRAC-associated excitatory amino acid release via one of the members of the src kinase family. Because ONOO–-induced up-regulation is only seen in substantially swollen cells, this mechanism is not likely to be of physiological significance. However, during pathological states associated with cell swelling, ONOO–-relevant VRAC activity may contribute to high extracellular EAA levels and subsequent excitotoxic neuronal damage. Acknowledgements The authors thank Ms C. J. Charniga for expert cell culture preparation and Drs J. A. Melendez and D. Jourd’heuil for their critical reading and many helpful suggestions on the manuscript. This study was supported by National Institute for Neurological Disorders and Stroke (NINDS) grant 35205 to HKK and by the Albany Medical College Graduate Studies Program.
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